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Standard Methods for the Examination of Water and Wastewater
Part 9000
9010
MICROBIOLOGICAL EXAMINATION
INTRODUCTION*#(1)
The following sections describe procedures for making microbiological examinations of
water samples to determine sanitary quality. The methods are intended to indicate the degree of
contamination with wastes. They are the best techniques currently available; however, their
limitations must be understood thoroughly.
Tests for detection and enumeration of indicator organisms, rather than of pathogens, are
used. The coliform group of bacteria, as herein defined, is the principal indicator of suitability of
a water for domestic, industrial, or other uses. The cultural reactions and characteristics of this
group of bacteria have been studied extensively.
Experience has established the significance of coliform group density as a criterion of the
degree of pollution and thus of sanitary quality. The significance of the tests and the
interpretation of results are well authenticated and have been used as a basis for standards of
bacteriological quality of water supplies.
The membrane filter technique, which involves a direct plating for detection and estimation
of coliform densities, is as effective as the multiple-tube fermentation test for detecting bacteria
of the coliform group. Modification of procedural details, particularly of the culture medium, has
made the results comparable with those given by the multiple-tube fermentation procedure.
Although there are limitations in the application of the membrane filter technique, it is
equivalent when used with strict adherence to these limitations and to the specified technical
details. Thus, two standard methods are presented for the detection and enumeration of bacteria
of the coliform group.
It is customary to report results of the coliform test by the multiple-tube fermentation
procedure as a Most Probable Number (MPN) index. This is an index of the number of coliform
bacteria that, more probably than any other number, would give the results shown by the
laboratory examination; it is not an actual enumeration. By contrast, direct plating methods such
as the membrane filter procedure permit a direct count of coliform colonies. In both procedures
coliform density is reported conventionally as the MPN or membrane filter count per 100 mL.
Use of either procedure permits appraising the sanitary quality of water and the effectiveness of
treatment processes. Because it is not necessary to provide a quantitative assessment of coliform
bacteria for all samples, a qualitative, presence-absence test is included.
Fecal streptococci and enterococci also are indicators of fecal pollution and methods for their
detection and enumeration are given. A multiple-tube dilution and a membrane filter procedure
are included.
Methods for the differentiation of the coliform group are included. Such differentiation
© Copyright 1999 by American Public Health Association, American Water Works Association, Water Environment Federation
generally is considered of limited value in assessing drinking water quality because the presence
of any coliform bacteria renders the water potentially unsatisfactory and unsafe. Speciation may
provide information on colonization of a distribution system and further confirm the validity of
coliform results.
Coliform group bacteria present in the gut and feces of warm-blooded animals generally
include organisms capable of producing gas from lactose in a suitable culture medium at 44.5 ±
0.2°C. Inasmuch as coliform organisms from other sources often cannot produce gas under these
conditions, this criterion is used to define the fecal component of the coliform group. Both the
multiple-tube dilution technique and the membrane filter procedure have been modified to
incorporate incubation in confirmatory tests at 44.5°C to provide estimates of the density of fecal
organisms, as defined. Procedures for fecal coliforms and Escherichia coli include a 24-h
multiple-tube test using A-1 medium, a 7-h rapid method, and chromogenic substrate coliform
tests. This differentiation yields valuable information concerning the possible source of pollution
in water, and especially its remoteness, because the nonfecal members of the coliform group may
be expected to survive longer than the fecal members in the unfavorable environment provided
by the water.
The heterotrophic plate count may be determined by pour plate, spread plate, or membrane
filter method. It provides an approximate enumeration of total numbers of viable bacteria that
may yield useful information about water quality and may provide supporting data on the
significance of coliform test results. The heterotrophic plate count is useful in judging the
efficiency of various treatment processes and may have significant application as an in-plant
control test. It also is valuable for checking quality of finished water in a distribution system as
an indicator of microbial regrowth and sediment buildup in slow-flow sections and dead ends.
Experience in the shipment of un-iced samples by mail indicates that noticeable changes may
occur in type or numbers of bacteria during such shipment for even limited periods of time.
Therefore, refrigeration during transportation is recommended to minimize changes, particularly
when ambient air temperature exceeds 13°C.
Procedures for the isolation of certain pathogenic bacteria and protozoa are presented. These
procedures are tedious and complicated and are not recommended for routine use. Likewise,
tentative procedures for enteric viruses are included but their routine use is not advocated.
Examination of routine bacteriological samples cannot be regarded as providing complete
information concerning water quality. Always consider bacteriological results in the light of
information available concerning the sanitary conditions surrounding the sample source. For a
water supply, precise evaluation of quality can be made only when the results of laboratory
examinations are interpreted in the light of sanitary survey data. Consider inadequate the results
of the examination of a single sample from a given source. When possible, base evaluation of
water quality on the examination of a series of samples collected over a known and protracted
period of time.
Pollution problems of tidal estuaries and other bodies of saline water have focused attention
on necessary modification of existing bacteriological techniques so that they may be used
effectively. In the following sections, applications of specific techniques to saline water are not
discussed because the methods used for fresh waters generally can be used satisfactorily with
saline waters.
Methods for examination of the waters of swimming pools and other bathing places are
included. The standard procedures for the plate count, fecal coliforms, and fecal streptococci are
identical with those used for other waters. Procedures for Staphylococcus and Pseudomonas
aeruginosa, organisms commonly associated with the upper respiratory tract or the skin, are
included.
Procedures for aquatic fungi and actinomycetes are included.
Sections on rapid methods for coliform testing and on the recovery of stressed organisms are
included. Because of increased interest and concern with analytical quality control, this section
continues to be expanded.
The bacteriological methods in Part 9000, developed primarily to permit prompt and rapid
examination of water samples, have been considered frequently to apply only to routine
examinations. However, these same methods are basic to, and equally valuable in, research
investigations in sanitary bacteriology and water treatment. Similarly, all techniques should be
the subject of investigations to establish their specificity, improve their procedural details, and
expand their application to the measurement of the sanitary quality of water supplies or polluted
waters.
9020
QUALITY ASSURANCE/QUALITY CONTROL*#(2)
9020 A.
Introduction
1. General Considerations
The growing emphasis on microorganisms in water quality standards and enforcement
activities and their continuing role in research, process control, and compliance monitoring
require the establishment and effective operation of a quality assurance (QA) program to
substantiate the validity of analytical data.
A laboratory quality assurance program is the integration of intralaboratory and
interlaboratory quality control (QC), standardization, and management practices into a formal,
documented program with clearly defined responsibilities and duties to ensure that the data are
of the type, quality, and quantity required.
The program must be practical and require only a reasonable amount of time or it will be
bypassed. Generally, about 15% of overall laboratory time should be spent on different aspects
of a quality assurance program. However, more time may be needed for more important
analytical data, e.g., data for enforcement actions. When properly administered, a balanced,
conscientiously applied QA program will optimize data quality without adversely affecting
laboratory productivity.
Because microbiological analyses measure constantly changing living organisms, they are
inherently variable. Some quality control tools used by chemists, such as reference standards,
instrument calibration, and quality control charts, may not be available to the microbiologist.
Because QA programs vary among laboratories as a result of differences in organizational
mission, responsibilities, and objectives; laboratory size, capabilities, and facilities; and staff
skills and training, this provides only general guidance. Each laboratory should determine the
appropriate QA level for its purpose.
2. Guidelines for a Quality Assurance Program
Develop a QA program to meet the laboratory’s specific needs and the planned use of the
data. Emphasis on the use of data is particularly important where significant and costly decisions
depend on analytical results. An effective QA program will confirm the quality of results and
increase confidence in the data.
a. Management responsibilities: Management must recognize the need for quality assurance,
commit monetary and personnel resources, assume a leadership role, and involve staff in
development and operation of the QA program. Management should meet with the laboratory
supervisor and staff to develop and maintain a comprehensive program and establish specific
responsibility for management, supervisors, and analysts.
b. Quality assurance officer: In large laboratories, a QA officer has the authority and
responsibility for application of the QA program. Ideally, this person should have a staff position
reporting directly to upper management, not a line position. The QA officer should have a
technical education, be acquainted with all aspects of laboratory work, and be familiar with
statistical techniques for data evaluation. The QA officer is responsible for initiating the
program, convincing staff of its value, and providing necessary information and training to the
staff. Once the QA program is functioning, the coordinator conducts frequent (weekly to
monthly) reviews with the laboratory supervisor and staff to determine the current status and
accomplishments of the program and to identify and resolve problems. The QA officer also
reports periodically to management to secure backing in actions necessary to correct problems
that threaten data quality.
c. Staff: Laboratory and field staffs participate with management in planning the QA
program, preparing standard operating procedures, and most importantly, implementing the QC
program in their daily tasks of collecting samples, conducting analyses, performing quality
control checks, and calculating and reporting results. Because the staffs are the first to see
potential problems, they should identify them and work with the supervisor to correct and avoid
them. It is critical to the success of the QA program that staff understand and actively support it.
3. Quality Assurance Program Objectives
The objectives of a QA program include providing data of known quality, ensuring a high
quality of laboratory performance, maintaining continuing assessment of laboratory operations,
identifying weaknesses in laboratory operations, detecting training needs, and improving
documentation and recordkeeping.
4. Elements of a Quality Assurance Program
Each laboratory should develop and implement a written QA plan describing the QA
program and QC activities of the laboratory. The plan should address the following basic
common aspects:
a. Statement of objectives, describing the specific goals of the laboratory.
b. Sampling procedures, including selection of representative sites and specified holding
time and temperature conditions. If data may be subjected to litigation, use chain-of-custody
procedures.
c. Personnel policies, describing specific qualification and training requirements for
supervisors and analysts.
d. Equipment and instrument requirements, providing calibration procedures and frequency
and maintenance requirements.
e. Specifications for supplies, to ensure that reagents and supplies are of high quality and are
tested for acceptability.
f. Analytical methods, i.e., standardized methods established by a standards-setting
organization and validated. Ideally, these laboratory methods have documented precision, bias,
sensitivity, selectivity, and specificity.
g. Analytical quality control measures, including such analytical checks as duplicate
analyses, positive and negative controls, sterility checks, and verification tests.
h. Standard operating procedures (SOPs), i.e., written statement and documentation of all
routine laboratory operations.
i. Documentation requirements, concerning data acquisition, recordkeeping, traceability, and
accountability.
j. Assessment requirements:
1) Internal audits of the laboratory operations, performed by the QA officer and supervisor.
2) On-site evaluations by outside experts to ensure that the laboratory and its personnel are
following an acceptable QA program.
3) Performance evaluation studies, in which the QA officer works with the supervisor to
incorporate unknown challenge samples into routine analytical runs and laboratories are
encouraged to participate in state and national proficiency testing and accreditation programs.
The collaborative studies confirm the abilities of a laboratory to generate acceptable data
comparable to those of other laboratories and identify potential problems.
k. Corrective actions: When problems are identified by the staff, supervisor, and/or QA
coordinator, use standard stepwise procedures to determine the causes and correct them.
Nonconformances identified by external laboratory evaluation are corrected, recorded, and
signed off by the laboratory manager and QA officer.
Detailed descriptions of quality assurance programs are available.1-4
The QC guidelines discussed in Section 9020B and Section 9020C are recommended as
useful source material, but all elements need to be addressed in developing a QA program.
5. References
1. GASKIN, J.E. 1992. Quality Assurance in Water Quality Monitoring. Inland Water
Directorate, Conservation & Protection, Ottawa, Ont., Canada.
2. RATLIFF, T.A., JR. 1990. The Laboratory Quality Assurance System. A Manual of
Quality Procedures with Related Forms. Van Nostrand Reinhold, New York, N.Y.
3. GARFIELD, F.M. 1984. Quality Assurance Principles of Analytical Laboratories. Assoc.
Official Analytical Chemists, Arlington, Va.
4. DUX, J.P. 1983. Quality assurance in the analytical laboratory. Amer. Lab. 26:54.
9020 B.
Intralaboratory Quality Control Guidelines
All laboratories have some intralaboratory QC practices that have evolved from common
sense and the principles of controlled experimentation. A QC program applies practices
necessary to minimize systematic and random errors resulting from personnel, instrumentation,
equipment, reagents, supplies, sampling and analytical methods, data handling, and data
reporting. It is especially important that laboratories performing only a limited amount of
microbiological testing exercise strict QC. A listing of key QC practices is given in Table 9020:I.
Other sources of QC practices are available.1-3 These practices and guidelines will assist
laboratories in establishing and improving QC programs. Laboratories should address all of the
QC guidelines discussed herein, but the depth and details may differ for each laboratory.
1. Personnel
Microbiological testing should be performed by a professional microbiologist or technician
trained in environmental microbiology whenever possible. If not, a professional microbiologist
should be available for guidance. Train and evaluate the analyst in basic laboratory procedures.
The supervisor periodically should review procedures of sample collecting and handling, media
and glassware preparation, sterilization, routine analytical testing, counting, data handling, and
QC techniques to identify and eliminate problems. Management should assist laboratory
personnel in obtaining additional training and course work to advance their skills and career.
2. Facilities
a. Ventilation: Plan well-ventilated laboratories that can be maintained free of dust, drafts,
and extreme temperature changes. Whenever possible, laboratories should have air conditioning
to reduce contamination, permit more stable operation of incubators, and decrease moisture
problems with media and instrumentation.
b. Space utilization: Design and operate the laboratory to minimize through traffic and
visitors, with a separate area for preparing and sterilizing media, glassware, and equipment. Use
a vented laminar-flow hood for dispensing and preparing sterile media, transferring microbial
cultures, or working with pathogenic materials. In smaller laboratories it may be necessary,
although undesirable, to carry out these activities in the same room.
c. Laboratory bench areas: Provide at least 2 m of linear bench space per analyst and
additional areas for preparation and support activities. For stand-up work, typical bench
dimensions are 90 to 97 cm high and 70 to 76 cm deep. For sit-down activities such as
microscopy and plate counting, benches are 75 to 80 cm high. Specify bench tops of stainless
steel, epoxy plastic, or other smooth, impervious surface that is inert and corrosion-resistant, has
a minimum number of seams, and has adequate sealing of any crevices. Install even, glare-free
lighting with about 1000 lux (100 ft-candles) intensity at the working surface.
d. Walls and floors: Assure that walls are covered with a smooth finish that is easily cleaned
and disinfected. Specify floors of smooth concrete, vinyl, asphalt tile, or other impervious, sealed
washable surfaces.
e. Work-area monitoring: Maintain high standards of cleanliness in work areas. Monitor air,
at least monthly, with air density plates. The number of colonies on the air density plate test
should not exceed 160/m2/15 min exposure (15 colonies/plate/15 min).
Plate or the swab method1 can be used weekly or more frequently to monitor bench surface
contamination. Although uniform limits for bacterial density have not been set, each laboratory
can use these tests to establish a base line and take action on a significant increase.
f. Laboratory cleanliness: Regularly clean laboratory rooms and wash benches, shelves,
floors, and windows. Wet-mop floors and treat with a disinfectant solution; do not sweep or
dry-mop. Wipe bench tops and treat with a disinfectant before and after use. Do not permit
laboratory to become cluttered.
3. Laboratory Equipment and Instrumentation
Verify that each item of equipment meets the user’s needs for precision and minimization of
bias. Perform equipment maintenance on a regular basis as recommended by the manufacturer or
obtain preventive maintenance contracts on autoclave, balances, microscopes, and other
equipment. Directly record all quality control checks in a permanent log book.
Use the following quality control procedures:
a. Thermometer/temperature-recording instruments: Check accuracy of thermometers or
temperature-recording instruments semiannually against a certified National Institute of
Standards and Technology (NIST) thermometer or one traceable to NIST and conforming to
NIST specifications. For general purposes use thermometers graduated in increments of 0.5°C or
less. Maintain in water or glycerol for air incubators and refrigerators and glycerol for freezers
and seal in a flask. For a 44.5°C water bath, use a submersible thermometer graduated to 0.2°C
or less. Record temperature check data in a quality control log. Mark the necessary NIST
calibration corrections on each thermometer and incubator, refrigerator, or freezer. When
possible, equip incubators and water baths with temperature-recording instruments that provide a
continuous record of operating temperature.
b. Balances: Follow manufacturer’s instructions in operation and routine maintenance of
analytical and top-loading balances. Balances should be serviced and recalibrated by a
manufacturer technician annually or more often as conditions change or problems occur. In
weighing 2 g or less, use an analytical balance with a sensitivity less than 1 mg at a 10-g load.
For larger quantities use a pan balance with sensitivity of 0.1 g at a 150-g load.
Wipe balance before use with a soft brush. Clean balance pans after use and wipe spills up
immediately with a laboratory tissue. Inspect weights with each use and replace if corroded. Use
only a plastic-tip forceps to handle weights. Check balance and working weights monthly against
a set of reference weights (ANSI/ASTM Class 1 or NIST Class S) for accuracy, precision, and
linearity.4 Record results.
c. pH meter: Use a meter graduated in 0.1 pH units or less, that includes temperature
compensation. Preferably use digital meters and commercial buffer solutions. With each use,
standardize meter with two buffers that bracket the pH of interest and record. Date buffer
solutions when opened and check monthly against another pH meter. Discard solution after each
use and replace buffer supply before expiration date. For full details of pH meter use and
maintenance, see Section 4500-H+.
d. Water purification system: Commercial systems are available that include some
combination of prefiltration, activated carbon, mixed-bed resins, and reverse-osmosis with final
filtration to produce a reagent-grade water. The life of such systems can be extended greatly if
the source water is pretreated by distillation or by reverse osmosis to remove dissolved solids.
Such systems tend to produce the same quality water until resins or activated carbon are near
exhaustion and quality abruptly becomes unacceptable. Some deionization components are
available now that automatically regenerate the ion exchange resins. Do not store reagent water
unless a commercial UV irradiation device is installed and is confirmed to maintain sterility.
Monitor reagent water continuously or daily with a calibrated conductivity meter and
analyze at least annually for trace metals. Replace cartridges at intervals recommended by the
manufacturer based on the estimated usage and source water quality. Do not wait for column
failure. If bacteria-free water is desired, include aseptic final filtration with a 0.22-µm-pore
membrane filter and collect in a sterile container. Monitor treated water for contamination and
replace the filter as necessary.
e. Water still: Stills produce water of a good grade that characteristically deteriorates slowly
over time as corrosion, leaching, and fouling occur. These conditions can be controlled with
proper maintenance and cleaning. Stills efficiently remove dissolved substances but not
dissolved gases or volatile organic chemicals. Freshly distilled water may contain chlorine and
ammonia (NH3). On storage, additional NH3 and CO2 are absorbed from the air. Use softened
water as the source water to reduce frequency of cleaning the still. Drain and clean still and
reservoir according to manufacturer’s instructions and usage.
f. Media dispensing apparatus: Check accuracy of volumes dispensed with a graduated
cylinder at start of each volume change and periodically throughout extended runs. If the unit is
used more than once per day, pump a large volume of hot reagent water through the unit to rinse
between runs. Correct leaks, loose connections, or malfunctions immediately. At the end of the
work day, break apparatus down into parts, wash, rinse with reagent water, and dry. Lubricate
parts according to manufacturer’s instructions or at least once per month.
g. Hot-air oven: Test performance monthly with commercially available Bacillus subtilis
spore strips or spore suspensions. Monitor temperature with a thermometer accurate in the 160 to
180°C range and record results. Use heat-indicating tape to identify supplies and materials that
have been exposed to sterilization temperatures.
h. Autoclave: Record items sterilized, temperature, pressure, and time for each run.
Optimally use a recording thermometer. Check and record operating temperature weekly with a
minimum/maximum thermometer. Test performance with Bacillus stearothermophilus spore
strips, suspensions, or capsules monthly. Use heat-indicating tape to identify supplies and
materials that have been sterilized.
i. Refrigerator: Maintain temperature at 1 to 4°C. Check and record temperature daily and
clean monthly. Identify and date materials stored. Defrost as required and discard outdated
materials quarterly.
j. Freezer: Maintain temperature at −20°C to −30°C. Check and record temperature daily. A
recording thermometer and alarm system are highly desirable. Identify and date materials stored.
Defrost and clean semiannually; discard outdated materials.
k. Membrane filtration equipment: Before use, assemble filtration units and check for leaks.
Discard units if inside surfaces are scratched. Wash and rinse filtration assemblies thoroughly
after use, wrap in nontoxic paper or foil, and sterilize.
l. Ultraviolet lamps: Disconnect lamps monthly and clean bulbs with a soft cloth moistened
with ethanol. Test lamps quarterly with an appropriate (short- or long-wave) UV light
meter*#(3) and replace bulbs if output is less than 70% of the original. For short-wave lamps
used in disinfecting work areas, expose plate count agar spread plates containing 200 to 300
organisms of interest, for 2 min. Incubate plates at 35°C for 48 h and count colonies. Replace
bulb if count is not reduced 99%.
CAUTION: Although short-wave (254-nm) UV light is known to be more dangerous than
long-wave UV (365-nm), both types of UV light can damage eyes and skin and potentially are
carcinogenic.5 Protect eyes and skin from exposure to UV light. (See Section 1090B .)
m. Biohazard hood: Once per month expose plate count agar plates to air flow for 1 h.
Incubate plates at 35°C for 48 h and examine for contamination. A properly operating biohazard
hood should produce no growth on the plates. Disconnect UV lamps and clean monthly by
wiping with a soft cloth moistened with ethanol. Check lamps’ efficiency as specified above.
Inspect cabinet for leaks and rate of air flow quarterly. Use a pressure monitoring device to
measure efficiency of hood performance. Have laminar-flow safety cabinets containing HEPA
filters serviced by the manufacturer. Maintain hoods as directed by the manufacturer.
n. Water bath incubator: Verify that incubators maintain test temperature, such as 35 ±
0.5°C or 44.5 ± 0.2°C. Keep an appropriate thermometer (¶ 3a, above) immersed in the water
bath; monitor and record temperature twice daily (morning and afternoon). For optimum
operation, equip water bath with a gable cover. Use only stainless steel, plastic-coated, or other
corrosion-proof racks. Clean bath as needed.
o. Incubator (air, water jacketed, or aluminum block): Verify that incubators maintain
appropriate test temperatures. Also, verify that cold samples are incubated at the test temperature
for the required time. Check and record temperature twice daily (morning and afternoon) on the
shelves in use. If a glass thermometer is used, submerge bulb and stem in water or glycerine to
the stem mark. For best results use a recording thermometer and alarm system. Place incubator in
an area where room temperature is maintained between 16 and 27°C (60 to 80°F).
p. Microscopes: Use lens paper to clean optics and stage after each use. Cover microscope
when not in use.
Permit only trained technicians to use fluorescence microscope and light source. Monitor
fluorescence lamp with a light meter and replace when a significant loss in fluorescence is
observed. Log lamp operation time, efficiency, and alignment. Periodically check lamp
alignment, particularly when the bulb has been changed; realign if necessary. Use known
positive 4 + fluorescence slides as controls.
4. Laboratory Supplies
a. Glassware: Before each use, examine glassware and discard items with chipped edges or
etched inner surfaces. Particularly examine screw-capped dilution bottles and flasks for chipped
edges that could leak and contaminate the analyst and the area. Inspect glassware after washing
for excessive water beading and rewash if necessary. Make the following tests for clean
glassware as necessary:
1) pH check—Because some cleaning solutions are difficult to remove completely, spot
check batches of clean glassware for pH reaction, especially if soaked in alkali or acid. To test
clean glassware for an alkaline or acid residue add a few drops of 0.04% bromthymol blue
(BTB) or other pH indicator and observe the color reaction. BTB should be blue-green (in the
neutral range).
To prepare 0.04% bromthymol blue indicator solution, add 16 mL 0.01N NaOH to 0.1 g BTB
and dilute to 250 mL with reagent water.
2) Test for inhibitory residues on glassware and plasticware—Certain wetting agents or
detergents used in washing glassware may contain bacteriostatic or inhibiting substances that
require 6 to 12 rinsings to remove all traces and insure freedom from residual bacteriostatic
action. Perform this test annually and before using a new supply of detergent. If prewashed,
presterilized plasticware is used, test it for inhibitory residues. Although the following procedure
describes testing of petri dishes for inhibitory residue, it is applicable to other glass or
plasticware.
a) Procedure—Wash and rinse six petri dishes according to usual laboratory practice and
designate as Group A.
Wash six petri dishes as above, rinse 12 times with successive portions of reagent water, and
designate as Group B.
Rinse six petri dishes with detergent wash water (in use concentration), and air-dry without
further rinsing, and designate as Group C.
Sterilize dishes in Groups A, B, and C by the usual procedure.
For presterilized plasticware, set up six plastic petri dishes and designate them as Group D.
Prepare and sterilize 200 mL plate count agar and hold in a 44 to 46°C water bath.
Prepare a culture of E. aerogenes known to contain 50 to 150 colony-forming units/mL.
Preliminary testing may be necessary to achieve this count range. Inoculate three dishes from
each test group with 0.1 mL and the other three dishes from each group with 1 mL culture.
Analyze the four sets of six plates each, following heterotrophic plate count method (Section
9215B), and incubate at 35°C for 48 h. Count plates with 30 to 300 colonies and record results as
CFU/ mL.
b) Interpretation of results—Difference in averaged counts on plates in Groups A through D
should be less than 15% if there are no toxic or inhibitory effects.
Differences in averaged counts of less than 15% between Groups A and B and greater than
15% between Groups A and C indicate that the cleaning detergent has inhibitory properties that
are eliminated during routine washing. Differences between B and D greater than 15% indicate
an inhibitory residue.
b. Utensils and containers for media preparation: Use utensils and containers of borosilicate
glass, stainless steel, aluminum, or other corrosion-resistant material (see Section 9030). Do not
use copper utensils.
c. Dilution water bottles: Use scribed bottles made of nonreactive borosilicate glass or
plastic with screwcaps containing inert liners. Clean before use. Disposable plastic bottles
prefilled with dilution water are available commercially and are acceptable. Before use of each
lot, check pH and volume and examine sterile bottles of dilution water for a precipitate; discard
if present. Reclean bottles with acid if necessary, and remake the dilution water. If precipitate
repeats, procure a different source of bottles.
d. Reagent-grade water quality: The quality of water obtainable from a water purification
system differs with the system used and its maintenance. See ¶ 3d and ¶ 3e above.
Recommended limits for reagent water quality are given in Table 9020:II. If these limits are not
met, investigate and correct or change water source. Although pH measurement of reagent water
is characterized by drift, extreme readings are indicative of chemical contamination.
e. Use test for evaluation of reagent water, media, and membranes: When a new lot of
culture medium, membrane filters, or a new source of reagent-grade water is to be used make
comparison tests, at least quarterly, of the current lot in use (reference lot) against the new lot
(test lot).
1) Procedure—Use a single batch of control water (redistilled or distilled water polished by
deionization), glassware, membrane filters, or other needed materials to control all variables
except the one factor under study. Make parallel pour or spread plate or membrane filter plate
tests on reference lot and test lot, according to procedures in Section 9215 and Section 9222. As
a minimum, make single analyses on five different water samples positive for the target
organism. Replicate analyses and additional samples can be tested to increase the sensitivity of
detecting differences between reference and test lots.
When conducting the use test on reagent water, perform the quantitative bacterial tests in
parallel using a known high-quality water as a control water. Prepare dilution/rinse water and
media with new source of reagent and control water. Test water for all uses (dilution, rinse,
media preparation, etc.).
2) Counting and calculations—After incubation, compare bacterial colonies from the two
lots for size and appearance. If colonies on the test lot plates are atypical or noticeably smaller
than colonies on the reference lot plates, record the evidence of inhibition or other problem,
regardless of count differences. Count plates and calculate the individual count per 1 mL or per
100 mL. Transform the count to logarithms and enter the log-transformed results for the two lots
in parallel columns. Calculate the difference, d, between the two transformed results for each
sample, including the + or − sign, the mean,
and the standard deviation sd of these
differences (see Section 1010B).
Calculate Student’s t statistic, using the number of samples as n:
These calculations may be made with various statistical software packages available for
personal computers.
3) Interpretation—Use the critical t value, from a Student’s t table for comparison against
the calculated value. At the 0.05 significance level this value is 2.78 for five samples (four
degrees of freedom). If the calculated t value does not exceed 2.78, the lots do not produce
significantly different results and the test lot is acceptable. If the calculated t value exceeds 2.78,
the lots produce significantly different results and the test lot is unacceptable.
If the colonies are atypical or noticeably smaller on the test lot or the Student’s t exceeds
2.78, review test conditions, repeat the test, and/or reject the test lot and obtain another one.
f. Reagents: Because reagents are an integral part of microbiological analyses, their quality
must be assured. Use only chemicals of ACS or equivalent grade because impurities can inhibit
bacterial growth, provide nutrients, or fail to produce the desired reaction. Date chemicals and
reagents when received and when first opened for use. Make reagents to volume in volumetric
flasks and transfer for storage to good-quality inert plastic or borosilicate glass bottles with
borosilicate, polyethylene, or other plastic stoppers or caps. Label prepared reagents with name
and concentration, date prepared, and initials of preparer. Include positive and negative control
cultures with each series of cultural or biochemical tests.
g. Dyes and stains: In microbiological analyses, organic chemicals are used as selective
agents (e.g., brilliant green), as indicators (e.g., phenol red), and as microbiological stains (e.g.,
Gram stain). Dyes from commercial suppliers vary from lot to lot in percent dye, dye complex,
insolubles, and inert materials. Because dyes for microbiology must be of proper strength and
stability to produce correct reactions, use only dyes certified by the Biological Stain
Commission. Check bacteriological stains before use with at least one positive and one negative
control culture and record results.
h. Membrane filters and pads: The quality and performance of membrane filters vary with
the manufacturer, type, brand, and lot. These variations result from differences in manufacturing
methods, materials, quality control, storage conditions, and application.
1) Membrane filters and pads for water analyses should meet the following specifications:
a) Filter diam 47 mm, mean pore diam 0.45 µm. Alternate filter and pore sizes may be used
if the manufacturer provides data verifying performance equal to or better than that of
47-mm-diam, 0.45-µm-pore size filter. At least 70% of filter area must be pores.
b) When filters are floated on reagent water, the water diffuses uniformly through the filters
in 15 s with no dry spots on the filters.
c) Flow rates are at least 55 mL/min/cm2 at 25°C and a differential pressure of 93 kPa.
d) Filters are nontoxic, free of bacterial-growth-inhibiting or stimulating substances, and free
of materials that directly or indirectly interfere with bacterial indicator systems in the medium;
ink grid is nontoxic. The arithmetic mean of five counts on filters must be at least 90% of the
arithmetic mean of the counts on five agar spread plates using the same sample volumes and agar
media.
e) Filters retain the organisms from a 100-mL suspension of Serratia marcescens containing
1 × 103 cells.
f) Water-extractables in filter do not exceed 2.5% after the membrane is boiled in 100 mL
reagent water for 20 min, dried, cooled, and brought to constant weight.
g) Absorbent pad has diam 47 mm, thickness 0.8 mm, and is capable of absorbing 2.0 ± 0.2
mL Endo broth.
h) Pads release less than 1 mg total acidity calculated as CaCO3 when titrated to the
phenolphthalein end point with 0.02N NaOH.
i) If filter and absorbent pad are not sterile, they should not be degraded by sterilization at
121°C for 10 min. Confirm sterility by absence of growth when a membrane filter is placed on a
pad saturated with tryptone glucose extract broth or tryptone glucose extract agar and incubated
at 35 ±0.5°C for 24 h.
j) Some lots of membrane filters yield low recoveries, poor differentiation, or malformation
of colonies due to toxicity, chemical composition, or structural defects.6 Perform the use test (¶
4e) on new lots of filters.
2) Standardized tests:
Standardized tests are available for evaluating retention, recovery, extractables, and flow rate
characteristics of membrane filters.7
Some manufacturers provide information beyond that required by specifications and certify
that their membranes are satisfactory for water analysis. They report retention, pore size, flow
rate, sterility, pH, percent recovery, and limits for specific inorganic and organic chemical
extractables. Although the standard membrane filter evaluation tests were developed for the
manufacturers, a laboratory can conduct its own tests.
To maintain quality control inspect each lot of membranes before use and during testing to
insure they are round and pliable, with undistorted gridlines after autoclaving. After incubation,
colonies should be well-developed with well-defined color and shape as defined by the test
procedure. The gridline ink should not channel growth along the ink line nor restrict colony
development. Colonies should be distributed evenly across the membrane surface.
i. Culture media: Because cultural methods depend on properly prepared media, use the best
available materials and techniques in media preparation, storage, and application. For control of
quality, use commercially prepared media whenever available but note that such media may vary
in quality among manufacturers and even from lot to lot from the same manufacturer.
Order media in quantities to last no longer than 1 year. Use media on a first-in, first-out
basis. When practical, order media in quarter pound (114 g) multiples rather than one pound
(454 g) bottles, to keep the supply sealed as long as possible. Record kind, amount, and
appearance of media received, lot number, expiration date, and dates received and opened.
Check inventory quarterly for reordering.
Store dehydrated media at an even temperature in a cool dry place, away from direct
sunlight. Discard media that cake, discolor, or show other signs of deterioration. If expiration
date is given by manufacturer, discard unused media after that date. A conservative time limit for
unopened bottles is 2 years at room temperature. Compare recovery of newly purchased lots of
media against proven lots, using recent pure-culture isolates and natural samples.
Use opened bottles of media within 6 months. Dehydrated media are hygroscopic. Protect
opened bottles from moisture. Close bottles as tightly as possible, immediately after use. If
caking or discoloration of media occurs, discard media. Store opened bottles in a dessicator.
1) Preparation of media—Prepare media in containers that are at least twice the volume of
the medium being prepared. Stir media, particularly agars, while heating. Avoid scorching or
boil-over by using a boiling water bath for small batches of media and by continually attending
to larger volumes heated on a hot plate or gas burner. Preferably use hot plate-magnetic stirrer
combinations. Label and date prepared media. Prepare media in reagent water. Measure water
volumes and media with graduates or pipets conforming to NIST and APHA standards,
respectively. Do not use blow-out pipets. After preparation and storage, remelt agar media in
boiling water or flowing steam.
Check and record pH of a portion of each medium after sterilization and cooling. Check pH
of solid medium with a surface probe. Record results. Make minor adjustments in pH (<0.5 pH
units) with 1N NaOH or HCl solution to the pH specified in formulation. If the pH difference is
larger than 0.5 units, discard the batch and check preparation instructions and pH of reagent
water to resolve the problem. Incorrect pH values may be due to reagent water quality, medium
deterioration, or improper preparation. Review instructions for preparation and check water pH.
If water pH is unsatisfactory, prepare a new batch of medium using water from a new source (see
Section 9020B.3d and e). If water is satisfactory, remake medium and check; if pH is again
incorrect, prepare medium from another bottle.
Record pH problems in the media record book and inform the manufacturer if the medium is
indicated as the source of error. Examine prepared media for unusual color, darkening, or
precipitation and record observations. Consider variations of sterilization time and temperature
as possible causes for problems. If any of the above occur, discard the medium.
2) Sterilization—Sterilize media at 121 to 124°C for the minimum time specified. A
double-walled autoclave permits maintenance of full pressure and temperature in the jacket
between loads and reduces chance for heat damage. Follow manufacturer’s directions for
sterilization of specific media. The required exposure time varies with form and type of material,
type of medium, presence of carbohydrates, and volume. Table 9020:III gives guidelines for
typical items. Do not expose media containing carbohydrates to the elevated temperatures for
more than 45 min. Exposure time is defined as the period from initial exposure to removal from
the autoclave.
Some currently available autoclave models are automatic and include features such as
vertical sliding, self-sealing and opening doors, programmable sterilization cycles, and
continuous multipoint monitoring of chamber temperature and pressure. These units also may
incorporate solution cooling and vapor removal features. When sterilizer design includes heat
exchangers and solution cooling features as part of a factory-programmed liquid cycle, strict
adherence to the 45-min total elapsed time in the autoclave is not necessary provided that
printout records verify normal cycle operation and chamber cooling during exhaust and vapor
removal.
Remove sterilized media from autoclave as soon as chamber pressure reaches zero, or, if a
fully automatic model is used, as soon as the door opens. Do not reautoclave media.
Check effectiveness of sterilization weekly by placing Bacillus stearothermophilus spore
suspensions or strips (commercially available) inside glassware. Sterilize at 121°C for 15 min.
Place in trypticase soy broth tubes and incubate at 55°C for 48 h. If growth of the autoclaved
spores occurs after incubation at 55°C, sterilization was inadequate. A small, relatively
inexpensive 55°C incubator is available commercially.†#(4)
Sterilize heat-sensitive solutions or media by filtration through a 0.22-µm-pore-diam filter in
a sterile filtration and receiving apparatus. Filter and dispense medium in a safety cabinet or
biohazard hood if available. Sterilize glassware (pipets, petri dishes, sample bottles) in an
autoclave or an oven at 170°C for 2 h. Sterilize equipment, supplies, and other solid or dry
materials that are heat-sensitive, by exposing to ethylene oxide in a gas sterilizer. Use
commercially available spore strips or suspensions to check dry heat and ethylene oxide
sterilization.
3) Use of agars and broths—Temper melted agars in a water bath at 44 to 46°C until used
but do not hold longer than 3 h. To monitor agar temperature, expose a bottle of water or
medium to the same heating and cooling conditions as the agar. Insert a thermometer in the
monitoring bottle to determine when the temperature is 45 to 46°C and suitable for use in pour
plates. If possible, prepare media on the day of use. After pouring agar plates for streaking, dry
agar surfaces by keeping dish slightly open for at least 15 min in a bacteriological hood to avoid
contamination. Discard unused liquid agar; do not let harden or remelt for later use.
Handle tubes of sterile fermentation media carefully to avoid entrapping air in inner tubes,
thereby producing false positive reactions. Examine freshly prepared tubes to determine that gas
bubbles are absent.
4) Storage of media—Prepare media in amounts that will be used within holding time limits
given in Table 9020:IV. Protect media containing dyes from light; if color changes occur,
discard the media. Refrigerate poured agar plates not used on the day of preparation. Seal agar
plates with loose-fitting lids in plastic bags if held more than 2 d. Prepare broth media that will
be stored for more than 2 weeks in screw-cap tubes, other tightly sealed tubes, or in loose-capped
tubes placed in a sealed plastic bag or other tightly sealed container to prevent evaporation.
Mark liquid level in several tubes and monitor for loss of liquid. If loss is 10% or more,
discard the batch. If media are refrigerated, incubate overnight at test temperature before use and
reject the batch if false positive responses occur. Prepared sterile broths and agars available from
commercial sources may offer advantages when analyses are done intermittently, when staff is
not available for preparation work, or when cost can be balanced against other factors of
laboratory operation. Check performance of these media as described in ¶ 5 below.
5) Quality control of prepared media—Maintain in a bound book a complete record of each
prepared batch of medium with name of preparer and date, name and lot number of medium,
amount of medium weighed, volume of medium prepared, sterilization time and temperature, pH
measurements and adjustments, and preparations of labile components. Compare quantitative
recoveries of new lots with previously acceptable ones. Include sterility and positive and
negative control culture checks on all media as described below.
5. Standard Operating Procedures (SOPs)
SOPs are the operational backbone of an analytical laboratory. SOPs describe in detail all
laboratory operations such as preparation of reagents, reagent water, standards, culture media,
proper use of balances, sterilization practices, and dishwashing procedures, as well as methods of
sampling, analysis, and quality control. The SOPs are unique to the laboratory. They describe the
tasks as performed on a day-to-day basis, tailored to the laboratory’s own equipment,
instrumentation, and sample types. The SOPs guide routine operations by each analyst, help to
assure uniform operations, and provide a solid training tool.
6. Sampling
a. Planning: Microbiologists should participate in the planning of monitoring programs that
will include microbial analyses. They can provide valuable expertise on the selection of
sampling sites, number of samples and analyses needed, workload, and equipment and supply
needs. For natural waters, knowledge of the probable microbial densities, and the impact of
season, weather, tide and wind patterns, known sources of pollution, and other variables, are
needed to formulate the most effective sampling plan.
b. Methods: Sampling plans must be specific for each sampling site. Prior sampling
guidance can be only general in nature, addressing the factors that must be considered for each
site. Sampling SOPs describe sampling equipment, techniques, frequency, holding times and
conditions, safety rules, etc., that will be used under different conditions for different sites. From
the information in these SOPs sampling plans will be drawn up.
7. Analytical Methods
a. Method selection: Because minor variations in technique can cause significant changes in
results, microbiological methods must be standardized so that uniform data result from multiple
laboratories. Select analytical methods appropriate for the sample type from Standard Methods
or other source of standardized methods and ensure that methods have been validated in a
multi-laboratory study with the sample types of interest.
b. Data objectives: Review available methods and determine which produce data to meet the
program’s needs for precision, bias, specificity, selectivity, and detection limit. Ensure that the
methods have been demonstrated to perform within the above specifications for the samples of
interest.
c. Internal QC: The written analytical methods should contain required QC checks of
positive and negative control cultures, sterile blank, replicate analyses (precision), and a known
quantitative culture, if available.
d. Method SOPs: As part of the series of SOPs, provide each analyst with a copy of the
analytical methods written in step-wise fashion exactly as they are to be performed and specific
to the sample type, equipment, and instrumentation used in the laboratory.
8. Analytical Quality Control Procedures
a. General quality control procedures:
1) New methods—Conduct parallel tests with the standard procedure and a new method to
determine applicability and comparability. Perform at least 100 parallel tests across seasons of
the year before replacement with the new method for routine use.
2) Comparison of plate counts—For routine performance evaluation, repeat counts on one or
more positive samples at least monthly and compare the counts with those of other analysts
testing the same samples. Replicate counts for the same analyst should agree within 5% and
those between analysts should agree within 10%. See Section 9020B.10b for a statistical
calculation of data precision.
3) Control cultures—For each lot of medium check analytical procedures by testing with
known positive and negative control cultures for the organism(s) under test. See Table 9020:V
for examples of test cultures.
4) Duplicate analyses—Perform duplicate analyses on 10% of samples and on at least one
sample per test run. A test run is defined as an uninterrupted series of analyses. If the laboratory
conducts less than 10 tests/week, make duplicate analyses on at least one sample each week.
5) Sterility checks—For membrane filter tests, check sterility of media, membrane filters,
buffered dilution and rinse water, pipets, flasks and dishes, and equipment as a minimum at the
end of each series of samples, using sterile reagent water as the sample. If contaminated, check
for the source. For multiple-tube and presence-absence procedures, check sterility of media,
dilution water, and glassware. To test sterility of media, incubate a representative portion of each
batch at an appropriate temperature for 24 to 48 h and observe for growth. Check each batch of
buffered dilution water for sterility by adding 20 mL water to 100 mL of a nonselective broth.
Alternatively, aseptically pass 100 mL or more dilution water through a membrane filter and
place filter on growth medium suitable for heterotrophic bacteria. Incubate at 35 ± 0.5°C for 24 h
and observe for growth. If any contamination is indicated, determine the cause and reject
analytical data from samples tested with these materials. Request immediate resampling and
reanalyze.
b. Precision of quantitative methods: Calculate precision of duplicate analyses for each
different type of sample examined, for example, drinking water, ambient water, wastewater, etc.,
according to the following procedure:
1) Perform duplicate analyses on first 15 positive samples of each type, with each set of
duplicates analyzed by a single analyst. If there is more than one analyst, include all analysts
regularly running the tests, with each analyst performing approximately an equal number of
tests. Record duplicate analyses as D1 and D2.
2) Calculate the logarithm of each result. If either of a set of duplicate results is <1, add 1 to
both values before calculating the logarithms.
3) Calculate the range (R) for each pair of transformed duplicates as the mean (î ) of these
ranges.
See sample calculation in Table 9020:VI.
4) Thereafter, analyze 10% of routine samples in duplicate. Transform the duplicates as in ¶
2) and calculate their range. If the range is greater than 3.27 R, there is greater than 99%
probability that the laboratory variability is excessive. Determine if increased imprecision is
acceptable; if not, discard all analytical results since the last precision check (see Table
9020:VII). Identify and resolve the analytical problem before making further analyses.
5) Update the criterion used in ¶ 4) by periodically repeating the procedures of ¶s 1) through
3) using the most recent sets of 15 duplicate results.
9. Verification
For the most part, the confirmation/verification procedures for drinking water differ from
those for other waters because of specific regulatory requirements.
a. Multiple-tube fermentation (MTF) methods:
1) Total coliform procedure (Section 9221B)
a) Drinking water—Carry samples through confirmed phase only. Verification is not
required. For QC purposes, if normally there are no positive results, analyze at least one positive
source water quarterly to confirm that the media produce appropriate responses. For samples
with a history of heavy growth without gas in presumptive-phase tubes, carry the tubes through
the confirmed phase to check for false negative responses for coliform bacteria. Verify any
positives for fecal coliforms or E. coli.
b) Other water types—Verify by performing the completed MTF Test on 10% of samples
positive through the confirmed phase.
2) Enzyme substrate coliform test (total coliform/E. coli) (Section 9223B)
a) Drinking water—Verify at least 5% of total coliform positive results from enzyme
substrate coliform tests by inoculating growth from a known positive sample and testing for
lactose fermentation or for β-D-galactopyranosidase by the o-nitrophenyl-β-D-galactopyranoside
(ONPG) test and indophenol by the cytochrome oxidase (CO) test. See Section 9225D for these
tests. Coliforms are ONPG-positive and cytochrome-oxidase-negative. Verify E. coli using the
EC MUG test (see Section 9221F).
b) Other water types—Verify at least 10% of total coliform positive samples as in ¶ 2a
above.
3) Fecal streptococci procedure—Verify as in 9230C.5. Growth of catalase-negative,
gram-positive cocci on bile esculin agar at 35°C and in brain-heart infusion broth at 45°C
verifies the organisms as fecal streptococci. Growth at 45°C and in 6.5% NaCl broth indicates
the streptococci are members of the enterococcus group.
4) Include known positive and negative pure cultures as a QC check.
b. Membrane filter methods:
1) Total coliform procedures
a) Drinking water—Pick all, up to 5 typical and 5 atypical (nonsheen) colonies from positive
samples on M-Endo medium and verify as in Section 9222B.5 f. Also verify any positives for
fecal coliforms or E. coli. If there are no positive samples, test at least one known positive source
water quarterly.
b) Other water types—Verify positives monthly by picking at least 10 sheen colonies from a
positive water sample as in Section 9222B.5 f. Adjust counts based on percent verification.
c) To determine false negatives, pick representative atypical colonies of different
morphological types and verify as in Section 9222B.5 f.
2) Fecal coliform procedure
a) Verify positives monthly by picking at least 10 blue colonies from one positive sample.
Verify in lauryl tryptose broth and EC broth as in Section 9221B.3 and Section 9221E. Adjust
counts based on percent verification.
b) To determine false negatives, pick representative atypical colonies of different
morphological types and verify as in Section 9221B.3 and Section 9221E.
3) Escherichia coli procedure
a) Drinking water—Verify at least 5% of MUG-positive and MUG-negative results. Pick
from well-isolated sheen colonies that fluoresce on nutrient agar with MUG (NA MUG), taking
care not to pick up medium, which can cause a false positive response. Also verify nonsheen
colonies that fluoresce. Verify by performing the citrate test and the indole test as described in
Section 9225D, but incubate indole test at 44.5°C. E. coli are indole-positive and yield no growth
on citrate.
b) Other water types—Verify one positive sample monthly as in ¶ a) above. Adjust counts
based on percentage of verification.
4) Fecal streptococci procedure—Pick to verify monthly at least 10 isolated esculin-positive
red colonies from m-Enterococcus agar to brain heart infusion (BHI) media. Verify as described
in Section 9230C. Adjust counts based on percentage of verification.
5) Enterococci procedures—Pick to verify monthly at least 10 well-isolated pink to red
colonies with black or reddish-brown precipitate from EIA agar. Transfer to BHI media as
described in Section 9230C. Adjust counts based on percentage of verification.
6) Include known positive and negative pure cultures as a quality control check.
9. Documentation and Recordkeeping
a. QA plan: The QA program documents management’s commitment to a QA policy and
sets forth the requirements needed to support program objectives. The program describes overall
policies, organization, objectives, and functional responsibilities for achieving the quality goals.
In addition, the program should develop a project plan that specifies the QC requirements for
each project. The plan specifies the QC activities required to achieve the data representativeness,
completeness, comparability, and compatibility. Also, the QA plan should include a program
implementation plan that ensures maximum coordination and integration of QC activities within
the overall program (sampling, analyses, and data handling).
b. Sampling records: A written SOP for sample handling records sample collection, transfer,
storage, analyses, and disposal. The record is most easily kept on a series of printed forms that
prompt the user to provide all the necessary information. It is especially critical that this record
be exact and complete if there is any chance that litigation may occur. Such record systems are
called chain of custody. Because laboratories do not always know whether analytical results will
be used in future litigation, some maintain chain-of-custody on all samples. Details on chain of
custody are available in Section 1060B and elsewhere.1
c. Recordkeeping: An acceptable recordkeeping system provides needed information on
sample collection and preservation, analytical methods, raw data, calculations through reported
results, and a record of persons responsible for sampling and analyses. Choose a format
agreeable to both the laboratory and the customer (the data user). Ensure that all data sheets are
signed and dated by the analyst and the supervisor. The preferable record form is a bound and
page-numbered notebook, with entries in ink and a single line drawn through any change with
the correction entered next to it.
Keep records of microbiological analyses for at least 5 years. Actual laboratory reports may
be kept, or data may be transferred to tabular summaries, provided that the following information
is included: date, place, and time of sampling, name of sample collector; identification of
sample; date of receipt of sample and analysis; person(s) responsible for performing analysis;
analytical method used; the raw data and the calculated results of analysis. Verify that each result
was entered correctly from the bench sheet and initialed by the analyst. If an information storage
and retrieval system is used, double check data on the printouts.
10. Data Handling
a. Distribution of bacterial populations: In most chemical analyses the distribution of
analytical results follows the Gaussian curve, which has symmetrical distribution of values about
the mean (see Section 1010B). Microbial distributions are not necessarily symmetrical. Bacterial
counts often are characterized as having a skewed distribution because of many low values and a
few high ones. These characteristics lead to an arithmetic mean that is considerably larger than
the median. The frequency curve of this distribution has a long right tail, such as that shown in
Figure 9020:1, and is said to display positive skewness.
Application of the most rigorous statistical techniques requires the assumption of
symmetrical distributions such as the normal curve. Therefore it usually is necessary to convert
skewed data so that a symmetrical distribution resembling the normal distribution results. An
approximately normal distribution can be obtained from positively skewed data by converting
numbers to their logarithms, as shown in Table 9020:VIII. Comparison of the frequency tables
for the original data (Table 9020:IX) and their logarithms (Table 9020:X) shows that the
logarithms approximate a symmetrical distribution.
b. Central tendency measures of skewed distribution: If the logarithms of numbers from a
positively skewed distribution are approximately normally distributed, the original data have a
log-normal distribution. The best estimate of central tendency of log-normal data is the
geometric mean, defined as:
and
that is, the geometric mean is equal to the antilog of the arithmetic mean of the logarithms. For
example, the following means calculated from the data in Table 9020:VIII are drastically
different.
geometric mean
and arithmetic mean
Therefore, although regulations or tradition may require or cause microbiological data to be
reported as the arithmetic mean or median, the preferred statistic for summarizing
microbiological monitoring data is the geometric mean. An exception may be in the evaluation
of data for risk assessment. The arithmetic mean may be a better measure for this purpose
because it may generate a higher central tendency value and possibly provide a greater safety
factor.8
c. ‘‘Less than’’ (<) values: There has always been uncertainty as to the proper way to
include ‘‘less than’’ values in calculation and evaluation of microbiological data because such
values cannot be treated statistically without modification. Proposed modifications involve
changing such numbers to zero, choosing values halfway between zero and the ‘‘less than’’
value, or assigning the ‘‘less than’’ value itself, i.e., changing <1 values to 1, 1/2, or 0.
There are valid reasons for not including < values, whether modified or not. If the database
is fairly large with just a few < values, the influence of these uncertain values will be minimal
and of no benefit. If the database is small or has a relatively large number of < values, inclusion
of modified < values would exert an undue influence on the final results and could result in an
artificial negative or positive bias. Including < values is particularly inappropriate if the < values
are <100, <1000, or higher because the unknown true values could be anywhere from 0 to 99, 0
to 999, etc. When < values are first noted, adjust or expand test volumes. The only exception to
this caution would be regulatory testing with defined compliance limits, such as the <1/100 mL
values reported for drinking water systems where the 100-mL volume is required.
11. References
1. BORDNER, R.H., J.A. WINTER & P.V. SCARPINO, eds. 1978. Microbiological Methods for
Monitoring the Environment, Water and Wastes. EPA-600/8-78-017, Environmental
Monitoring & Support Lab., U.S. Environmental Protection Agency, Cincinnati, Ohio.
2. AMERICAN SOCIETY FOR TESTING AND MATERIALS. 1995. Standard guide for good
laboratory practices in laboratories engaged in sampling and analysis of water.
D-3856-95, Annual Book of ASTM Standards, Vol. 11.01, American Soc. Testing &
Materials, Philadelphia, Pa.
3. AMERICAN SOCIETY FOR TESTING AND MATERIALS. 1996. Standard practice for
writing quality control specifications for standard test methods for water analysis.
D-5847-96, Annual Book of ASTM Standards, Vol. 11.01, American Soc. Testing &
Materials, West Conshohocken, Pa.
4. AMERICAN SOCIETY FOR TESTING AND MATERIALS. 1993. Annual Book of ASTM
Standards, Vol. 14.02, General Methods and Instrumentation. E-319-86 (reapproved
1993), Standard Practice for Evaluation of Single-Pan Mechanical Balances, and
E-898-88 (reapproved 1993), Standard Method of Testing Top-Loading,
Direct-Reading Laboratory Scales and Balances. American Soc. Testing & Materials,
Philadelphia, Pa.
5. SCHMITZ, S., C. GARBE, B. TEBBE & C. ORFANOS. 1994. Long wave ultraviolet radiation
(UVA) and skin cancer. Hautarzt 45:517.
6. BRENNER, K. & C.C. RANKIN. 1990. New screening test to determine the acceptability of
0.45 µm membrane filters for analysis of water. Appl. Environ. Bacteriol. 56:54.
7. AMERICAN SOCIETY FOR TESTING AND MATERIALS. 1977. Annual Book of ASTM
Standards. Part 31, Water. American Soc. Testing & Materials, Philadelphia, Pa.
8. HAAS, C.N. 1996. How to average microbial densities to characterize risk. Water Res.
30:1036.
9020 C.
Interlaboratory Quality Control
1. Background
Interlaboratory QC programs are a means of establishing an agreed-upon, common
performance criteria system that will assure an acceptable level of data quality and comparability
among laboratories with similar interests and/or needs.
These systems may be volunteer, such as that for the cities in the Ohio River Valley Water
Sanitation Commission (ORSANCO), or regulatory, such as the Federal Drinking Water
Laboratory Certification Program (see below). Often, the term ‘‘accreditation’’ is used
interchangeably with certification. Usually, interlaboratory quality control programs have three
elements: uniform criteria for laboratory operations, external review of the program, and
external proficiency testing.
2. Uniform Criteria
Interlaboratory quality control programs begin as a volunteer or mandatory means of
establishing uniform laboratory standards for a specific purpose. The participants may be from
one organization or a group of organizations having common interests or falling under common
regulations. Often one group or person may agree to draft the criteria. If under regulation, the
regulating authority may set the criteria for compliance-monitoring analyses.
Uniform sampling and analytical methods and quality control criteria for personnel,
facilities, equipment, instrumentation, supplies, and data handling and reporting are proposed,
discussed, reviewed, modified if necessary, and approved by the group for common use. Criteria
identified as necessary for acceptable data quality should be mandatory. A formal document is
prepared and provided to all participants.
The QA/QC responsibilities of management, supervisors, and technical staff are described in
9020A. In large laboratories, a QA officer is assigned as a staff position but may be the
supervisor or other senior person in smaller laboratories.
After incorporation into laboratory operations and confirmation that the QA program has
been adapted and is in routine use, the laboratory supervisor and the QA officer conduct an
internal program review of all operations and records for acceptability, to identify possible
problems and assist in their resolution. If this is done properly, there should be little concern that
subsequent external reviews will find major problems.
3. External Program Review
Once a laboratory has a QA program in place, management informs the organization and a
qualified external QA person or team arranges an on-site visit to evaluate the QA program for
acceptability and to work with the laboratory to solve any problems. An acceptable rating
confirms that the laboratory’s QA program is operating properly and that the laboratory has the
capability of generating valid defensible data. Such on-site evaluations are repeated and may be
announced or unannounced.
4. External Proficiency Testing
Whenever practical, the external organization conducts formal performance evaluation
studies among all participant laboratories. Challenge samples are prepared and sent as unknowns
on a set schedule for analyses and reporting of results. The reported data are coded for
confidentiality and evaluated according to an agreed-upon scheme. The results are summarized
for all laboratories and individual laboratory reports are sent to participants. Results of such
studies indicate the quality of routine analyses of each laboratory as compared to group
performance. Also, results of the group as a whole characterize the performance that can be
expected for the analytical methods tested.
5. Example Program
In the Federal Drinking Water Laboratory Certification Program, public water supply
laboratories must be certified according to minimal criteria and procedures and quality assurance
described in the EPA manual on certification:1 criteria are established for laboratory operations
and methodology; on-site inspections are required by the certifying state agency or its surrogate
to verify minimal standards; annually, laboratories are required to perform acceptably on
unknown samples in formal studies, as samples are available; the responsible authority follows
up on problems identified in the on-site inspection or performance evaluation and requires
corrections within a set period of time. Individual state programs may exceed the federal criteria.
On-site inspections of laboratories in the present certification program show that primary
causes for discrepancies in drinking water laboratories have been inadequate equipment,
improperly prepared media, incorrect analytical procedures, and insufficiently trained personnel.
6. References
1. U.S. ENVIRONMENTAL PROTECTION AGENCY. 1997. Manual for the Certification of
Laboratories Analyzing Drinking Water, 4th ed. EPA-814B-92-002, U.S.
Environmental Protection Agency, Cincinnati, Ohio.
9030
LABORATORY APPARATUS*#(5)
9030 A.
Introduction
This section contains specifications for microbiological laboratory equipment. For testing
and maintenance procedures related to quality control, see Section 9020.
9030 B.
Equipment Specifications
1. Incubators
Incubators must maintain a uniform and constant temperature at all times in all areas, that is,
they must not vary more than ±0.5°C in the areas used. Obtain such accuracy by using a
water-jacketed or anhydric-type incubator with thermostatically controlled low-temperature
electric heating units properly insulated and located in or adjacent to the walls or floor of the
chamber and preferably equipped with mechanical means of circulating air.
Incubators equipped with high-temperature heating units are unsatisfactory, because such
sources of heat, when improperly placed, frequently cause localized overheating and excessive
drying of media, with consequent inhibition of bacterial growth. Incubators so heated may be
operated satisfactorily by replacing high-temperature units with suitable wiring arranged to
operate at a lower temperature and by installing mechanical air-circulation devices. It is
desirable, where ordinary room temperatures vary excessively, to keep laboratory incubators in
special rooms maintained at a few degrees below the recommended incubator temperature.
Alternatively, use special incubating rooms well insulated and equipped with properly
distributed heating units, forced air circulation, and air exchange ports, provided that they
conform to desired temperature limits. When such rooms are used, record the daily temperature
range in areas where plates or tubes are incubated. Provide incubators with open metal wire or
perforated sheet shelves so spaced as to assure temperature uniformity throughout the chamber.
Leave a 2.5-cm space between walls and stacks of dishes or baskets of tubes.
Maintain an accurate thermometer, traceable to the National Institute of Standards and
Technology (NIST), with the bulb immersed in liquid (glycerine, water, or mineral oil) on each
shelf in use within the incubator and record daily temperature readings (preferably morning and
afternoon). It is desirable, in addition, to maintain a maximum and minimum registering
thermometer within the incubator on the middle shelf to record the gross temperature range over
a 24-h period. At intervals, determine temperature variations within the incubator when filled to
maximum capacity. Install a recording thermometer whenever possible, to maintain a continuous
and permanent record of temperature.
Ordinarily, a water bath with a gabled cover to reduce water and heat loss, or a solid heat
sink incubator, is required to maintain a temperature of 44.5 ± 0.2°C. If satisfactory temperature
control is not achieved, provide water recirculation. Keep water depth in the incubator sufficient
to immerse tubes to upper level of media.
2. Hot-Air Sterilizing Ovens
Use hot-air sterilizing ovens of sufficient size to prevent internal crowding; constructed to
give uniform and adequate sterilizing temperatures of 170 ± 10°C; and equipped with suitable
thermometers. Optionally use a temperature-recording instrument.
3. Autoclaves
Use autoclaves of sufficient size to prevent internal crowding; constructed to provide
uniform temperatures within the chambers (up to and including the sterilizing temperature of
121°C); equipped with an accurate thermometer the bulb of which is located properly on the
exhaust line so as to register minimum temperature within the sterilizing chambers
(temperature-recording instrument is optional); equipped with pressure gauge and properly
adjusted safety valves connected directly with saturated-steam supply lines equipped with
appropriate filters to remove particulates and oil droplets or directly to a suitable special steam
generator (do not use steam from a boiler treated with amines for corrosion control); and capable
of reaching the desired temperature within 30 min. Confirm, by chemical or toxicity tests, that
the steam supply has not been treated with amines or other corrosion-control chemicals that will
impart toxicity.
Use of a vertical autoclave or pressure cooker is not recommended because of difficulty in
adjusting and maintaining sterilization temperature and the potential hazard. If a pressure cooker
is used in emergency or special circumstances, equip it with an efficient pressure gauge and a
thermometer the bulb of which is 2.5 cm above the water level.
4. Gas Sterilizers
Use a sterilizer equipped with automatic controls capable of carrying out a complete
sterilization cycle. As a sterilizing gas use ethylene oxide (CAUTION: Ethylene oxide is
toxic—avoid inhalation, ingestion, and contact with the skin. Also, ethylene oxide forms an
explosive mixture with air at 3-80% proportion.) diluted to 10 to 12% with an inert gas. Provide
an automatic control cycle to evacuate sterilizing chamber to at least 0.06 kPa, to hold the
vacuum for 30 min, to adjust humidity and temperature, to charge with the ethylene oxide
mixture to a pressure dependent on mixture used, to hold such pressure for at least 4 h, to vent
gas, to evacuate to 0.06 kPa, and finally, to bring to atmospheric pressure with sterile air. The
humidity, temperature, pressure, and time of sterilizing cycle depend on the gas mixture used.
Store overnight sample bottles with loosened caps that were sterilized by gas, to allow last
traces of gas mixture to dissipate. Incubate overnight media sterilized by gas, to insure
dissipation of gas.
In general, mixtures of ethylene oxide with chlorinated hydrocarbons such as freon are
harmful to plastics, although at temperatures below 55°C, gas pressure not over 35 kPa, and time
of sterilization less than 6 h, the effect is minimal. If carbon dioxide is used as a diluent of
ethylene oxide, increase exposure time and pressure, depending on temperature and humidity
that can be used.
Determine proper cycle and gas mixture for objects to be sterilized and confirm by sterility
tests.
5. Optical Counting Equipment
a. Pour and spread plates: Use Quebec-type colony counter, dark-field model preferred, or
one providing equivalent magnification (1.5 diameters) and satisfactory visibility.
b. Membrane filters: Use a binocular microscope with magnification of 10 to 15×. Provide
daylight fluorescent light source at angle of 60 to 80° above the colonies; use low-angle lighting
for nonpigmented colonies.
c. Mechanical tally.
6. pH Equipment
Use electrometric pH meters, accurate to at least 0.1 pH units, for determining pH values of
media.
7. Balances
Use balances providing a sensitivity of at least 0.1 g at a load of 150 g, with appropriate
weights. Use an analytical balance having a sensitivity of 1 mg under a load of 10 g for weighing
small quantities (less than 2 g) of materials. Single-pan rapid-weigh balances are most
convenient.
8. Media Preparation Utensils
Use borosilicate glass or other suitable noncorrosive equipment such as stainless steel. Use
glassware that is clean and free of residues, dried agar, or other foreign materials that may
contaminate media.
9. Pipets and Graduated Cylinders
Use pipets of any convenient size, provided that they deliver the required volume accurately
and quickly. The error of calibration for a given manufacturer’s lot must not exceed 2.5%. Use
pipets having graduations distinctly marked and with unbroken tips. Bacteriological transfer
pipets or pipets conforming to APHA standards may be used. Do not pipet by mouth; use a pipet
aid.
Use graduated cylinders meeting ASTM Standards (D-86 and D-216) and with accuracy
limits established by NIST where appropriate.
10. Pipet Containers
Use boxes of aluminum or stainless steel, end measurement 5 to 7.5 cm, cylindrical or
rectangular, and length about 40 cm. When these are not available, paper wrappings for
individual pipets may be substituted. To avoid excessive charring during sterilization, use
best-quality sulfate pulp (kraft) paper. Do not use copper or copper alloy cans or boxes as pipet
containers.
11. Refrigerator
Use a refrigerator maintaining a temperature of 1 to 4.4°C to store samples, media, reagents,
etc. Do not store volatile solvents, food, or beverages in a refrigerator with media. Frost-free
refrigerators may cause excessive media dehydration on storage longer than 1 week.
12. Temperature-Monitoring Devices
Use glass or metal thermometers graduated to 0.5°C to monitor most incubators and
refrigerators. Use thermometers graduated to 0.1°C for incubators operated above 40°C. Use
continuous recording devices that are equally sensitive. Verify accuracy by comparison with a
NIST-certified thermometer, or equivalent.
13. Dilution Bottles or Tubes
Use bottles or tubes of resistant glass, preferably borosilicate glass, closed with glass
stoppers or screw caps equipped with liners that do not produce toxic or bacteriostatic
compounds on sterilization. Do not use cotton plugs as closures. Mark graduation levels
indelibly on side of dilution bottle or tube. Plastic bottles of nontoxic material and acceptable
size may be substituted for glass provided that they can be sterilized properly.
14. Petri Dishes
For the plate count, use glass or plastic petri dishes about 100 × 15 mm. Use dishes the
bottoms of which are free from bubbles and scratches and flat so that the medium will be of
uniform thickness throughout the plate. For the membrane filter technique use loose-lid glass or
plastic dishes, 60 × 15 mm, or tight-lid dishes, 50 × 12 mm. Sterilize petri dishes and store in
metal cans (aluminum or stainless steel, but not copper), or wrap in paper—preferably
best-quality sulfate pulp (kraft)—before sterilizing. Presterilized petri dishes are commercially
available.
15. Membrane Filtration Equipment
Use filter funnel and membrane holder made of seamless stainless steel, glass, or
autoclavable plastic that does not leak and is not subject to corrosion. Field laboratory kits are
acceptable but standard laboratory filtration equipment and procedures are required.
16. Fermentation Tubes and Vials
Use fermentation tubes of any type, if their design permits conforming to medium and
volume requirements for concentration of nutritive ingredients as described subsequently. Where
tubes are used for a test of gas production, enclose a shell vial, inverted. Use tube and vial of
such size that the vial will be filled completely with medium, at least partly submerged in the
tube, and large enough to make gas bubbles easily visible.
17. Inoculating Equipment
Use wire loops made of 22- or 24-gauge nickel alloy*#(6) or platinum-iridium for flame
sterilization. Use loops at least 3 mm in diameter. Sterilize by dry heat or steam. Single-service
hardwood or plastic applicators also may be used. Make these 0.2 to 0.3 cm in diameter and at
least 2.5 cm longer than the fermentation tube; sterilize by dry heat and store in glass or other
nontoxic containers.
18. Sample Bottles
For bacteriological samples, use sterilizable bottles of glass or plastic of any suitable size
and shape. Use bottles capable of holding a sufficient volume of sample for all required tests and
an adequate air space, permitting proper washing, and maintaining samples uncontaminated until
examinations are completed. Ground-glass-stoppered bottles, preferably wide-mouthed and of
resistant glass, are recommended. Plastic bottles of suitable size, wide-mouthed, and made of
nontoxic materials such as polypropylene that can be sterilized repeatedly are satisfactory as
sample containers. Presterilized plastic bags, with or without dechlorinating agent, are available
commercially and may be used. Plastic containers eliminate the possibility of breakage during
shipment and reduce shipping weight.
Metal or plastic screw-cap closures with liners may be used on sample bottles provided that
no toxic compounds are produced on sterilization.
Before sterilization, cover tops and necks of sample bottles having glass closures with
aluminium foil or heavy kraft paper.
19. Bibliography
COLLINS, W.D. & H.B. RIFFENBURG. 1923. Contamination of water samples with material
dissolved from glass containers. Ind. Eng. Chem. 15:48.
CLARK, W.M. 1928. The Determination of Hydrogen Ion Concentration, 3rd ed. Williams &
Wilkins, Baltimore, Md.
ARCHAMBAULT, J., J. CUROT & M.H. MCCRADY. 1937. The need of uniformity of conditions for
counting plates (with suggestions for a standard colony counter). Amer. J. Pub. Health
27:809.
BARKWORTH, H. & J.O. IRWIN. 1941. The effect of the shape of the container and size of gas tube
in the presumptive coliform test. J. Hyg. 41:180.
RICHARDS, O.W. & P.C. HEIJN. 1945. An improved dark-field Quebec colony counter. J. Milk
Technol. 8:253.
COHEN, B. 1957. The measurement of pH, titratable acidity, and oxidation-reduction potentials.
In Manual of Microbiological Methods. Society of American Bacteriologists. McGraw-Hill
Book Co., New York, N.Y.
MORTON, H.E. 1957. Stainless-steel closures for replacement of cotton plugs in culture tubes.
Science. 126:1248.
MCGUIRE, O.E. 1964. Wood applicators for the confirmatory test in the bacteriological analysis
of water. Pub. Health Rep. 79:812.
BORDNER, R.H., J.A. WINTER & P.V. SCARPINO, eds. 1978. Microbiological Methods for
Monitoring the Environment, Water and Wastes. EPA-600/8-78-017, Environmental
Monitoring & Support Lab., U.S. Environmental Protection Agency, Cincinnati, Ohio.
AMERICAN PUBLIC HEALTH ASSOCIATION. 1993. Standard Methods for the Examination of
Dairy Products, 16th ed. American Public Health Assoc., Washington, D.C.
9040
WASHING AND STERILIZATION*#(7)
Cleanse all glassware thoroughly with a suitable detergent and hot water, rinse with hot
water to remove all traces of residual washing compound, and finally rinse with laboratory-pure
water. If mechanical glassware washers are used, equip them with influent plumbing of stainless
steel or other nontoxic material. Do not use copper piping to distribute water. Use stainless steel
or other nontoxic material for the rinse water system.
Sterilize glassware, except when in metal containers, for not less than 60 min at a
temperature of 170°C, unless it is known from recording thermometers that oven temperatures
are uniform, under which exceptional condition use 160°C. Heat glassware in metal containers to
170°C for not less than 2 h.
Sterilize sample bottles not made of plastic as above or in an autoclave at 121°C for 15 min.
For plastic bottles loosen caps before autoclaving to prevent distortion.
9050
PREPARATION OF CULTURE MEDIA*#(8)
9050 A.
General Procedures
1. Storage of Culture Media
Store dehydrated media (powders) in tightly closed bottles in the dark at less than 30°C in an
atmosphere of low humidity. Do not use them if they discolor or become caked and lose the
character of a free-flowing powder. Purchase dehydrated media in small quantities that will be
used within 6 months after opening. Additionally, use stocks of dehydrated media containing
selective agents such as sodium azide, bile salts or derivatives, antibiotics, sulfur-containing
amino acids, etc., of relatively current lot number (within a year of purchase) so as to maintain
optimum selectivity. See also Section 9020.
Prepare culture media in batches that will be used in less than 1 week. However, if the media
are contained in screw-capped tubes they may be stored for up to 3 months. See Table 9020:IV
for specific details. Store media out of direct sun and avoid contamination and excessive
evaporation.
Liquid media in fermentation tubes, if stored at refrigeration or even moderately low
temperatures, may dissolve sufficient air to produce, upon incubation at 35°C, a bubble of air in
the tube. Incubate fermentation tubes that have been stored at a low temperature overnight before
use and discard tubes containing air.
Fermentation tubes may be stored at approximately 25°C; but because evaporation may
proceed rapidly under these conditions—resulting in marked changes in concentration of the
ingredients—do not store at this temperature for more than 1 week. Discard tubes with an
evaporation loss exceeding 1 mL.
2. Adjustment of Reaction
State reaction of culture media in terms of hydrogen ion concentration, expressed as pH.
The decrease in pH during sterilization will vary slightly with the individual sterilizer in use,
and the initial reaction required to obtain the correct final reaction will have to be determined.
The decrease in pH usually will be 0.1 to 0.2 but occasionally may be as great as 0.3 in
double-strength media. When buffering salts such as phosphates are present in the media, the
decrease in pH value will be negligible.
Make tests to control adjustment to required pH with a pH meter. Measure pH of prepared
medium as directed in Section 4500-H+. Titrate a known volume of medium with a solution of
NaOH to the desired pH. Calculate amount of NaOH solution that must be added to the bulk
medium to reach this reaction. After adding and mixing thoroughly, check reaction and adjust if
necessary. The required final pH is given in the directions for preparing each medium. If a
specific pH is not prescribed, adjustment is unnecessary.
The pH of reconstituted dehydrated media seldom will require adjustment if made according
to directions. Such factors as errors in weighing dehydrated medium or overheating reconstituted
medium may produce an unacceptable final pH. Measure pH, especially of rehydrated selective
media, regularly to insure quality control and media specifications.
3. Sterilization
After rehydrating a medium, dispense promptly to culture vessels and sterilize within 2 h.
Do not store nonsterile media.
Sterilize all media, except sugar broths or broths with other specifications, in an autoclave at
121°C for 15 min after the temperature has reached 121°C. When the pressure reaches zero,
remove medium from autoclave and cool quickly to avoid decomposition of sugars by prolonged
exposure to heat. To permit uniform heating and rapid cooling, pack materials loosely and in
small containers. Sterilize sugar broths at 121°C for 12 to 15 min. The maximum elapsed time
for exposure of sugar broths to any heat (from time of closing loaded autoclave to unloading) is
45 min. Preferably use a double-walled autoclave to permit preheating before loading to reduce
total needed heating time to within the 45-min limit. Presterilized media may be available
commercially.
4. Bibliography
BUNKER, G.C. & H. SCHUBER. 1922. The reaction of culture media. J. Amer. Water Works Assoc.
9:63.
RICHARDSON, G.H., ed. 1985. Standard Methods for the Examination of Dairy Products, 15th ed.
American Public Health Assoc., Washington, D.C.
BALOWS, A., W.J. HAUSLER, JR., K.L. HERRMANN, H.D. ISENBERG & H.J. SHADOMY, eds. 1991.
Manual of Clinical Microbiology, 5th ed. American Soc. Microbiology, Washington, D.C.
9050 B.
Water
1. Specifications
To prepare culture media and reagents, use only distilled or demineralized reagent-grade
water that has been tested and found free from traces of dissolved metals and bactericidal or
inhibitory compounds. Toxicity in distilled water may be derived from fluoridated water high in
silica. Other sources of toxicity are silver, lead, and various unidentified organic complexes.
Where condensate return is used as feed for a still, toxic amines or other boiler compounds may
be present in distilled water. Residual chlorine or chloramines also may be found in distilled
water prepared from chlorinated water supplies. If chlorine compounds are found in distilled
water, neutralize them by adding an equivalent amount of sodium thiosulfate or sodium sulfite.
Distilled water also should be free of contaminating nutrients. Such contamination may be
derived from flashover of organics during distillation, continued use of exhausted carbon filter
beds, deionizing columns in need of recharging, solder flux residues in new piping, dust and
chemical fumes, and storage of water in unclean bottles. Store distilled water out of direct
sunlight to prevent growth of algae and turn supplies over as rapidly as possible. Aged distilled
water may contain toxic volatile organic compounds absorbed from the atmosphere if stored for
prolonged periods in unsealed containers. Good housekeeping practices usually will eliminate
nutrient contamination.
See Section 9020.
2. Bibliography
STRAKA, R.P. & J.L. STOKES. 1957. Rapid destruction of bacteria in commonly used diluents and
its elimination. Appl. Microbiol. 5:21.
GELDREICH, E.E. & H.F. CLARK. 1965. Distilled water suitability for microbiological applications.
J. Milk Food Technol. 28:351.
MACLEOD, R.A., S.C. KUO & R. GELINAS. 1967. Metabolic injury to bacteria. II. Metabolic injury
induced by distilled water or Cu++ in the plating diluent. J. Bacteriol. 93:961.
9050 C.
Media Specifications
The need for uniformity dictates the use of dehydrated media. Never prepare media from
basic ingredients when suitable dehydrated media are available. Follow manufacturer’s
directions for rehydration and sterilization. Commercially prepared media in liquid form (sterile
ampule or other) also may be used if known to give equivalent results. See Section 9020 for
quality-control specifications.
The terms used for protein source in most media, for example, peptone, tryptone, tryptose,
were coined by the developers of the media and may reflect commercial products rather than
clearly defined entities. It is not intended to preclude the use of alternative materials provided
that they produce equivalent results.
NOTE—The term ‘‘percent solution’’ as used in these directions is to be understood to mean
‘‘grams of solute per 100 mL solution.’’
1. Dilution Water
a. Buffered water: To prepare stock phosphate buffer solution, dissolve 34.0 g potassium
dihydrogen phosphate (KH2PO4), in 500 mL reagent-grade water, adjust to pH 7.2 ± 0.5 with 1N
sodium hydroxide (NaOH), and dilute to 1 L with reagent-grade water.
Add 1.25 mL stock phosphate buffer solution and 5.0 mL magnesium chloride solution (81.1
g MgCl2⋅6H2O/L reagent-grade water) to 1 L reagent-grade water. Dispense in amounts that will
provide 99 ± 2.0 mL or 9 ± 0.2 mL after autoclaving for 15 min.
b. Peptone water: Prepare a 10% solution of peptone in distilled water. Dilute a measured
volume to provide a final 0.1% solution. Final pH should be 6.8.
Dispense in amounts to provide 99 ± 2.0 mL or 9 ± 0.2 mL after autoclaving for 15 min.
Do not suspend bacteria in any dilution water for more than 30 min at room temperature
because death or multiplication may occur.
2. Culture Media
Specifications for individual media are included in subsequent sections. Details are provided
where use of a medium first is described.
9060
9060 A.
SAMPLES*#(9)
Collection
1. Containers
Collect samples for microbiological examination in nonreactive borosilicate glass or plastic
bottles that have been cleansed and rinsed carefully, given a final rinse with deionized or
distilled water, and sterilized as directed in Section 9030 and Section 9040. For some
applications samples may be collected in presterilized plastic bags.
2. Dechlorination
Add a reducing agent to containers intended for the collection of water having residual
chlorine or other halogen unless they contain broth for direct planting of sample. Sodium
thiosulfate (Na2S2O3) is a satisfactory dechlorinating agent that neutralizes any residual halogen
and prevents continuation of bactericidal action during sample transit. The examination then will
indicate more accurately the true microbial content of the water at the time of sampling.
For sampling chlorinated wastewater effluents add sufficient Na2S2O3 to a clean sterile
sample bottle to give a concentration of 100 mg/L in the sample. In a 120-mL bottle 0.1 mL of a
10% solution of Na2S2O3 will neutralize a sample containing about 15 mg/L residual chlorine.
For drinking water samples, the concentration of dechlorinating agent may be reduced: 0.1 mL
of a 3% solution of Na2S2O3 in a 120-mL bottle will neutralize up to 5 mg/L residual chlorine.
Cap bottle and sterilize by either dry or moist heat, as directed (Section 9040). Presterilized
plastic bags or bottles containing Na2S2O3 are available commercially.
Collect water samples high in metals, including copper or zinc (>1.0 mg/L), and wastewater
samples high in heavy metals in sample bottles containing a chelating agent that will reduce
metal toxicity. This is particularly significant when such samples are in transit for 4 h or more.
Use 372 mg/L of the disodium salt of ethylenediaminetetraacetic acid (EDTA). Adjust EDTA
solution to pH 6.5 before use. Add EDTA separately to sample bottle before bottle sterilization
(0.3 mL 15% solution in a 120-mL bottle) or combine it with the Na2S2O3 solution before
addition.
3. Sampling Procedures
When the sample is collected, leave ample air space in the bottle (at least 2.5 cm) to
facilitate mixing by shaking, before examination. Collect samples that are representative of the
water being tested, flush or disinfect sample ports, and use aseptic techniques to avoid sample
contamination.
Keep sampling bottle closed until it is to be filled. Remove stopper and cap as a unit; do not
contaminate inner surface of stopper or cap and neck of bottle. Fill container without rinsing,
replace stopper or cap immediately, and if used, secure hood around neck of bottle.
a. Potable water: If the water sample is to be taken from a distribution-system tap without
attachments, select a tap that is supplying water from a service pipe directly connected with the
main, and is not, for example, served from a cistern or storage tank. Open tap fully and let water
run to waste for 2 or 3 min, or for a time sufficient to permit clearing the service line. Reduce
water flow to permit filling bottle without splashing. If tap cleanliness is questionable, choose
another tap. If a questionable tap is required for special sampling purposes, disinfect the faucet
(inside and outside) by applying a solution of sodium hypochlorite (100 mg NaOCl/L) to faucet
before sampling; let water run for additional 2 to 3 min after treatment. Do not sample from
leaking taps that allow water to flow over the outside of the tap. In sampling from a mixing
faucet remove faucet attachments such as screen or splash guard, run hot water for 2 min, then
cold water for 2 to 3 min, and collect sample as indicated above.
If the sample is to be taken from a well fitted with a hand pump, pump water to waste for
about 5 to 10 min or until water temperature has stabilized before collecting sample. If an
outdoor sampling location must be used, avoid collecting samples from frost-proof hydrants. If
there is no pumping machinery, collect a sample directly from the well by means of a sterilized
bottle fitted with a weight at the base; take care to avoid contaminating samples by any surface
scum. Other sterile sampling devices, such as a trip bailer, also may be used.
In drinking water evaluation, collect samples of finished water from distribution sites
selected to assure systematic coverage during each month. Carefully choose distribution system
sample locations to include dead-end sections to demonstrate bacteriological quality throughout
the network and to ensure that localized contamination does not occur through
cross-connections, breaks in the distribution lines, or reduction in positive pressure. Sample
locations may be public sites (police and fire stations, government office buildings, schools, bus
and train stations, airports, community parks), commercial establishments (restaurants, gas
stations, office buildings, industrial plants), private residences (single residences, apartment
buildings, and townhouse complexes), and special sampling stations built into the distribution
network. Preferably avoid outdoor taps, fire hydrants, water treatment units, and backflow
prevention devices. Establish sampling program in consultation with state and local health
authorities.
b. Raw water supply: In collecting samples directly from a river, stream, lake, reservoir,
spring, or shallow well, obtain samples representative of the water that is the source of supply to
consumers. It is undesirable to take samples too near the bank or too far from the point of
drawoff, or at a depth above or below the point of drawoff.
c. Surface waters: Stream studies may be short-term, high-intensity efforts. Select
bacteriological sampling locations to include a baseline location upstream from the study area,
industrial and municipal waste outfalls into the main stream study area, tributaries except those
with a flow less than 10% of the main stream, intake points for municipal or industrial water
facilities, downstream samples based on stream flow time, and downstream recreational areas.
Dispersion of wastewaters into the receiving stream may necessitate preliminary cross-section
studies to determine completeness of mixing. Where a tributary stream is involved, select the
sampling point near the confluence with the main stream. Samples may be collected from a boat
or from bridges near critical study points. Choose sampling frequency to be reflective of
changing stream or water body conditions. For example, to evaluate waste discharges, sample
every 4 to 6 h and advance the time over a 7- to 10-d period.
To monitor stream and lake water quality establish sampling locations at critical sites.
Sampling frequency may be seasonal for recreational waters, daily for water supply intakes,
hourly where waste treatment control is erratic and effluents are discharged into shellfish
harvesting areas, or even continuous.
d. Bathing beaches: Sampling locations for recreational areas should reflect water quality
within the entire recreational zone. Include sites from upstream peripheral areas and locations
adjacent to drains or natural contours that would discharge stormwater collections or septic
wastes. Collect samples in the swimming area from a uniform depth of approximately 1 m.
Consider sediment sampling of the water-beach (soil) interface because of exposure of young
children at the water’s edge.
To obtain baseline data on marine and estuarine bathing water quality include sampling at
low, high, and ebb tides.
Relate sampling frequency directly to the peak bathing period, which generally occurs in the
afternoon. Preferably, collect daily samples during the recognized bathing season; as a minimum
include Friday, Saturday, Sunday, and holidays. When limiting sampling to days of peak
recreational use, preferably collect a sample in the morning and the afternoon. Correlate
bacteriological data with turbidity levels or rainfall over the watershed to make rapid assessment
of water quality changes.
e. Sediments and biosolids: The bacteriology of bottom sediments is important in water
supply reservoirs, in lakes, rivers, and coastal waters used for recreational purposes, and in
shellfish-growing waters. Sediments may provide a stable index of the general quality of the
overlying water, particularly where there is great variability in its bacteriological quality.
Sampling frequency in reservoirs and lakes may be related more to seasonal changes in
water temperatures and stormwater runoff. Bottom sediment changes in river and estuarine
waters may be more erratic, being influenced by stormwater runoff, increased flow velocities,
and sudden changes in the quality of effluent discharges.
Microbiological examination of biosolids from water and wastewater treatment processes is
desirable to determine the impact of their disposal into receiving waters, ocean dumping, land
application, or burial in landfill operations.
Collect and handle biosolids with less than 7% total solids using the procedures discussed for
other water samples. Biosolids with more than 7% solids and exhibiting a ‘‘plastic’’ consistency
or ‘‘semisolid’’ state typical of thickened sludges require a finite shear stress to cause them to
flow. This resistance to flow results in heterogeneous distribution of biosolids in tanks and
lagoons. Use cross-section sampling of accumulated biosolids to determine distribution of
organisms within these impoundments. Establish a length-width grid across the top of the
impoundment, and sample at intercepts. A thief sampler that samples only the solids layer may
be useful. Alternatively use weighted bottle samplers that can be opened up at a desired depth to
collect samples at specific locations.
Processed biosolids having no free liquids are best sampled when they are being transferred.
Collect grab samples across the entire width of the conveyor and combine into a composite
sample. If solids are stored in piles, classification occurs. Exteriors of uncovered piles are subject
to various environmental stresses such as precipitation, wind, fugitive dusts, and fecal
contamination from scavengers. Consequently, surface samples may not reflect the
microbiological quality of the pile. Therefore, use cross-section sampling of these piles to
determine the degree of heterogeneity within the pile. Establish a length-width grid across the
top of the pile, and sample intercepts. Sample augers and corers may prove to be ineffective for
sampling piles of variable composition. In such cases use hand shovels to remove overburden.
f. Nonpotable samples (manual sampling): Take samples from a river, stream, lake, or
reservoir by holding the bottle near its base in the hand and plunging it, neck downward, below
the surface. Turn bottle until neck points slightly upward and mouth is directed toward the
current. If there is no current, as in the case of a reservoir, create a current artificially by pushing
bottle forward horizontally in a direction away from the hand. When sampling from a boat,
obtain samples from upstream side of boat. If it is not possible to collect samples from these
situations in this way, attach a weight to base of bottle and lower it into the water. In any case,
take care to avoid contact with bank or stream bed; otherwise, water fouling may occur.
g. Sampling apparatus: Special apparatus that permits mechanical removal of bottle stopper
below water surface is required to collect samples from depths of a lake or reservoir. Various
types of deep sampling devices are available. The most common is the ZoBell J-Z sampler,1
which uses a sterile 350-mL bottle and a rubber stopper through which a piece of glass tubing
has been passed. This tubing is connected to another piece of glass tubing by a rubber connecting
hose. The unit is mounted on a metal frame containing a cable and a messenger. When the
messenger is released, it strikes the glass tubing at a point that has been slightly weakened by a
file mark. The glass tube is broken by the messenger and the tension set up by the rubber
connecting hose is released and the tubing swings to the side. Water is sucked into the bottle as a
consequence of the partial vacuum created by sealing the unit at time of autoclaving.
Commercial adaptations of this sampler and others are available.
Bottom sediment sampling also requires special apparatus. The sampler described by Van
Donsel and Geldreich2 has been found effective for a variety of bottom materials for remote
(deep water) or hand (shallow water) sampling. Construct this sampler preferably of stainless
steel and fit with a sterile plastic bag. A nylon cord closes the bag after the sampler penetrates
the sediment. A slide bar keeps the bag closed during descent and is opened, thereby opening the
bag, during sediment sampling.
For sampling wastewaters or effluents the techniques described above generally are
adequate; in addition see Section 1060.
4. Size of Sample
The volume of sample should be sufficient to carry out all tests required, preferably not less
than 100 mL.
5. Identifying Data
Accompany samples by complete and accurate identifying and descriptive data. Do not
accept for examination inadequately identified samples.
6. References
1. ZOBELL, C.E. 1941. Apparatus for collecting water samples from different depths for
bacteriological analysis. J. Mar. Res. 4:173.
2. VAN DONSEL, D.J. & E.E. GELDREICH. 1971. Relationships of Salmonellae to fecal
coliforms in bottom sediments. Water Res. 5:1079.
7. Bibliography
PUBLIC HEALTH LABORATORY SERVICE WATER SUB-COMMITTEE. 1953. The effect of sodium
thiosulphate on the coliform and Bacterium coli counts of non-chlorinated water samples. J.
Hyg. 51:572.
SHIPE, E.L. & A. FIELDS. 1956. Chelation as a method for maintaining the coliform index in water
samples. Pub. Health Rep. 71:974.
HOATHER, R.C. 1961. The bacteriological examination of water. J. Inst. Water Eng. 61:426.
COLES, H.G. 1964. Ethylenediamine tetra-acetic acid and sodium thiosulphate as protective
agents for coliform organisms in water samples stored for one day at atmospheric
temperature. Proc. Soc. Water Treat. Exam. 13:350.
DAHLING, D.R. & B.A. WRIGHT. 1984. Processing and transport of environmental virus samples.
Appl. Environ. Microbiol. 47:1272.
U.S. ENVIRONMENTAL PROTECTION AGENCY. 1992. Environmental Regulations and Technology
Control of Pathogens and Vector Attraction in Sewage Sludge. EPA-625/R-92-013.
Washington, D.C.
9060 B.
Preservation and Storage
1. Holding Time and Temperature
a. General: Start microbiological analysis of water samples as soon as possible after
collection to avoid unpredictable changes in the microbial population. For most accurate results,
ice samples during transport to the laboratory, if they cannot be processed within 1 h after
collection. If the results may be used in legal action, employ special means (rapid transport,
express mail, courier service, etc.) to deliver the samples to the laboratory within the specified
time limits and maintain chain of custody. Follow the guidelines and requirements given below
for specific water types.
b. Drinking water for compliance purposes: Preferably hold samples at <10°C during transit
to the laboratory. Analyze samples on day of receipt whenever possible and refrigerate overnight
if arrival is too late for processing on same day. Do not exceed 30 h holding time from collection
to analysis for coliform bacteria. Do not exceed 8 h holding time for heterotrophic plate counts.
c. Nonpotable water for compliance purposes: Hold source water, stream pollution,
recreational water, and wastewater samples below 10°C during a maximum transport time of 6 h.
Refrigerate these samples upon receipt in the laboratory and process within 2 h. When transport
conditions necessitate delays in delivery of samples longer than 6 h, consider using either field
laboratory facilities located at the site of collection or delayed incubation procedures.
d. Other water types for noncompliance purposes: Hold samples below 10°C during
transport and until time of analysis. Do not exceed 24 h holding time.
2. Bibliography
CALDWELL, E.L. & L.W. PARR. 1933. Present status of handling water samples—Comparison of
bacteriological analyses under varying temperatures and holding conditions, with special
reference to the direct method. Amer. J. Pub. Health 23:467.
COX, K.E. & F.B. CLAIBORNE. 1949. Effect of age and storage temperature on bacteriological
water samples. J. Amer. Water Works Assoc. 41: 948.
PUBLIC HEALTH LABORATORY SERVICE WATER SUB-COMMITTEE. 1952. The effect of storage
on the coliform and Bacterium coli counts of water samples. Overnight storage at room and
refrigerator temperatures. J. Hyg. 50:107.
PUBLIC HEALTH LABORATORY SERVICE WATER SUB-COMMITTEE. 1953. The effect of storage
on the coliform and Bacterium coli counts of water samples. Storage for six hours at room
and refrigerator temperatures. J. Hyg. 51:559.
MCCARTHY, J.A. 1957. Storage of water sample for bacteriological examinations. Amer. J. Pub.
Health 47:971.
LONSANE, B.K., N.M. PARHAD & N.U. RAO. 1967. Effect of storage temperature and time on the
coliform in water samples. Water Res. 1: 309.
LUCKING, H.E. 1967. Death rate of coliform bacteria in stored Montana water samples. J.
Environ. Health 29:576.
MCDANIELS, A.E. & R.H. BORDNER. 1983. Effect of holding time and temperature on coliform
numbers in drinking water. J. Amer. Water Works Assoc. 75:458.
MCDANIELS, A.E. et al. 1985. Holding effects on coliform enumeration in drinking water
samples. Appl. Environ. Microbiol. 50:755.
9211
RAPID DETECTION METHODS*#(10)
9211 A.
Introduction
There is a generally recognized need for methods that permit rapid estimation of the
bacteriological quality of water. Applications of rapid methods may range from analysis of
wastewater to potable water quality assessment. In the latter case, during emergencies involving
water treatment plant failure, line breaks in a distribution network, or other disruptions to water
supply caused by disasters, there is urgent need for rapid assessment of the sanitary quality of
water.
Ideally, rapid procedures would be reliable and have sensitivity levels equal to those of the
standard tests routinely used. However, sensitivity of a rapid test may be compromised because
the bacterial limit sought may be below the minimum bacterial concentration essential to rapid
detection. Rapid tests fall into two categories, those involving modified conventional procedures
and those requiring special instrumentation and materials.
9211 B.
Seven-Hour Fecal Coliform Test (SPECIALIZED)
This method1,2 is similar to the fecal coliform membrane filter procedure (see Section
9222D) but uses a different medium and incubation temperature to yield results in 7 h that
generally are comparable to those obtained by the standard fecal coliform method.
1. Medium
M-7 h FC agar: This medium may not be available in dehydrated form and may require
preparation from the basic ingredients.
Proteose peptone No. 3 or polypeptone
Yeast extract
Lactose
d-Mannitol
Sodium chloride, NaCl
Sodium lauryl sulfate
Sodium desoxycholate
Bromcresol purple
Phenol red
Agar
Reagent-grade water
5.0
3.0
10.0
5.0
7.5
0.2
0.1
0.35
0.3
15.0
1
g
g
g
g
g
g
g
g
g
g
L
Heat in boiling water bath. After ingredients are dissolved heat additional 5 min. Cool to 55
to 60°C and adjust pH to 7.3 ± 0.1 with 0.1N NaOH (0.35 mL/L usually required). Cool to about
45°C and dispense in 4- to 5-mL quantities to petri plates with tight-fitting covers. Store at 2 to
10°C. Discard after 30 d.
2. Procedure
Filter an appropriate sample volume through a membrane filter, place filter on the surface of
a plate containing M-7 h FC agar medium, and incubate at 41.5°C for 7 h. Fecal coliform
colonies are yellow (indicative of lactose fermentation).
3. References
1. VAN DONSEL, D.J., R.M. TWEDT & E.E. GELDREICH. 1969. Optimum temperature for
quantitation of fecal coliforms in seven hours on the membrane filter. Bacteriol. Proc.
Abs. No. G46, p. 25.
2. REASONER, D.J., J.C. BLANNON & E.E. GELDREICH. 1979. Rapid seven hour fecal
coliform test. Appl. Environ. Microbiol. 38:229.
9211 C.
Special Techniques (SPECIALIZED)
Special rapid techniques are summarized in Table 9211:I. Most are not sensitive enough for
potable water quality measurement or are not specific. They may be useful in monitoring
wastewater effluents and natural waters but require reagents not generally available, are tedious,
or require special handling or incubation schemes incompatible with most water laboratory
schedules. Except for the colorimetric test, none are suitable for routine use but they may be used
as research tools. The user should refer to the literature citations for the technique listed in the
table for procedural details, conditions for use, and method limitations. Only the adenosine
triphosphate (ATP) procedure (the firefly bioluminescence system), the colorimetric test to
estimate total microbial density, and a radiometric fecal coliform procedure that uses a
14C-labeled substrate can be recommended.
Correlate initial concentration of bacteria with ATP concentration by extracting ATP from
serial dilutions of a bacterial suspension, or for the 14C radiometric method, standardize by
determining the 14CO2 released by known concentrations of fecal coliform organisms in natural
samples, not pure cultures. In using any rapid procedure, determine the initial bacterial density
by using an appropriate procedure such as heterotrophic plate count (Section 9215) or total
(Section 9221) or fecal (Section 9222) coliforms, and correlate with results from the special
rapid technique.
1. Bioluminescence Test (Total Viable Microbial Measurement)
The firefly luciferase test for ATP in living cells is based on the reaction between the
luciferase enzyme, luciferin (enzyme substrate), magnesium ions, and ATP. Light is emitted
during the reaction and can be measured quantitatively and correlated with the quantity of ATP
extracted from known numbers of bacteria. When all reactants except ATP are in excess, ATP is
the limiting factor. Addition of ATP drives the reactions, producing a pulse of light that is
proportional to the ATP concentration.
The assay is completed in less than 1 h.1-3 For monitoring microbial populations in water,
the ATP assay is limited primarily by the need to concentrate bacteria from the sample to
achieve the minimum ATP sensitivity level, which is 105 cells/mL. When combined with
membrane filtration of a 1-L sample, ATP assay can provide the sensitivity level needed.
2. Radiometric Detection (Fecal Coliforms)
In this test, 14CO2 is released from a 14C-labeled substrate.14 The technique permits
presumptive detection of as few as 2 to 20 fecal coliform bacteria in 4.5 h. The test uses M-FC
broth, uniformly labeled 14C-mannitol, and two-temperature incubation; 2 h at 35°C followed by
2.5 h at 44.5°C for fecal coliform specificity. Add labeled substrate at start of 44.5°C incubation.
Use membrane filtration to concentrate organisms from sample and place membrane filter in
M-FC broth in a sealable container. The 14CO2 released is trapped by exposure to
Ba(OH)2-saturated filter paper disk. 14C activity is assayed by liquid scintillation spectrometry.
Except for the use of the 14C-mannitol substrate and liquid scintillation spectrometry to count the
activity of the 14CO2 released by the fecal coliforms, this procedure is similar to those given in
Section 9222.
3. References
1. CHAPPELLE, E.W. & G.L. PICCIOLO. 1975. Laboratory Procedures Manual for the Firefly
Luciferase Assay for Adenosine Triphosphate (ATP). NASA GSFC Doc. X-726-75-1,
National Aeronautics & Space Admin., Washington, D.C.
2. PICCIOLO, G.L., E.W. CHAPPELLE, J.W. DEMING, R.R. THOMAS, D.A. NIBLE & H. OKREND.
1981. Firefly luciferase ATP assay development for monitoring bacterial concentration
in water supplies. EPA-600/S2:81-014, U.S. Environmental Protection Agency,
Cincinnati, Ohio; NTIS No. PB 88-103809/AS, National Technical Information Serv.,
Springfield, Va.
3. NELSON, W.H., ed. 1985. Instrumental Methods for Rapid Microbiological Analysis.
VCH Publishers, Inc., Deerfield Beach, Fla.
4. SEITZ, W.R. & M.P. NEARY. 1974. Chemiluminescence and bioluminescence. Anal.
Chem. 46:188A.
5. OLENIAZ, W.S., M.A. PISANO, M.H. ROSENFELD & R.L. ELGART. 1968. Chemiluminescent
method for detecting microorganisms in water. Environ. Sci. Technol. 2:1030.
6. WHEELER, T.G. & M.C. GOLDSCHMIDT. 1975. Determination of bacterial cell
concentrations by electrical measurements. J. Clin. Microbiol. 1:25.
7. SILVERMAN, M.P. & E.F. MUNOZ. 1979. Automated electrical impedance technique for
rapid enumeration of fecal coliforms in effluents from sewage treatment plants. Appl.
Environ. Microbiol. 37:521.
8. MUNOZ, E.F. & M.P. SILVERMAN. 1979. Rapid, single-step most-probable-number
method for enumerating fecal coliforms in effluents from sewage treatment plants.
9. FIRSTENBERG-EDEN, R. & G. EDEN. 1984. Impedance Microbiology. John Wiley &
Sons, Inc., New York, N.Y.
10. WALLIS, C. & J.L. MELNICK. 1985. An instrument for the immediate quantification of
bacteria in potable waters. Appl. Environ. Microbiol. 49:1251.
11. BITTON, G., R.J. DUTTON & J.A. FORAN. 1984. A new rapid technique for counting
microorganisms directly on membrane filters. Stain Technol. 58:343.
12. SIERACKI, M.E., P.W. JOHNSON & J.M. SIEBURTH. 1985. Detection, enumeration, and
sizing of planktonic bacteria by image-analyzed epifluoresence microscopy. Appl.
13. MCCOY, W.F. & B.H. OLSON. 1985. Fluorometric determination of the DNA
concentration in municipal drinking water. Appl. Environ. Microbiol. 49:811.
14. REASONER, D.J. & E.E. GELDREICH. 1978. Rapid detection of water-borne fecal
coliforms by 14CO2 release. In A.N. Sharpe & D.S. Clark, eds. Mechanizing
Microbiology. Charles C. Thomas, Publisher, Springfield, Ill.
15. MORAN, J.W. & L.D. WITTER. 1976. An automated rapid test for Escherichia coli in milk.
J. Food Sci. 41:165.
16. MORAN, J.W. & L.D. WITTER. 1976. An automated rapid method for measuring fecal
pollution. Water Sewage Works 123:66.
17. TRINEL, P.A., N. HANOUNE & H. LECLERC. 1980. Automation of water bacteriological
analysis: running test of an experimental prototype. Appl. Environ. Microbiol. 39:976.
18. WILKINS, J.R., G.E. STONER & E.H. BOYKIN. 1974. Microbial detection method based on
sensing molecular hydrogen. Appl. Microbiol. 27:947.
19. WILKINS, J.R. & E.H. BOYKIN. 1976. Analytical notes—electrochemical method for early
detection of monitoring of coliforms. J. Amer. Water Works Assoc. 68:257.
20. GRANA, D.C. & J.R. WILKINS. 1979. Description and field test results of an in situ
coliform monitoring system. NASA Tech. Paper 1334, National Aeronautics & Space
Admin., Washington, D.C.
21. NEWMAN, J.S. & R.T. O’BRIEN. 1975. Gas chromatographic presumptive test for
coliform bacteria in water. Appl. Environ. Microbiol. 30:584.
22. WARREN, L.S., R.E. BENOIT & J.A. JESSEE. 1978. Rapid enumeration of faecal coliforms
in water by a colorimetric β-galactosidase assay. Appl. Environ. Microbiol. 35:136.
23. JOUENNE, T., G.-A. JUNTER & G. CARRIERE. 1985. Selective detection and enumeration
of fecal coliforms in water by potentiometric measurement of lipoic acid reduction.
24. TENCATE, J.W., H.R. BULER, A. STURK & J. LEVIN. 1985. Bacterial Endotoxins. Structure,
Biomedical Significance, and Detection with the Limulus Amebocyte Lysate Test.
Alan R. Liss, Inc., New York, N.Y.
25. JORGENSEN, J.H., J.C. LEE, G.A. ALEXANDER & H.W. WOLF. 1979. Comparison of
Limulus assay, standard plate count, and total coliform count for microbiological
assessment of renovated wastewater. Appl. Environ. Microbiol. 37:928.
26. JORGENSEN, J.H. & G.A. ALEXANDER. 1981. Automation of the Limulus amebocyte
lysate test by using the Abbott MS-2 microbiology system. Appl. Environ. Microbiol.
41:1316.
27. TSUGI, K., P.A. MARTIN & D.M. BUSSEY. 1984. Automation of chromogenic substrate
Limulus amebocyte lysate assay method for endotoxin by robotic system. Appl.
28. ABSHIRE, R.L. 1976. Detection of enteropathogenic Escherichia coli strains in
wastewater by fluorescent antibody. Can. J. Microbiol. 22:365.
29. ABSHIRE, R.L. & R.K. GUTHRIE. 1973. Fluorescent antibody techniques as a method for
the detection of fecal pollution. Can. J. Microbiol. 19:201.
30. THOMASON, B.M. 1981. Current status of immunofluorescent methodology. J. Food
Protect. 44:381.
9211 D.
Coliphage Detection
Coliphages are bacteriophages that infect and replicate in coliform bacteria and appear to be
present wherever total and fecal coliforms are found. Correlations between coliphages and
coliform bacteria in fresh water generally show that coliphages may be used to indicate the
sanitary quality of water.1-5 Because coliphages are more resistant to chlorine disinfection than
total or fecal coliforms, they may be a better indicator of disinfection efficiency than coliform
bacteria.4 The quantitative relationship between coliphages and coliform bacteria in disinfected
waters is different from that in natural fresh waters because of differences in their survival rates.
1. Materials and Culture Media
a. Host culture: Escherichia coli C, ATCC No. 13706.
b. Media:
1) Tryptic(ase) soy agar (TSA), to maintain E. coli C host stock cultures:
Tryptone (pancreatic digest of casein) or equivalent 15.0
Soytone (soybean peptone) or equivalent
5.0
5.0
Agar
15.0
Reagent-grade water
1
g
g
g
g
L
pH should be 7.3 ± 0.1 at 25°C; if necessary, adjust pH with 0.1 or 1.0N NaOH or HCl. Heat
to boiling to dissolve, then autoclave for 15 min at 121°C. For agar slants, dispense 5 to 8 mL in
16- × 125-mm screw-capped tubes before sterilizing; for plates, dispense 20 to 25 mL per petri
dish after autoclaving and cooling to about 45°C.
2) Tryptic(ase) soy broth (TSB):
Tryptone (pancreatic digest of casein), or equivalent
Soytone (soybean peptone), or equivalent
Dextrose
Dipotassium hydrogen phosphate, K2HPO4
Reagent-grade water
17.0
3.0
2.5
5.0
2.5
g
g
g
g
g
1
L
pH should be 7.3 ± 0.1 at 25°C; adjust with 0.1 or 1.0N NaOH or HCl, if necessary. Warm
and agitate to dissolve completely. Dispense in appropriate volumes as needed; sterilize in
autoclave for 15 min at 121°C.
3) Modified tryptic(ase) soy agar (MTSA): To the ingredients of TSB, add ammonium
nitrate, NH4NO3, 1.60 g; strontium nitrate, Sr(NO3)2, 0.21 g; and agar, 15 g. pH should be 7.3 ±
0.1 at 25°C; if necessary, adjust pH with 0.1 or 1.0N NaOH or HCl. Heat to boiling to dissolve,
dispense 5.5 mL in 16- × 25-mm screw-capped tubes, and autoclave for 15 min at 121°C.
4) Glycerine: Add 10% (w/v) to tryptic(ase) soy broth before autoclave sterilization.
5) 2,3,5-triphenyl tetrazolium chloride (TPTZ), 1% (w/v) in
ethanol. Add to MTSA tempered at 45 to 46°C to enhance plaque visibility. Prepare fresh
weekly.
2. Procedure
a. Frozen host preparation: Inoculate E. coli C from a stock agar slant (on TSA) into a
tube(s) containing 10 mL TSB and 10% glycerine (w/v) and incubate overnight at 35°C. Then
inoculate each tube into a flask containing 25 mL TSB plus 10% glycerine and incubate at 35 ±
0.5°C until an optical density of 0.5 at 520 nm is obtained (equivalent to about 1 × 109 E. coli C
cells/mL). Measure optical density using a spectrometer. Zero spectrometer with sterile TSB plus
10% glycerine.
Aseptically dispense 4.5-mL portions of cell suspension in sterile plastic test tubes, cap, chill
to 9°C, and freeze at −20°C. Store for no longer than 6 weeks in non-frost-free freezer to reduce
loss of frozen host culture viability.
b. Assay procedure: The procedure is directly applicable to samples containing more than 5
coliphage/100 mL; if sample contains more than 1000 coliphage/100 mL, dilute sample 1:5 or 1:
10 with sterile distilled water before proceeding.
Thaw tube(s) of frozen host E. coli C in 44.5°C water bath. Use one tube of host culture per
sample. Add 1.0 mL of host E. coli C culture, 5 mL sample or dilution, and 0.08 mL TPTZ6 to
each of four tubes of MTSA (melted and held at about 45°C).
Mix thoroughly and pour into separate 100- × 15-mm labeled petri dishes, cover, and let agar
gel. Incubate inverted plates at 35°C. Count plaques after incubating for 4 to 6 h.
3. Interpreting and Reporting Results
Bacteriophage infect and multiply in sensitive bacteria. This results in lysis of the bacterial
cells and a release of phage particles to infect adjacent cells. As the infected coliform bacteria
are lysed, visible clear areas known as plaques develop in the lawn of confluent bacterial growth.
Count plaques on each plate and record. Obtain the number of plaques/100 mL of sample by
summing the plaques on the four plates and multiplying by 5. If a diluted sample has been used,
additionally multiply by the reciprocal of the dilution factor.
Based on coliphage counts, estimate total and fecal coliform numbers as shown below.4
Independently verify equations for specific types of samples and locations.
Total coliforms:
log y = 0.627 (log x) + 1.864
where:
y = total coliforms/100 mL and
x = coliphages/100 mL.
Fecal coliforms:
log y = 0.805 (log x) + 0.895
where:
y = fecal coliforms/100 mL and
x = coliphages/100 mL.
4. References
1. WENTZEL, R.S., P.E. O’NEILL & J.F. KITCHENS. 1982. Evaluation of coliphage detection
as a rapid indicator of water quality. Appl. Environ. Microbiol. 43:430.
2. ISBISTER, J.D. & J.L. ALM. 1982. Rapid coliphage procedure for water treatment
processes. In Proc. Amer. Water Works Assoc. Water Quality Technol. Conf., Seattle,
Wash., Dec. 6–9, 1981.
3. ISBISTER, J.D., J.A. SIMMONS, W.M. SCOTT & J.F. KITCHENS. 1983. A simplified method
for coliphage detection in natural waters. Acta Microbiol. Polonica 32:197.
4. KOTT, Y., N. ROZE, S. SPERBER & N. BETZER. 1974. Bacteriophages as viral pollution
indicators. Water Res. 8:165.
5. KENNEDY, J.D., JR., G. BITTON & J.L. OBLINGER. 1985. Comparison of selective media
for assay of coliphages in sewage effluent and lake water. Appl. Environ. Microbiol.
49:33.
6. HURST, C.J., J.C. BLANNON, R.L. HARDAWAY & W.C. JACKSON. 1994. Differential effect
of tetrazolium dyes upon bacteriophage plaque assay titers. Appl. Environ. Microbiol.
60:3462.
9212
STRESSED ORGANISMS*#(11)
9212 A.
Introduction
1. General Discussion
Indicator bacteria, including total coliforms, fecal coliforms, and fecal streptococci, may
become stressed or injured in waters and wastewaters. These injured bacteria are incapable of
growth and colony formation under standard conditions because of structural or metabolic
damage. As a result, a substantial portion of the indicator bacteria present, i.e., 10 to greater than
90%, may not be detected.1,2 These false negative bacteriological findings could result in an
inaccurate definition of water quality and lead to the acceptance of a potentially hazardous
condition resulting from contamination by resistant pathogens3 or the penetration of undetected
indicator bacteria through treatment barriers.4
Stressed organisms are present under ordinary circumstances in treated drinking water and
wastewater effluents, saline waters, polluted natural waters, and relatively clean surface waters.
High numbers of injured indicator bacteria may be associated with partial or inadequate
disinfection and the presence of metal ions or other toxic substances. These and other factors,
including extremes of temperature and pH and solar radiation, may lead collectively to
significant underestimations of the number of viable indicator bacteria.
Publications support the health significance of injured coliform bacteria.2,5-7 These reports
show that enteropathogenic bacteria are less susceptible than coliforms to injury under
conditions similar to those in treated drinking water and wastewater, that injured pathogens
retain the potential for virulence, and that they recover after being ingested. Hence, methods
allowing for the enumeration of injured coliform bacteria yield more sensitive determinations of
potential health risks. This conclusion is further supported by the observation that viruses and
waterborne pathogens that form cysts also are more resistant than indicator bacteria to
environmental stressors.
2. Sample Handling and Collection
Certain laboratory manipulations following sample collection also may produce injury or act
as a secondary stress to the organisms.2,8 These include excessive sample storage time,
prolonged holding time (more than 30 min) of diluted samples before inoculation into growth
media and of inoculated samples before incubation at the proper temperature, incorrect media
formulations, incomplete mixing of sample with concentrated medium, and exposure to
untempered liquefied agar media. Excessive numbers of nonindicator bacteria also interfere with
detection of indicators by causing injury.9
3. References
1. MCFETERS, G.A., J.S. KIPPIN & M.W. LECHEVALLIER. 1986. Injured coliforms in drinking
water. Appl. Environ. Microbiol. 51:1.
2. MCFETERS, G.A. 1990. Enumeration, occurrence, and significance of injured indicator
bacteria in drinking water. In G.A. McFeters, ed. Drinking Water Microbiology:
Progress and Recent Developments, p. 478. Springer-Verlag, New York.
3. LECHEVALLIER, M.W. & G.A. MCFETERS. 1985. Enumerating injured coliforms in
drinking water. J. Amer. Water Works Assoc. 77:81.
4. BUCKLIN, K.E., G.A. MCFETERS & A. AMIRTHARAJAH. 1991. Penetration of coliforms
through municipal drinking water filters. Water Res. 25:1013.
5. LECHEVALLIER, M.W., A. SINGH, D.A. SCHIEMANN & G.A. MCFETERS. 1985. Changes in
virulence of waterborne enteropathogens with chlorine injury. Appl. Environ.
Microbiol. 50:412.
6. SINGH, A. & G.A. MCFETERS. 1986. Repair, growth and production of heat-stable
enterotoxin by E. coli following copper injury. Appl. Environ. Microbiol. 51:738.
7. SINGH, A., R. YEAGER & G.A. MCFETERS. 1986. Assessment of in vivo revival, growth,
and pathogenicity of Escherichia coli strains after copper- and chlorine-induced injury.
Appl. Environ. Microbiol. 52: 832.
8. MCFETERS, G.A., S.C. CAMERON & M.W. LECHEVALLIER. 1982. Influence of diluents,
media and membrane filters on the detection of injured waterborne coliform bacteria.
9. LECHEVALLIER, M.W. & G.A. MCFETERS. 1985. Interactions between heterotrophic plate
count bacteria and coliform organisms. Appl. Environ. Microbiol. 49:1338.
4. Bibliography
CLARK, H.F., E.E. GELDREICH, H.L. JETER & P.W. KABLER. 1951. The membrane filter in sanitary
bacteriology. Pub. Health Rep. 66:951.
MCKEE, J.E., R.T. MCLAUGHLIN & P. LESGOURGUES. 1958. Application of molecular filter
techniques to the bacterial assay of sewage. III. Effects of physical and chemical disinfection.
Sewage Ind. Wastes 30:245.
ROSE, R.E. & W. LITSKY. 1965. Enrichment procedure for use with the membrane filter for the
isolation and enumeration of fecal streptococci from water. Appl. Microbiol. 13:106.
MAXCY, R.B. 1970. Non-lethal injury and limitations of recovery of coliform organisms on
selective media. J. Milk Food Technol. 33:445.
LIN, S.D. 1973. Evaluation of coliform tests for chlorinated secondary effluents. J. Water Pollut.
Control Fed. 45:498.
BRASWELL, J.R. & A.W. HOADLEY. 1974. Recovery of Escherichia coli from chlorinated
secondary sewage. Appl. Microbiol. 28:328.
STEVENS, A.P., R.J. GRASSO & J.E. DELANEY. 1974. Measurements of fecal coliform in estuarine
water. In D.D. Wilt, ed., Proceedings of the 8th National Shellfish Sanitation Workshop, U.S.
Dep. Health, Education, & Welfare, Washington, D.C.
BISSONNETTE, G.K., J.J. JEZESKI, G.A. MCFETERS & D.S. STUART. 1975. Influence of
environmental stress on enumeration of indicator bacteria from natural waters. Appl.
Microbiol. 29:186.
BISSONNETTE, G.K., J.J. JEZESKI, G.A. MCFETERS & D.S. STUART. 1977. Evaluation of recovery
methods to detect coliforms in water. Appl. Environ. Microbiol. 33:590.
9212 B.
Recovery Enhancement
This section describes some general procedures and considerations regarding recovery of
stressed indicator organisms.
For chlorinated samples, insure that sufficient dechlorinating agent is present in the sample
bottle (see Section 9060A.2). 1 Collect water samples with elevated concentrations of
heavy-metal ions in a sample bottle containing a chelating agent2 (see Section 9060A.2) and
minimize sample storage time (see Section 9060B). Use buffered peptone dilution water rather
than buffered water (see Section 9050C.1) when preparing dilutions of samples containing
heavy-metal ions. After making dilutions, inoculate test media within 30 min.
Resuscitation of stressed or injured organisms is enhanced by inoculating samples and
initially culturing organisms in an enriched, noninhibitory medium at a moderate temperature.
Although no simple test is available to establish the presence of injured bacteria in a given
sample, bacteria in water known to contain stressors such as disinfectants or heavy metals
frequently will be injured.1,3 When multiple-tube fermentation test results consistently are higher
than those obtained from parallel membrane filter tests, or there is other indication of suboptimal
recovery, consider injury probable and use one or more of the following procedures.
1. Recovery of Injured Total Coliform Bacteria Using Membrane Filtration
a. m-T7 agar: Use m-T7 agar4 in the procedure described for the membrane filter test (see
Section 9222B).
Proteose peptone No. 3
Yeast extract
Lactose
Tergitol 7
Polyoxyethylene ether W1
Bromthymol blue
Bromcresol purple
Agar
Reagent-grade water
5.0
3.0
20.0
0.4
5.0
0.1
0.1
15.0
1
g
g
g
mL
g
g
g
g
L
Adjust to pH 7.4 with 0.1N NaOH after sterilization at 121°C for 15 min. Aseptically add 1.0
µg penicillin G/mL when medium has cooled to about 45°C.
After filtering sample place filter on m-T7 agar and incubate at 35°C for 22 to 24 h. Coliform
colonies are yellow. Verify not less than 10% of coliform colonies by the procedure in Section
9222B.5 f. With some drinking water samples containing many non-coliform bacteria, confluent
growth occurs. To obtain reliable results, carefully distinguish target yellow colonies from
background growth.
b. Addition of sodium sulfite: The addition of sodium sulfite to some media (0.05 to 0.1%)
can improve the detection of coliform bacteria following exposure to chloramine but not
chlorine.5 Such modified medium is applicable to clean-water systems using chloramination or
to chlorinated discharges such as wastewater effluent containing high levels of organic
compounds.
2. Recovery of Injured Fecal Coliform Bacteria Using Membrane Filtration
a. Enrichment-temperature acclimation: Use two-layer agar (M-FC agar with a nonselective
overlay medium that does not contain glucose, i.e., tryptic soy agar) with a 2-h incubation at
35°C followed by 22 h at 44.5°C.6 Prepare the M-FC agar plate in advance but do not add the
overlay agar more than 1 h before use.
Alternatively, use a pre-enrichment in phenol red lactose broth incubated at 35°C for 4 h
followed by M-FC agar at 44.5°C for 22 h.7
As a third option, prepare enrichment two-layer medium containing specific additives and
incubate for 1.5 h at room temperature (22 to 26°C) followed by 35°C for 4.5 h and 44.5°C for
18 h.8
b. Temperature acclimation:9 Modify elevated temperature procedure by preincubation of
M-FC cultures for 5 h at 35°C, followed by 18 ± 1 h at 44.5°C. Use a commercially available
temperature-programmed incubator to make the change from 35 to 44.5°C after the 5 h
preincubation period to eliminate inconvenience and provide a practical method of analysis.
c. Deletion of suppressive agent:10 Eliminate rosolic acid from M-FC medium and incubate
cultures at 44.5°C ± 0.2°C for 24 h. Fecal coliform colonies are intense blue on the modified
medium and are distinguished from the cream, gray, and pale-green colonies typically produced
by nonfecal coliforms.
d. Alternative medium-temperature acclimation: Use m-T7 medium with an 8 h incubation at
37°C followed by 12 h at 44.5°C.11
e. Verification of stressed fecal coliform bacteria: Modifications of media and procedures
may decrease selectivity and differentiation of fecal coliform colonies. Therefore, if any
procedural modifications are used, verify not less than 10% of the blue colonies from a variety of
samples. Use lauryl tryptose broth (Section 9221B) (35°C for 48 h) with transfer of
gas-producing cultures to EC broth (Section 9221E) (44.5°C for 24 h). Gas production at 44.5°C
confirms the presence of fecal coliforms.
3. Recovery of Stressed Fecal Streptococci Using Membrane Filtration
Using bile broth medium yields fecal streptococcus recoveries comparable with
multiple-tube fermentation tests.12 Preincubate membrane filters on an enrichment medium for 2
h at 35°C and follow by plating on m-Enterococcus agar (Section 9230) for 48 ± 2 h at 35°C.
Verification of stressed fecal streptococci—Verify not less than 10% of the colonies from a
variety of samples using the confirmed test procedure given in Section 9230B.3.
4. References
1. MCFETERS, G.A. & A.K. CAMPER. 1983. Enumeration of coliform bacteria exposed to
chlorine. In A.I. Laskin, ed. Advances in Applied Microbiology, Vol. 29, p. 177.
2. DOMEK, M.J., M.W. LECHEVALLIER, S.C. CAMERON & G.A. MCFETERS. 1984. Evidence
for the role of metals in the injury process of coliforms in drinking water. Appl.
3. LECHEVALLIER, M.W. & G.A. MCFETERS. 1985. Interactions between heterotrophic plate
count bacteria and coliform organisms. Appl. Environ. Microbiol. 49:1338.
4. LECHEVALLIER, M.W., S.C. CAMERON & G.A. MCFETERS. 1983. New medium for the
improved recovery of coliform bacteria from drinking water. Appl. Environ. Microbiol.
45:484.
5. WATTERS, S.K., B.H. PYLE, M.W. LECHEVALLIER & G.A. MCFETERS. 1989. Enumeration
of E. cloacae after chlorine exposure. Appl. Environ. Microbiol. 55:3226.
6. ROSE, R.E., E.E. GELDREICH & W. LITSKY. 1975. Improved membrane filter method for
fecal coliform analysis. Appl. Microbiol. 29:532.
7. LIN, S.D. 1976. Membrane filter method for recovery of fecal coliforms in chlorinated
sewage effluents. Appl. Environ. Microbiol. 32: 547.
8. STUART, D.S, G.A. MCFETERS & J.E. SCHILLINGER. 1977. Membrane filter technique for
quantification of stressed fecal coliforms in the aquatic environment. Appl. Environ.
Microbiol. 34:42.
9. GREEN, B.L., E.M. CLAUSEN & W. LITSKY. 1977. Two-temperature membrane filter
method for enumerating fecal coliform bacteria from chlorinated effluents. Appl.
10. PRESSWOOD, W.G. & D. STRONG. 1977. Modification of M-FC medium by eliminating
rosolic acid. Amer. Soc. Microbiol. Abs. Annu. Meeting. ISSN-0067-2777:272.
11. LECHEVALLIER, M.W., P.E. JAKANOSKI, A.K. CAMPER & G.A. MCFETERS. 1984.
Evaluation of m-T7 agar as a fecal coliform medium. Appl. Environ. Microbiol.
48:371.
12. LIN, S.D. 1974. Evaluation of fecal streptococci tests for chlorinated secondary
effluents. J. Environ. Eng. Div., Proc. Amer. Soc. Civil Engr. 100:253.
9213
RECREATIONAL WATERS*#(12)
9213 A.
Introduction
1. Microbiological Indicators
Recreational waters include freshwater swimming pools, whirlpools, and naturally occurring
fresh and marine waters. Many local and state health departments require microbiological
monitoring of recreational waters. Historically, the most common microbiological tests to assess
sanitary quality have been heterotrophic counts and total and fecal coliform tests. Total coliform
tests and heterotrophic counts usually are performed on treated waters and fecal coliform tests
performed on untreated waters. Although detection of coliform bacteria in water indicates that it
may be unsafe to drink, other bacteria have been isolated from recreational waters that may
suggest health risks through body contact, ingestion, or inhalation. Other bacteria suggested as
indicators of recreational water quality include Pseudomonas aeruginosa, fecal streptococci,
enterococci, and staphylococci. Ideally, recreational water quality indicators are microorganisms
for which densities in the water can be related quantitatively to potential health hazards resulting
from recreational use, particularly where upper body orifices are exposed to water. The ideal
indicator is the one with the best correlation between density and the health hazards associated
with a given type of pollution. The most common potential sources of infectious agents in
recreational waters include untreated or poorly treated municipal and industrial effluents or
sludge, sanitary wastes from seaside residences, fecal wastes from pleasure craft, drainage from
sanitary landfills, stormwater runoff, and excretions from animals. In addition, the source of
infectious agents may be the aquatic environment itself. The potential health hazards from each
of these sources are not equal. Exposure to untreated or inadequately treated human fecal wastes
is considered the greatest health hazard. The presence of microbiological indicators in treated
swimming pools or whirlpools indicate possible insufficient water exchange, disinfection, and
maintenance. Bather density is a major factor in determining the probability of
swimmer-associated illnesses with swimming pools, particularly when there is insufficient
disinfection and water circulation. The bathers themselves may be the source of pollution by
shedding organisms associated with the mouth, nose, and skin.
2. Infectious Diseases from Water Exposure
In general, infections or disease associated with recreational water contact fall into two
categories. The first group is gastroenteritis resulting from unintentional ingestion of water
contaminated with fecal wastes. Enteric microorganisms that have been shown to cause
gastroenteritis from recreational water contact include Giardia, Cryptosporidium, Shigella,
Salmonella, E. coli 0157:H7, Hepatitis A, Coxsackie A and B, and Norwalk virus. Leptospirosis
is not an enteric infection but also is transmitted through contact with waters contaminated with
human or animal wastes. The second group or category of infections or disease is associated
mainly with microorganisms that are indigenous to the environment, which include the
following: Pseudomonas aeruginosa, Staphylococcus sp., Legionella sp., Naegleria fowleri,
Mycobacterium sp., and Vibrio sp. The illnesses or waterborne diseases caused by these
organisms include dermatitis or folliculitis, otitis externa, Pontiac fever, granulomas, primary
amebic meningoencephalitis (PAM), and conjunctivitis. Commonly occurring illnesses or
infections associated with recreational water contact are dermatitis caused by Pseudomonas
aeruginosa and otitis externa, ‘‘swimmer’s ear,’’ frequently caused by Pseudomonas aeruginosa
and Staphylococcus aureus.
3. Microbiological Monitoring Limitations
Routine examination for pathogenic microorganisms is not recommended except for
investigations of water-related illness and special studies; in such cases, focus microbiological
analyses on the known or suspected pathogen. Methods for several of these pathogens are given
in Section 9260, Detection of Pathogenic Bacteria, Section 9510, Detection of Enteric Viruses,
and Section 9711, Pathogenic Protozoa. Because some pathogenic organisms such as Giardia,
Cryptosporidium, Mycobacterium, and Naegleria are more resistant to changes in environmental
conditions than indicator bacteria, routine monitoring may not always reflect the risk of infection
from these organisms. Described below are recommended methods for microbial indicators of
recreational water quality. Consider the type(s) of water examined in selecting the
microbiological method(s) or indicator(s) to be used. No single procedure is adequate to isolate
all microorganisms from contaminated water. While bacterial indicators may not adequately
reflect risk of viral, fungal, or parasitic infection from recreational waters, available technology
limits monitoring for such organisms in routine laboratory operations.
4. Bibliography
CABELLI, V.J. 1977. Indicators of recreational water quality. In Bacterial Indicators/Health
Hazards Associated with Waters. STP 635, American Soc. Testing & Materials,
Philadelphia, Pa.
DUFOUR, A.P. 1986. Diseases caused by water contact. In Waterborne Diseases in the United
States. CRC Press Inc., Boca Raton, Fla.
MOE, C.L. 1996. Waterborne transmission of infectious agents. In Manual of Environmental
Microbiology. American Soc. Microbiology, ASM Press, Washington, D.C.
9213 B.
Swimming Pools
a. Characteristics: A swimming pool is a body of water of limited size contained in a
holding structure.1 The pool water generally is potable and treated with additional disinfectant
but also may come from thermal springs or salt water. Modern pools have a recirculating system
for filtration and disinfection.
b. Monitoring requirements:
1) General—Monitor water quality in pools for changes in chemical and physical
characteristics that may result in irritation to the bather’s skin, eyes, and mucosal barriers or may
adversely affect disinfection. Microorganisms of concern typically are those from the bather’s
body and its orifices and include those causing infections of the eye, ear, upper respiratory tract,
skin, and intestinal or genitourinary tracts. Water quality depends on the efficacy of disinfection,
sanitary conditions, the number of bathers in the pool at any one time, and the total number of
bathers per day.
2) Disinfected indoor pools—Swimming pools should be disinfected continuously when in
use. Test swimming pool water for residual chlorine and pH when the pool is initially opened
and at least three times/d. Collect samples from at least two locations for these determinations.
Evaluate clarity of the swimming pool water before opening for the day and during periods of
heavy usage.2 The heterotrophic plate count is the primary indicator of disinfection efficacy.
Indicators of health risk include normal skin flora that are shed, such as Pseudomonas and
Staphylococcus.3-6 These organisms account for a large percentage of
swimming-pool-associated illnesses. In special circumstances Mycobacterium, Legionella, or
Candida albicans may be associated with health risks related to recreational waters. Take
samples for microbiological examination while the pool is in use. APHA recommends for public
swimming pools that not more than 15% of the samples collected during any 30-d period shall
have a heterotrophic plate count of 200/mL or show a positive confirmed total coliform test in
any of five 10-mL portions of sample examined with the multiple-tube fermentation test or more
than 1 total coliform/50 mL when the membrane filter test is used. Whenever swimming pool
samples are examined for total staphylococci or Staphylococcus aureus, not more than 50
organisms/100 mL should be present.2
3) Disinfected outdoor pools—Fecal coliform bacteria and Pseudomonas species are the
primary indicators of contamination from animal pets, rodents, stormwater runoff, and human
sources. Supporting indicators include coliform bacteria, the heterotrophic plate count, and
staphylococci.
4) Untreated pools—The primary indicator may be fecal coliform bacteria. Supporting
indicators are those described for disinfected pools. Untreated pools are not recommended for
recreational use due to increased health risks.
2. Samples
a. Containers: Collect samples for bacteriological examination of swimming pool water as
directed in Section 9060A. Use containers with capacities of 120 to 480 mL, depending on
analyses to be made. Add sufficient sodium thiosulfate, Na2S2O3, to the sample to provide a
concentration of approximately 100 mg/L in the sample. Do this by adding 0.1 mL of 10%
solution of Na2S2O3 to a 120-mL bottle or 0.4 mL to a 480-mL bottle. After adding Na2S2O3,
stopper or cap and sterilize container.
b. Sampling procedure: Collect samples during periods of maximum bather load.
Information on number of bathers may be helpful in subsequent interpretation of laboratory
results. Use sampling frequency consistent with state and local health regulations.
Collect samples by carefully removing cap of a sterile sample bottle and holding bottle near
the base at an angle of 45 deg. Fill in one slow sweep down through the water, with the mouth of
the bottle always ahead of the hand. Avoid contamination of the sample by floating debris.
Replace cap. Do not rinse bottle (i.e., retain sodium thiosulfate). For pools equipped with a filter,
samples may be collected from sampling cocks provided in the return and discharge lines from
the filter.
Most bacteria shed by bathers are in body oils, saliva, and mucus discharges that occur near
the surface; collect additional samples of the surface microlayer from the area in 1-m-deep water.
Collect microlayer samples by plunging a sterile glass plate (approximately 20 cm by 20 cm)
vertically through the water surface and withdrawing it upward at a rate of approximately 6 cm/s.
Remove surface film and water layer adhering to both sides of plate with a sterile silicone rubber
scraper and collect in a sterile glass bottle. Repeat until desired volume is obtained. To minimize
microbial contamination, wrap glass plate and scraper in metal foil and sterilize by autoclaving
before use. Wear sterile rubber or plastic gloves during sampling or hold glass plate with
forceps, clips, or tongs.
Determine residual chlorine or other disinfectant at poolside at the time of sample collection
(see Section 4500-Cl.G). Residual disinfectant levels, chemical, and physical quality of pool
water should be consistent with local, state, or APHA standards. The permissible bathing load
should adhere to local, state, or APHA-recommended regulations.
c. Sample storage: Analyze microbiological samples as soon as possible after collection (see
Section 9060B).
d. Sample volume: See Section 9222B.5.
e. Sample dilution: If sample dilutions are required, use 0.1% peptone water or buffered
dilution water as diluent to optimize recovery of stressed organisms (see Section 9222 for
suggested sample volume). Because peptone water has a tendency to foam, avoid including air
bubbles when pipetting to assure accurate measure.
3. Heterotrophic Plate Count
Determine the heterotrophic plate count as directed in Section 9215. Use at least two plates
per dilution.
4. Tests for Total Coliforms
Determine total coliform bacteria as directed in Section 9221, Section 9222, or Section
9223.
5. Tests for Fecal Coliforms
Test for fecal coliforms according to the multiple-tube fermentation technique (Section
9221), the membrane filter technique (Section 9222), or rapid methods (Section 9211).
6. Test for Staphylococci or Staphylococcus aureus
a. Baird-Parker agar base:
Tryptone
Beef extract
Yeast extract
Glycine
Sodium pyruvate
Lithium chloride
Agar
Reagent-grade water
10.0
5.0
1.0
12.0
10.0
5.0
20.0
1
g
g
g
g
g
g
g
L
Sterilize by autoclaving. Cool to 50°C and aseptically add 50 mL commercial egg yolk
tellurite enrichment/L. Mix well. Final pH should be 7.0 ± 0.2.
b. Procedure: Use membrane filter technique to prepare samples. Place membrane filter on
Baird-Parker agar and incubate at 35 ± 0.5°C for 48 ± 4 h. Staphylococci typically form slate
gray to jet black, smooth, entire colonies. If S. aureus is present egg yolk clearing may be
observed if the membrane filter is raised from the medium. Verify some differentiated colonies
with a commercial multi-test system or on the basis of such key characteristics as catalase
reaction, coagulase production, aerobic and anaerobic acid production from certain
carbohydrates, and typical microscopic morphology.
7. Test for Staphylococcus aureus
Use a modified multiple-tube procedure.
a. Media:
1) M-staphylococcus broth:
Tryptone
Yeast extract
Lactose
Mannitol
10.0
2.0
2.0
10.0
5.0
Sodium azide, NaN3
75.0
g
0.049 g
Reagent-grade water
1
g
g
g
g
g
L
Sterilize by boiling for 4 min; pH should be 7.0 ± 0.2. For 10-mL inocula prepare and use
double-strength medium.
2) Lipovitellenin-salt-mannitol agar: This medium may not be available in dehydrated form
and may require preparation from the basic ingredients or by addition of egg yolk to a
dehydrated base.
Beef extract
Polypeptone
d-Mannitol
Agar
Phenol red
Egg yolk
Reagent-grade water
1.0
10.0
75.0
10.0
15.0
0.025
20.0
1
g
g
g
g
g
g
g
L
Sterilize by autoclaving; pH should be 7.4 ± 0.2.
b. Procedure: Inoculate tubes of M-staphylococcus broth as directed in Section 9221.
Incubate at 35 ± 1°C for 24 h. Hold original enrichment sample but streak from positive (turbid)
tubes on plates of lipovitellenin-salt-mannitol agar and incubate at 35 ± 1°C for 48 h. Opaque
(24 h), yellow (48 h) zones around the colonies are positive evidence of lipovitellenin-lipase
activity (opaque) and mannitol fermentation (yellow).
If the plate is negative, streak another plate from the original enrichment tube before
discarding. Lipovitellenin-lipase activity has a 95% positive correlation with coagulase
production. If necessary, confirm positive isolates as catalase-positive, coagulase-positive,
fermenting mannitol, fermenting glucose anaerobically, yielding typical microscopic
morphology, and gram-positive.
8. Tests for Pseudomonas aeruginosa
Tests for P. aeruginosa are presented in Section 9213E and Section 9213F and include a
membrane filter procedure and a multiple-tube technique.
9. Test for Streptococci or Enterococci
Determine fecal streptococci or enterococci as described in Section 9230, and if necessary,
perform additional biochemical tests to identify species.
10. References
1. CENTERS FOR DISEASE CONTROL. 1983. Swimming Pools—Safety and Disease Control
2.
3.
4.
5.
6.
through Proper Design and Operation. DHHS—CDC No. 83-8319, Centers for Disease
Control, Atlanta, Ga.
AMERICAN PUBLIC HEALTH ASSOCIATION. 1981. Public Swimming Pools.
Recommended Regulations for Design and Construction, Operation and Maintenance.
SEYFRIED, P.L., R.S. TOBIN, N.E. BROWN & P.F. NESS. 1985. A prospective study of
swimming-related illness. II. Morbidity and the microbiological quality of water. Amer.
J. Pub. Health 75:1071.
KLAPES, N.A. & D. VESLEY. 1988. Rapid assay for in situ identification of
coagulase-positive staphylococci recovered by membrane filtration from swimming
pool water. Appl. Environ. Microbiol. 52:589.
COVERT, T.C. & P.V. SCARPINO. 1987. Comparison of Baird-Parker agar, Vogel-Johnson
agar, and M-Staphylococcus broth for the isolation and enumeration of Staphylococcus
aureus in swimming pool waters. Abstr. Annu. Meeting American Soc. Microbiology,
Atlanta, Ga., American Soc. Microbiology, Washington, D.C.
CHAROENCA, N. & R.S. FUJIOKA. 1995. Association of staphylococcal skin infections
and swimming. Water Sci. Technol. 32:11.
11. Bibliography
WORKING PARTY OF THE PUBLIC HEALTH LABORATORY SERVICE. 1965. A bacteriological
survey of swimming baths in primary schools. Monthly Bull. Min. Health & Pub. Health
Lab. Serv. 24:116.
GUNN, B.A., W.E. DUNKELBERG, JR. & J.R. CRUTZ. 1972. Clinical evaluation of 2% LSM medium
for primary isolation and identification of staphylococci. Amer. J. Clin. Pathol. 57:236.
HATCHER, R.F. & B.C. PARKER. 1974. Investigations of Freshwater Surface Microlayers.
VPI-SRRC-BULL 64. Virginia Polytechnic Inst. and State Univ., Blacksburg.
U.S. ENVIRONMENTAL PROTECTION AGENCY. 1985. Test Methods for Escherichia coli and
Enterococci in Water by the Membrane Filter Procedure. EPA-600/4-85/076.
HURST, C.J. 1991. Disinfection of drinking water, swimming-pool-water and treated sewage
effluent. In S.S. Block. Disinfection, Sterilization and Preservation, 4th ed. Lea & Febiger,
Philadelphia, Pa.
9213 C.
Whirlpools
a. Characteristics: A whirlpool is a shallow pool with a maximum water depth of 1.2 m; it
has a closed-cycle water system, a heated water supply, and usually a hydrojet recirculation
system. It may be constructed of plastic, fiberglass, redwood, or epoxy-lined surfaces.
Whirlpools are designed for recreational as well as therapeutic use and may accommodate one or
more bathers. These pools usually are not cleaned, drained, and refilled after each use. They are
located in homes, apartments, hotels, athletic facilities, rehabilitation centers, and hospitals.
b. Monitoring requirements: Whirlpool-associated infections are common because of the
inherent design and characteristics of whirlpools, which include high temperature, reduced
disinfection efficacy, and increased organic material. All these factors contribute to favorable
conditions for growth of microorganisms, especially Pseudomonas aeruginosa. Studies have
shown that whirlpools can serve as a reservoir of Legionella pneumophila. Therefore, frequent
testing for residual disinfectant levels and pH, along with scheduled maintenance, is necessary
for safe whirlpool water quality.1-5
c. Microbiological indicators: The primary indicator of disinfection efficacy is P.
aeruginosa, with total coliforms, heterotrophic plate count, and staphylococci as supporting
indicators of water quality. The standard index of water quality, i.e., total coliforms, may be
insufficient to judge the microbiological quality of whirlpool water. Pseudomonas aeruginosa is
frequently isolated from whirlpool water that is coliform-negative.6 In the event of a
whirlpool-associated outbreak, collect samples as close as possible to the time of the outbreak.
Analyze for the suspected pathogen and P. aeruginosa. Methods for P. aeruginosa are described
in Section 9213E and Section 9213F.
d. Sample preservation: Examine samples as soon as possible after collection. See Section
9060B.
2. References
1. CENTERS FOR DISEASE CONTROL. 1981. Suggested Health and Safety Guidelines for
Public Spas and Hot Tubs. DHHS-CDC #99-960. United States Government Printing
Off., Washington, D.C.
2. SOLOMON, S.L. 1985. Host factors in whirlpool-associated Pseudomonas aeruginosa
skin disease. Infect. Control 6:402.
3. HIGHSMITH, A.K., P.N. LEE, R.F. KHABBAZ & V.P. MUNN. 1985. Characteristics of
Pseudomonas aeruginosa isolated from whirlpools and bathers. Infect. Control 6:407.
4. GROOTHUIS, D.G., A.H. HAVELAAR & H.R. VEENENDAAL. 1985. A note on legionellas in
whirlpools. J. Appl. Bacteriol. 58:479.
5. HIGHSMITH, A.K. & M.S. FAVERO. 1985. Microbiological aspects of public whirlpools.
Clin. Microbiol. Newsletter 7:9.
6. HALL, N. 1984. Whirlpools and Pseudomonas aeruginosa. UHL Lab Hotline 21:9.
3. Bibliography
GELDREICH, E.E., A.K. HIGHSMITH & W.J. MARTONE. 1985. Public whirlpools—the epidemiology
and microbiology of disease. Infect. Control 6:392.
9213 D.
Natural Bathing Beaches
a. Characteristics: A natural bathing beach is any area of a stream, lake, ocean,
impoundment, or hot spring that is used for recreation. A wide variety of pathogenic
microorganisms can be transmitted to humans through use of natural fresh and marine
recreational waters contaminated by wastewater.1,2 These include enteric pathogens such as
Salmonella, Shigella, enteroviruses, protozoa, multicellular parasites, and ‘‘opportunists’’ such
as P. aeruginosa, Klebsiella sp., Vibrio sp., and Aeromonas hydrophila, which can multiply in
recreational waters with sufficient nutrients. Other organisms of concern are those associated
with the skin, mouth, or nose of bathers, such as Staphylococcus aureus and other organisms,
e.g., nontuberculous mycobacteria and leptospira, and Naegleria sp..3-9
b. Monitoring requirements: Historically, fecal coliforms have been recommended as the
indicator of choice for evaluating the microbiological quality of recreational waters. Many states
have adopted use of this indicator in their water quality standards. Recent studies have
demonstrated that E. coli and enterococci showed a stronger correlation with
swimming-associated gastroenteritis than do fecal coliforms, and that both indicators were
equally acceptable for monitoring fresh-water quality. For marine water, enterococci showed the
strongest relationship of density to gastroenteritis. The recommended densities of these indicator
organisms were calculated to approximate the degree of protection previously accepted for fecal
coliforms. EPA-recommended water quality criteria are based on these findings.10 While the
primary indicators of water quality are E. coli and enterococci, the enumeration of P.
aeruginosa, Aeromonas hydrophila, and Klebsiella sp. in recreational waters may be useful in
cases of discharge of pulp and paper wastes and effluents from textile finishing plants into
receiving waters.
2. Samples
a. Containers: Collect samples as directed in Section 9060A. The size of the container
varies with the number and variety of tests to be performed. Adding Na2S2O3 to the bottle is
unnecessary.
b. Sampling procedure: Collect samples 0.3 m below the water surface in the areas of
greatest bather load. Take samples over the range of environmental and climatic conditions,
especially during times when maximal pollution can be expected, i.e., periods of tidal, current,
and wind influences, stormwater runoff, wastewater treatment bypasses. See Section 9213B.2b
for methods of sample collection and Section 9222 for suggested sample volumes.
c. Sample storage: See Section 9060B.
3. Tests for Escherichia coli
a. Media:
1) mTEC agar:*#(13)
Proteose peptone
Yeast extract
Lactose
Dipotassium phosphate, K2HPO4
Monopotassium phosphate KH2PO4
Bromcresol purple
Bromphenol red
Agar
Reagent-grade water
5.0
3.0
10.0
7.5
3.3
g
g
g
g
g
1.0
g
0.2
0.1
0.08
0.08
15.0
1
g
g
g
g
g
L
Sterilize by autoclaving; pH should be 7.3 ± 0.2. Pour 4 to 5 mL liquefied agar into culture
dishes (50 × 10 mm). Store in refrigerator.
2) Urea substrate:* #(14)
Urea
Phenol red
Reagent-grade water
2.0 g
10 mg
100 mL
Adjust pH to between 3 and 4. Store at 2 to 8°C. Use within 1 week.
b. Procedure: Filter sample through a membrane filter (see Section 9222), place membrane
on mTEC agar, incubate at 35 ± 0.5°C for 2 h to rejuvenate injured or stressed bacteria, and then
incubate at 44.5 ± 0.2°C for 22 h. Transfer filter to a filter pad saturated with urea substrate.
After 15 min, count yellow or yellow-brown colonies, using a fluorescent lamp and a magnifying
lens. E. coli produces yellow or yellow-brown colonies. Verify a portion of these differentiated
colonies with a commercial multi-test system [see Section 9222B.5 f2)b)].
4. Tests for Enterococci
Perform tests for enterococci by the multiple-tube technique (Section 9230B) or membrane
filter technique (Section 9230C).
5. Tests for Pseudomonas aeruginosa
Perform tests for P. aeruginosa as directed in Section 9213E and Section 9213F. Use the
multiple-tube test with samples but note that the procedures may not be applicable to marine
samples.
6. Tests for Salmonella/Shigella
See Section 9260.
7. References
1. CABELLI, V.J. 1980. Health Effects Criteria for Marine Recreational Waters.
EPA-600/1-80-031, U.S. Environmental Protection Agency, Research Triangle Park,
N.C.
2. DUFOUR, A.P. 1984. Health Effects Criteria for Fresh Recreational Waters.
EPA-600/1-84-004, U.S. Environmental Protection Agency, Research Triangle Park,
N.C.
3. KESWICK, B.H., C.P. GERBA & S.M. GOYAL. 1981. Occurrence of enteroviruses in
community swimming pools. Amer. J. Pub. Health 71: 1026.
4. DUTKA, B.J. & K.K. KWAN. 1978. Health indicator bacteria in water surface microlayers.
Can. J. Microbiol. 24:187.
5. CABELLI, V.J., H. KENNEDY & M.A. LEVIN. 1976. Pseudomonas aeruginosa and fresh
recreational waters. J. Water Pollut. Control Fed. 48: 367.
6. SHERRY, J.P., S.R. KUCHMA & B.J. DUTKA. 1979. The occurrence of Candida albicans in
Lake Ontario bathing beaches. Can. J. Microbiol. 25:1036.
7. STEVENS, A.R., R.L. TYNDALL, C.C. COUTANT & E. WILLAERT. 1977. Isolation of the
etiological agent of primary amoebic meningoencephalitis from artificially heated
waters. Appl. Environ. Microbiol. 34:701.
8. WELLINGS, F.M., P.T. AMUSO, S.L. CHANG & A.L. LEWIS. 1977. Isolation and
identification of pathogenic Naegleria from Florida lakes. Appl. Environ. Microbiol.
34:661.
9. N’DIAYE, A., P. GEORGES, A. N’GO & B. FESTY. 1985. Soil amoebas as biological
markers to estimate the quality of swimming pool waters. Appl. Environ. Microbiol.
49:1072.
10. U.S. ENVIRONMENTAL PROTECTION AGENCY. 1986. Ambient Water Quality Criteria
for Bacteria—1986. EPA-440/5-84-002, U.S. Environmental Protection Agency,
Washington, D.C.
8. Bibliography
OLIVIERI, V.P., C.W. DRUSE & K. KAWATA. 1977. Microorganisms in Urban Stormwater.
EPA-600/2-77-087, U.S. Environmental Protection Agency, Cincinnati, Ohio.
RICE, E.W., T.C. COVERT, D.K. WILD, D. BERMAN, S.A. JOHNSON & C.H. JOHNSON. 1993.
Comparative resistance of Escherichia coli and Enterococci to chlorination. J. Environ.
Health. A28:89.
9213 E.
Membrane Filter Technique for Pseudomonas aeruginosa
1. Laboratory Apparatus
See Section 9222B.1.
2. Culture Media
a. M-PA agar: This agar may not be available in dehydrated form and may require
L-lysine HCl
Yeast extract
Xylose
Sucrose
Lactose
Phenol red
Ferric ammonium citrate
Sodium thiosulfate, Na2S2O3
Agar
Reagent-grade water
5.0
5.0
2.0
2.5
1.25
1.25
0.08
0.8
6.8
15.0
1
g
g
g
g
g
g
g
g
g
g
L
Adjust to pH 6.5 ± 0.1 and sterilize by autoclaving. Cool to 55 to 60°C; readjust to pH 7.1 ±
0.2 and add the following dry antibiotics per liter of agar base: sulfapyridine,*#(15) 176 mg;
kanamycin,* 8.5 mg; nalidixic acid,* 37.0 mg; and cycloheximide,* 150 mg. After mixing
dispense in 3-mL quantities in 50- × 12- mm petri plates. Store poured plates at 2 to 8°C. Discard
unused medium after 1 month.
b. Modified M-PA agar.†#(16)
c. Milk agar (Brown and Scott Foster Modification):
Mixture A:
Instant nonfat milk‡#(17)
Reagent-grade water
100
500
g
mL
Nutrient broth
Agar
Reagent-grade water
12.5
2.5
15.0
500
g
g
g
mL
Mixture B:
Separately prepare and sterilize Mixtures A and B; cool rapidly to 55°C; aseptically combine
mixtures and dispense 20 to 25 mL per petri dish.
3. Procedure
a. Presumptive tests: Filter 200 mL or less of natural waters or up to 500 mL of swimming
pool waters through sterile membrane filters. Place each membrane on a poured plate of
modified M-PA agar so that there is no air space between the membrane and the agar surface.
Invert plates and incubate at 41.5 ± 0.5°C for 72 h.
Typically, P. aeruginosa colonies are 0.8 to 2.2 mm in diameter and flat in appearance with
light outer rims and brownish to greenish-black centers. Count typical colonies, preferably from
filters containing 20 to 80 colonies. Use a 10- to 15-power magnifier as an aid in colony
counting.
b. Confirmation tests: Use milk agar to confirm a number of typical and atypical colonies.
Make a single streak (2 to 4 cm long) from an isolated colony on a milk agar plate and incubate
at 35 ± 1.0 °C for 24 h. P. aeruginosa hydrolyzes casein and produces a yellowish to green
diffusible pigment.
4. Interpretation and Calculation of Density
Confirmation is not routinely required. In the absence of confirmation, report results as the
number of presumptive P. aeruginosa/100 mL.
5. Bibliography
DRAKE, C.H. 1966. Evaluation of culture media for the isolation and enumeration of
Pseudomonas aeruginosa. Health Lab. Sci. 3:10.
BROWN, M.R.W. & J.H. SCOTT FOSTER. 1970. A simple diagnostic milk medium for Pseudomonas
aeruginosa. J. Clin. Pathol. 23:172.
LEVIN, M.A. & V.J. CABELLI. 1972. Membrane filter technique for enumeration of Pseudomonas
aeruginosa. Appl. Microbiol. 24:864.
DUTKA, B.J. & K.K. KWAN. 1977. Confirmation of the single-step membrane filter procedure for
estimating Pseudomonas aeruginosa densities in water. Appl. Environ. Microbiol. 33:240.
BRODSKY, M.H. & B.W. CIEBIN. 1978. Improved medium for recovery and enumeration of
Pseudomonas aeruginosa from water using membrane filters. Appl. Environ. Microbiol.
36:26.
9213 F.
Multiple-Tube Technique for Pseudomonas aeruginosa
See Section 9221.
2. Culture Media
a. Asparagine broth: This medium may not be available in dehydrated form and may require
Asparagine, DL
Anhydrous dipotassium hydrogen phosphate, K2HPO4
3.0 g
1.0 g
Magnesium sulfate, MgSO4⋅7H2O
0.5 g
Reagent-grade water
1
L
Adjust pH to 6.9 to 7.2 before sterilization.
b. Acetamide broth: This medium may not be available in dehydrated form and may require
Acetamide
Anhydrous dipotassium hydrogen phosphate, K2HPO4
10.0 g
5.0 g
1.39 g
Anhydrous potassium dihydrogen phosphate, KH2PO4
0.73 g
0.5
g
Dissolve 1.2 g phenol red in 100 mL 0.01N NaOH and add 1 mL/L of acetamide broth. Use
phenol red stock solution within 1 year. Adjust pH to 7.1 to 7.3 before sterilization. Final pH
should be 7.0 ± 0.2. Prepare acetamide broth as described above. If agar slants are preferred,
prepare as described above but add 15 g agar/L, heat to dissolve agar, and dispense 8-mL
quantities in 16-mm tubes. After autoclaving, incline tubes while cooling to provide a large slant
surface.
3. Procedure
a. Presumptive test: Perform a five-tube multiple-tube test. Use 10 mL single-strength
asparagine broth for inocula of 1 mL or less and 10 mL double-strength asparagine broth for
10-mL inocula. For swimming pools, higher dilutions may be necessary. Incubate inoculated
tubes at 35 to 37°C. After 24 h and again after 48 h of incubation, examine tubes under
long-wave ultraviolet light (black light) in a darkened room. Production of a green fluorescent
pigment constitutes a positive presumptive test.
b. Confirmed test: Confirm positive tubes by inoculating 0.1 mL of culture into acetamide
broth or onto the surface of acetamide agar slants. Development of purple color (alkaline pH)
within 24 to 36 h of incubation at 35 to 37°C is a positive confirmed test for Pseudomonas
aeruginosa.
c. Computing and reporting results: Refer to Table 9221:IV and to Section 9221D.
9215
HETEROTROPHIC PLATE COUNT*#(18)
9215 A.
Introduction
1. Applications
The heterotrophic plate count (HPC), formerly known as the standard plate count, is a
procedure for estimating the number of live heterotrophic bacteria in water and measuring
changes during water treatment and distribution or in swimming pools. Colonies may arise from
pairs, chains, clusters, or single cells, all of which are included in the term ‘‘colony-forming
units’’ (CFU). The final count also depends on interaction among the developing colonies;
choose that combination of procedure and medium that produces the greatest number of colonies
within the designated incubation time. To compare data, use the same procedure and medium.
Three different methods and four different media are described.
2. Selection of Method
a. Pour plate method: The pour plate method (9215B) is simple to perform and can
accommodate volumes of sample or diluted sample ranging from 0.1 to 2.0 mL. The colonies
produced are relatively small and compact, showing less tendency to encroach on each other
than those produced by surface growth. On the other hand, submerged colonies often are slower
growing and are difficult to transfer. A thermostatically controlled water bath is essential for
tempering the agar, but even so, significant heat shock to bacteria from the transient exposure of
the sample to 45 to 46°C agar may occur.
b. Spread plate method: The spread plate method (9215C) causes no heat shock and all
colonies are on the agar surface where they can be distinguished readily from particles and
bubbles. Colonies can be transferred quickly, and colony morphology easily can be discerned
and compared to published descriptions. However, this method is limited by the small volume of
sample or diluted sample that can be absorbed by the agar: 0.1 to 0.5 mL, depending on the
degree to which the prepoured plates have been dried. To use this procedure, maintain a supply
of suitable predried, absorbent agar plates.
c. Membrane filter method: The membrane filter method (9215D) permits testing large
volumes of low-turbidity water and is the method of choice for low-count waters (< 1 to 10 CFU/
mL). This method produces no heat shock but adds the expense of the membrane filter. Further
disadvantages include the smaller display area, the need to detect colonies by reflected light
against a white background if colored filters or contrast stains are not used, possible damage to
cells by excessive filtration pressures, and possible variations in membrane filter quality (see
Section 9020B.4h).
3. Work Area
Provide a level table or bench top with ample area in a clean, draft-free, well-lighted room
or within a horizontal-flow laminar hood. Use table and bench tops having nonporous surfaces
and disinfect before any analysis is made.
4. Samples
Collect water as directed in Section 9060A. Initiate analysis as soon as possible after
collection to minimize changes in bacterial population. The recommended maximum elapsed
time between collection and analysis of samples is 8 h (maximum transit time 6 h, maximum
processing time 2 h). When analysis cannot begin within 8 h, maintain sample at a temperature
below 4°C but do not freeze. Maximum elapsed time between collection and analysis must not
exceed 24 h.
5. Sample Preparation
Mark each plate with sample number, dilution, date, and any other necessary information
before examination. Prepare at least duplicate plates for each volume of sample or dilution
examined. For the pour or spread plate methods use sterile glass (65 cm2) or presterilized
disposable plastic (57 cm2) petri dishes.
Thoroughly mix all samples or dilutions by rapidly making about 25 complete up-and-down
(or back-and-forth) movements. Optionally, use a mechanical shaker to shake samples or
dilutions for 15 s.
6. Media
Compare new lots of media with current lot in use according to Section 9020B.4i.
a. Plate count agar (tryptone glucose yeast agar): Use for pour and spread plate methods.
This high-nutrient agar, widely used in the past, gives lower counts than R2A or NWRI agar. It
is included for laboratories wishing to make comparisons of media or to extend the continuity of
old data.
Tryptone
Yeast extract
Glucose
Agar
Reagent-grade water
5.0
2.5
1.0
15.0
1
g
g
g
g
L
pH should be 7.0 ± 0.2 after autoclaving at 121°C for 15 min.
b. m-HPC agar:†#(19) Use this high-nutrient medium only for the membrane filter method.
Peptone
Gelatin
Glycerol
Agar
Reagent-grade water
20.0
25.0
10.0
15.0
1
g
g
mL
g
L
Mix all ingredients except glycerol. Adjust pH to 7.1, if necessary, with 1N NaOH, heat
slowly to boili ng to dissolve thoroughly, add glycerol, and autoclave at 121°C for 5 min.‡#(20)
c. R2A agar: Use for pour, spread plate, and membrane filter methods. This low-nutrient agar
gives higher counts than high-nutrient formulations.
Yeast extract
Casamino acids
Glucose
Soluble starch
0.5
0.5
0.5
0.5
0.5
0.3
Magnesium sulfate heptahydrate, MgSO4⋅7H2O
0.05 g
Sodium pyruvate
Agar
Reagent-grade water
0.3
15.0
1
g
g
g
g
g
g
g
g
L
Adjust pH to 7.2 with solid K2HPO4 or KH2PO4 before adding agar. Heat to dissolve agar
and sterilize at 121°C for 15 min.
d. NWRI agar (HPCA): Use for pour, spread plate, and membrane filter methods. This
low-nutrient medium is likely to produce higher colony counts than high-nutrient media. It is not
currently available in dehydrated form and requires preparation from the basic ingredients; this
makes its usage less desirable.
Peptone
Soluble casein
K2HPO4
3.0
0.5
0.2
g
g
g
MgSO4
0.05
g
FeCl3
0.001 g
Agar
Reagent-grade water
15.0
1
g
L
Adjust pH to 7.2 before autoclaving for 15 min at 121°C.
7. Incubation
For compliance monitoring purposes under U.S. EPA’s Surface Water Treatment Rule (40
CFR 141.74), provision on heterotrophic bacteria, incubate pour plates at 35°C for 48 h.
Otherwise, select from among recommended times and temperatures for monitoring changes in
water quality. The highest counts typically will be obtained from 5- to 7-d incubation at a
temperature of 20 to 28°C.
During incubation maintain humidity within the incubator so that plates will have no
moisture weight loss greater than 15%. This is especially important if prolonged incubation is
used. A pan of water placed at the bottom of the incubator may be sufficient but note that to
prevent rusting or oxidation the inside walls and shelving should be of high-grade stainless steel
or anodized aluminum. For long incubation in nonhumidified incubators, seal plates in plastic
bags.
8. Counting and Recording
a. Pour and spread plates: Count all colonies on selected plates promptly after incubation. If
counting must be delayed temporarily, store plates at 5 to 10°C for no more than 24 h, but avoid
this as routine practice. Record results of sterility controls on the report for each lot of samples.
Use an approved counting aid, such as the Quebec colony counter, for manual counting. If
such equipment is not available, count with any other counter provided that it gives equivalent
magnification and illumination. Automatic plate counting instruments are available. These
generally use a television scanner coupled to a magnifying lens and an electronics package.
Their use is acceptable if evaluation in parallel with manual counting gives comparable results.
In preparing plates, pipet sample volumes that will yield from 30 to 300 colonies/plate. The
aim is to have at least one dilution giving colony counts between these limits, except as provided
below.
Ordinarily, do not pipet more than 2.0 mL of sample; however, when the total number of
colonies developing from 2.0 mL is less than 30, disregard the rule above and record result
observed. With this exception, consider only plates having 30 to 300 colonies in determining the
plate count. Compute bacterial count per milliliter by the following equation:
If there is no plate with 30 to 300 colonies, and one or more plates have more than 300
colonies, use the plate(s) having a count nearest 300 colonies. Compute the count as above and
report as estimated CFU per milliliter.
If plates from all dilutions of any sample have no colonies, report the count as less than one
(< 1) divided by the corresponding largest sample volume used. For example, if no colonies
develop from the 0.01-mL sample volume, report the count as less than 100 (< 100) estimated
CFU/mL.
If the number of colonies per plate far exceeds 300, do not report result as ‘‘too numerous to
count’’ (TNTC). If there are fewer than 10 colonies/cm2, count colonies in 13 squares (of the
colony counter) having representative colony distribution. If possible, select seven consecutive
squares horizontally across the plate and six consecutive squares vertically, being careful not to
count a square more than once. Multiply sum of the number of colonies in 13 representative
square centimeters by 5 to compute estimated colonies per plate when the plate area is 65 cm2.
When there are more than 10 colonies/cm2, count four representative squares, take average count
per square centimeter, and multiply by the appropriate factor to estimate colonies per plate. The
factor is 57 for disposable plastic plates and 65 for glass plates. When bacterial counts on
crowded plates are greater than 100 colonies/cm2, report result as greater than (>) 6500 divided
by the smallest sample volume plated for glass plates or greater than (>) 5700 divided by the
smallest sample volume plated for plastic plates. Report as estimated colony-forming units per
milliliter.
If spreading colonies (spreaders) are encountered on the plate(s) selected, count colonies on
representative portions only when colonies are well distributed in spreader-free areas and the
area covered by the spreader(s) does not exceed one-half the plate area.
When spreading colonies must be counted, count each of the following types as one: a chain
of colonies that appears to be caused by disintegration of a bacterial clump as agar and sample
were mixed; a spreader that develops as a film of growth between the agar and bottom of petri
dish; and a colony that forms in a film of water at the edge or over the agar surface. The last two
types largely develop because of an accumulation of moisture at the point from which the
spreader originates. They frequently cover more than half the plate and interfere with obtaining a
reliable plate count.
Count as individual colonies similar-appearing colonies growing in close proximity but not
touching, provided that the distance between them is at least equal to the diameter of the smallest
colony. Count impinging colonies that differ in appearance, such as morphology or color, as
individual colonies.
If plates have excessive spreader growth, report as ‘‘spreaders’’ (Spr). When plates are
uncountable because of missed dilution, accidental dropping, and contamination, or the control
plates indicate that the medium or other material or labware was contaminated, report as
‘‘laboratory accident’’ (LA).
b. Membrane filter method: Count colonies on membrane filters using a stereoscopic
microscope at 10 to 15 × magnification. Preferably slant petri dish at 45° angle on microscope
stage and adjust light source vertical to the colonies. Optimal colony density per filter is 20 to
200. If colonies are small and there is no crowding, a higher limit is acceptable.
Count all colonies on the membrane when there are ≤ 2 colonies per square. For 3 to 10
colonies per square count 10 squares and obtain average count per square. For 10 to 20 colonies
per square count 5 squares and obtain average count per square. Multiply average count per
square by 100 and divide by the sample volume to give colonies per milliliter. If there are more
than 20 colonies per square, record count as > 2000 divided by the sample volume. Report
averaged counts as estimated colony-forming units. Make estimated counts only when there are
discrete, separated colonies without spreaders.
9. Computing and Reporting Counts
The term ‘‘colony-forming units’’ (CFU) is descriptive of the methods used; therefore,
report all counts as colony-forming units. Include in the report the method used, the incubation
temperature and time, and the medium. For example: CFU/mL, pour plate method, 35°C/48 h,
plate count agar.
To compute the heterotrophic plate count, CFU/mL, divide total number of colonies or
average number (if duplicate plates of the same dilution) per plate by the sample volume. Record
sample volumes used and number of colonies on each plate counted or estimated.
When colonies on duplicate plates and/or consecutive dilutions are counted and results are
averaged before being recorded, round off counts to two significant figures only when
converting to colony-forming units.
Avoid creating fictitious precision and accuracy when computing colony-forming units by
recording only the first two left-hand digits. Raise the second digit to the next higher number
when the third digit from the left is 5, 6, 7, 8, or 9; use zeros for each successive digit toward the
right from the second digit. For example, report a count of 142 as 140 and a count of 155 as 160,
but report a count of 35 as 35.
10. Analytical Bias
Avoid inaccuracies in counting due to carelessness, damaged or dirty optics that impair
vision, or failure to recognize colonies. Laboratory workers who cannot duplicate their own
counts on the same plate within 5% and the counts of other analysts within 10% should discover
the cause and correct such disagreements.
9215 B.
Pour Plate Method
1. Samples and Sample Preparation
See Section 9215A.4 and Section 9215A.5.
2. Sample Dilution
Prepare water used for dilution blanks as directed in Section 9050C.
a. Selecting dilutions: Select the dilution(s) so that the total number of colonies on a plate
will be between 30 and 300 (Figure 9215:1). For example, where a heterotrophic plate count as
high as 3000 is suspected, prepare plates with 10−2 dilution.
For most potable water samples, plates suitable for counting will be obtained by plating 1
mL and 0.1 mL undiluted sample and 1 mL of the 10−2 dilution.
b. Measuring sample portions: Use a sterile pipet for initial and subsequent transfers from
each container. If pipet becomes contaminated before transfers are completed, replace with a
sterile pipet. Use a separate sterile pipet for transfers from each different dilution. Do not prepare
dilutions and pour plates in direct sunlight. Use caution when removing sterile pipets from the
container; to avoid contamination, do not drag pipet tip across exposed ends of pipets in the pipet
container or across lips and necks of dilution bottles. When removing sample, do not insert
pipets more than 2.5 cm below the surface of sample or dilution.
c. Measuring dilutions: When discharging sample portions, hold pipet at an angle of about
45° with tip touching bottom of petri dish or inside neck of dilution bottle. Lift cover of petri
dish just high enough to insert pipet. Allow 2 to 4 s for liquid to drain from 1-mL graduation
mark to tip of pipet. If pipet is not a blow-out type, touch tip of pipet once against a dry spot on
petri dish bottom. Less preferably, use a cotton-plugged blow-out-type pipet and gently blow out
remaining volume of sample dilution. When 0.1-mL quantities are measured, let diluted sample
drain from chosen reference graduation until 0.1 mL has been delivered. Remove pipet without
retouching it to dish. Pipet 1 mL, 0.1 mL, or other suitable volume into sterile petri dish before
adding melted culture medium. Use decimal dilutions in preparing sample volumes of less than
0.1 mL; in examining sewage or turbid water, do not measure a 0.1-mL inoculum of original
sample, but prepare an appropriate dilution. Prepare at least two replicate plates for each sample
dilution used. After depositing test portions for each series of plates, pour culture medium and
mix carefully. Do not let more than 20 min elapse between starting pipetting and pouring plates.
3. Plating
a. Melting medium: Melt sterile solid agar medium in boiling water or by exposure to
flowing steam in a partially closed container, but avoid prolonged exposure to unnecessarily high
temperatures during and after melting. Do not resterilize plating medium. If the medium is
melted in two or more batches, use all of each batch in order of melting, provided that the
contents remain fully melted. Discard melted agar that contains precipitate.
Maintain melted medium in a water bath between 44 and 46°C until used, preferably no
longer than 3 h. In a separate container place a thermometer in water or medium that has been
exposed to the same heating and cooling as the plating medium. Do not depend on the sense of
touch to indicate proper medium temperature when pouring agar.
Use plate count agar, R2A agar, or NWRI agar as specified in Section 9215A.6. Before using
a new lot of medium test its suitability.
b. Pouring plates: Limit the number of samples to be plated in any one series so that no more
than 20 min (preferably 10 min) elapse between dilution of the first sample and pouring of the
last plate in the series. Pour at least 10 to 12 mL liquefied medium maintained at 44 to 46°C into
each dish by gently lifting cover just high enough to pour. Carefully avoid spilling medium on
outside of container or on inside of dish lid when pouring. When pouring agar from flasks or
tubes that have been held in a water bath, wipe with clean paper towel and flame the neck before
pouring. As each plate is poured mix melted medium thoroughly with test portions in petri dish,
taking care not to splash mixture over the edge, by rotating the dish first in one direction and
then in the opposite direction, or by rotating and tilting. Let plates solidify (within 10 min) on a
level surface. After medium solidifies, invert plates and place in incubator.
c. Sterility controls: Check sterility of medium and dilution water blanks by pouring control
plates for each series of samples. Prepare additional controls to determine contamination of
plates, pipets, and room air.
4. Incubation
See Section 9215A.7.
5. Counting, Recording, Computing, and Reporting
6. Bibliography
BREED, R.S. & W.D. DOTTERER. 1916. The number of colonies allowable on satisfactory agar
plates. Tech. Bull. 53, New York Agricultural Experiment Sta.
BUTTERFIELD, C.T. 1933. The selection of a dilution water for bacteriological examinations. J.
Bacteriol. 23:355; Pub. Health Rep. 48: 681.
ARCHAMBAULT, J., J. CUROT & M.H. MCCRADY. 1937. The need of uniformity of conditions for
counting plates (with suggestions for a standard colony counter). Amer. J. Pub. Health
27:809.
RICHARDS, O.W. & P.C. HEIJN. 1945. An improved dark-field Quebec colony counter. J. Milk
Technol. 8:253.
BERRY, J.M., D.A. MCNEILL & L.D. WITTER. 1969. Effect of delays in pour plating on bacterial
counts. J. Dairy Sci. 52:1456.
GELDREICH, E.E., H.D. NASH, D.J. REASONER & R.H. TAYLOR. 1972. The necessity of controlling
bacterial populations in potable waters: Community water supply. J. Amer. Water Works
Assoc. 64:596.
GELDREICH, E.E. 1973. Is the total count necessary? Proc. 1st Annu. AWWA Water Quality
Technol. Conf., Dec. 3-4, 1973. Cincinnati, Ohio, p. VII-1. American Water Works Assoc.,
Denver, Colo.
GINSBURG, W. 1973. Improved total count techniques. Proc. 1st Annu. AWWA Water Quality
Technol. Conf., Dec. 3-4, 1973. Cincinnati, Ohio, p. VIII-1. American Water Works Assoc.,
Denver, Colo.
DUTKA, B.J., A.S.Y. CHAU & J. COBURN. 1974. Relationship of heterotrophic bacterial indicators
of water pollution and fecal sterols. Water Res. 8:1047.
KLEIN, D.A. & S. WU. 1974. Stress: a factor to be considered in heterotrophic microorganism
enumeration from aquatic environments. Appl. Microbiol. 37:429.
GELDREICH, E.E., H.D. NASH, D.J. REASONER & R.H. TAYLOR. 1975. The necessity for controlling
bacterial populations in potable waters: Bottled water and emergency water supplies. J.
Amer. Water Works Assoc. 67:117.
BELL, C.R., M.A. HOLDER-FRANKLIN & M. FRANKLIN. 1980. Heterotrophic bacteria in two
Canadian rivers.—I. Seasonal variation in the predominant bacterial populations. Water Res.
14:449.
MEANS, E.G., L. HANAMI, G.F. RIDGWAY & B.H. OLSON. 1981. Evaluating mediums and plating
techniques for enumerating bacteria in water distribution systems. J. Amer. Water Works
Assoc. 73: 585.
AMERICAN PUBLIC HEALTH ASSOCIATION. 1993. Standard Methods for the Examination of
Dairy Products, 16th ed. American Public Health Assoc., Washington, D.C.
REASONER, D.J. & E.E. GELDREICH. 1985. A new medium for the enumeration and subculture of
bacteria from potable water. Appl. Environ. Microbiol. 49:1.
9215 C.
Spread Plate Method
a. Glass rods: Bend 4-mm-diam fire-polished glass rods, 200 mm in length, 45° about 40
mm from one end. Sterilize before using.
b. Pipet, glass, 1.1 mL, with tempered, rounded tip. Do not use disposable plastic pipets.
c. Turntable (optional).*#(21)
d. Incubator or drying oven, set at 42°C, or laminar-flow hood.
2. Media
See Section 9215A.6a, Section 9215A.6c, and Section 9215A.6d. If R2A agar is used best
results are obtained at 28°C with 7 d incubation; if NWRI is used, incubate at 20°C for 7 d.
3. Preparation of Plates
Pour 15 mL of the desired medium into sterile 100 × 15 or 90 × 15 petri dishes; let agar
solidify. Predry plates inverted so that there is a 2- to 3-g water loss overnight with lids on. See
Figure 9215:2, Table 9215:I, or Figure 9215:3. Use predried plates immediately after drying or
store for up to 2 weeks in sealed plastic bags at 4°C. For predrying and using plates the same
day, pour 25 mL agar into petri dish and dry in a laminar-flow hood at room temperature (24 to
26°C) with the lid off to obtain the desired 2- to 3-g weight loss. See Figure 9215:3.
4. Procedure
Prepare sample dilutions as directed in 9215B.2.
a. Glass rod: Pipet 0.1 or 0.5 mL sample onto surface of predried agar plate. Using a sterile
bent glass rod, distribute inoculum over surface of the medium by rotating the dish by hand or on
a turntable. Let inoculum be absorbed completely into the medium before incubating.
b. Pipet: Pipet desired sample volume (0.1, 0.5 mL) onto the surface of the predried agar
plate while dish is being rotated on a turntable. Slowly release sample from pipet while making
one to-and-fro motion, starting at center of the plate and stopping 0.5 cm from the plate edge
before returning to the center. Lightly touch the pipet to the plate surface. Let inoculum be
absorbed completely by the medium before incubating.
5. Incubation
See Section 9215A.7.
7. Bibliography
BUCK, J.D. & R.C. CLEVERDON. 1960. The spread plate as a method for the enumeration of
marine bacteria. Limnol. Oceanogr. 5:78.
CLARK, D.S. 1967. Comparison of pour and surface plate methods for determination of bacterial
counts. Can. J. Microbiol. 13:1409.
VAN SOESTBERGAN, A.A. & C.H. LEE. 1969. Pour plates or streak plates. Appl. Microbiol.
18:1092.
CLARK, D.S. 1971. Studies on the surface plate method of counting bacteria. Can. J. Microbiol.
17:943.
GILCHRIST, J.E., J.E. CAMPBELL, C.B. DONNELLY, J.T. PELLER & J.M. DELANEY. 1973. Spiral plate
method for bacterial determination. Appl. Microbiol. 25:244.
PTAK, D.M. & W. GINSBURG. 1976. Pour plate vs. streak plate method. Proc. 4th Annu. AWWA
Water Quality Technol. Conf., Dec. 6-7, 1976. San Diego, Cal., p. 2B-5. American Water
Works Assoc., Denver, Colo.
DUTKA, B.J., ed. 1978. Methods for Microbiological Analysis of Waters, Wastewaters and
Sediments. Inland Waters Directorate, Scientific Operation Div., Canada Centre for Inland
Waters, Burlington, Ont.
KAPER, J.B., A.L. MILLS & R.R. COLWELL. 1978. Evaluation of the accuracy and precision of
enumerating aerobic heterotrophs in water samples by the spread method. Appl. Environ.
Microbiol. 35:756.
YOUNG, M. 1979. A modified spread plate technique for the determination of concentrations of
viable heterotrophic bacteria. STP 673:41-51, American Soc. Testing & Materials,
Philadelphia, Pa.
GELDREICH, E.E. 1981. Current status of microbiological water quality criteria. ASM News
47:23.
TAYLOR, R.H., M.J. ALLEN & E.E. GELDREICH. 1981. Standard plate count: A comparison of pour
plate and spread plate methods. Proc. 9th Annu. AWWA Water Quality Technol. Conf., Dec.
6-9, 1981. Seattle, Wash., p. 223. American Water Works Assoc. Denver, Colo.
9215 D.
Membrane Filter Method
2. Media
See Section 9215A.6. Use m-HPC agar, or alternatively R2A or NWRI agar.
3. Preparation of Plates
Dispense 5-mL portions of sterile medium*#(22) into 50- × 9-mm petri dishes. Let solidify
at room temperature. Prepared plates may be stored inverted in a plastic bag or tight container in
a refrigerator, for no longer than 2 weeks.
4. Sample Size
The volume to be filtered will vary with the sample. Select a maximum sample size to give
20 to 200 CFU per filter.
5. Procedure
Filter appropriate volume through a sterile 47-mm, 0.45-µm, gridded membrane filter, under
partial vacuum. Rinse funnel with three 20- to 30-mL portions of sterile dilution water. Place
filter on agar in petri dish.
6. Incubation
Place dishes in close-fitting box or plastic bag containing moistened paper towels. Incubate
at 35 ± 0.5°C for 48 h if using m-HPC agar, or longer if using R2A medium, or at 20 to 28°C for
5 to 7 d if using NWRI or R2A agar. Duplicate plates may be incubated for other time and
temperature conditions as desired.
See Section 9215A.8 and Section 9215A.9. Report as CFU/mL, membrane filter method,
time, medium.
8. Bibliography
STOPERT, E.M., W.T. SOKOSKI & J.T. NORTHAM. 1962. The factor of temperature in the better
recovery of bacteria from water by filtration. Can. J. Microbiol. 8:809.
TAYLOR, R.H. & E.E. GELDREICH. 1979. A new membrane filter procedure for bacterial counts in
potable water and swimming pool samples. J. Amer. Water Works Assoc. 71:402.
CLARK, J.A. 1980. The influence of increasing numbers of non-indicator organisms upon the
detection of indicator organisms by the membrane filter and presence-absence tests. Can. J.
Microbiol. 20: 827.
DUTKA, B.J., ed. 1981. Membrane Filtration, Applications, Techniques, and Problems. Marcel
Dekker, Inc., New York, N.Y. and Basel, Switzerland.
HOADLEY, A.W. 1981. Effect of injury on the recovery of bacteria on membrane filters. In B. J.
Dutka, ed. Membrane Filtration, Applications, Techniques, and Problems, p. 413. Marcel
Dekker, Inc., New York, N.Y. and Basel, Switzerland.
9216
DIRECT TOTAL MICROBIAL COUNT*#(23)
9216 A.
Introduction
Direct total cell counts of bacteria in water or wastewater usually exceed counts obtained
from heterotrophic plate counts and most probable number methods because, unlike those
procedures, direct counts preclude errors caused by viability-related phenomena such as
selectivity of growth media, cell clumping, and slow growth rates.
9216 B.
Epifluorescence Microscopic Method
The epifluorescence microscopic method produces direct total cell counts with relative
speed (20 to 30 min from time of sampling) and sensitivity. It does not permit differentiation of
bacterial cells on the basis of taxonomy, metabolic activity, or viability, and it cannot be used to
estimate the microbial biomass because of considerable variation in the volume of individual
cells. The method requires an experienced technician who can distinguish microbial cells from
debris on the basis of morphology.
The method consists of sample fixation for storage, staining with a chemical fluorochrome,
vacuum filtration onto a nonfluorescing polycarbonate membrane, and enumeration by counting
with an epifluorescence microscope.
2. Apparatus
a. Microscope, vertical UV illuminator for epifluorescence with flat field 100× oil
immersion objective lens, to give total magnification of at least 1000×.
b. Counting graticule, ocular lens micrometer* calibrated with stage micrometer.*#(24)
c. Filters,†#(25) including excitation filters (KP 490 and LP 455), beam splitter (LP 510),
and barrier filter (LP 520 using mercury lamp, HBO 50).
d. Blender or vortex mixer.
e. Filtration unit, suitable for use with 25-mm-diam membrane filters.
f. Membrane filters, polycarbonate,‡#(26) 25-mm-diam, 0.2-µm pore size (purchase
nonfluorescent or prepare by soaking membrane in Irgalan black [2 g/L in 2% acetic acid] for 24
h, then rinse in water and air dry); cellulosic§#(27) 25-mm-diam, 5-µm pore size.
g. Syringes, 3-mL, disposable, with disposable syringe filters, 0.2-µm pore size.
h. Test tubes, glass, screw-capped, 13- × 125-mm.
3. Reagents
a. Phosphate buffer: Dissolve 13.6 g KH2PO4 in water and dilute to 1 L. Adjust to pH 7.2 if
necessary; filter through 0.2-µm membrane filter.
b. Fixative, 5.0% (w/v) glutaraldehyde in phosphate buffer. Prepare fresh daily.
c. Fluorochrome, 0.1% (w/v) acridine orangei#(28) in phosphate buffer.
d. Immersion oil, low fluorescing.##(29)
4. Procedure
Collect water samples as directed in Section 9060. Add 9.0 mL sample to test tube
containing 1.0 mL fixative. Fixed samples can be stored at 4°C for up to 3 weeks without
significant decrease in cell numbers.
Disperse and dilute samples from mesotrophic or eutrophic sources to obtain reproducible
results. Mix sample using blender or vortex mixer, then make tenfold dilutions in phosphate
buffer as necessary. Clean water samples may not require dilution but larger sample volumes
(>100 mL) may be required to obtain reliable counts.
Place 1 mL sample or dilution on a nonfluorescent polycarbonate filter supported by a
cellulosic membrane filter in filter holder. Using disposable sterile syringe filters, add 1 mL
fluorochrome and wait 2 min, then add about 3 mL filtered phosphate buffer to promote more
even cell distribution. Alternatively, combine fluorochrome with sample in a small clean vial, let
react, and add mixture to filter holder. Filter with vacuum (about 13 kPa). Wash with 2 mL
phosphate buffer and filter. Remove polycarbonate filter with forceps and air dry for 1 to 2 min.
The filter can be cut into quarter sections and saved if needed. Place dried filter on a drop of
immersion oil on a clean glass microscope slide. Add a small drop of immersion oil to filter
surface. Gently cover filter with a clean glass cover slip. Samples can be stored in the dark for
several months without significant loss of fluorescence.
Examine at least 10 randomly selected fields on the filter using the 100× oil immersion lens
to establish that distribution of microbial cells is uniform and that individual cells can be
enumerated (if not, dilute sample and repeat). Preferably count 10 to 50 cells per field. Count
number of cells in at least 20 squares using the calibrated counting graticule.
5. Calculations
Calculate the average number of cells per filter. Obtain effective filter area from
specifications of filtration unit. Extrapolate to determine number of cells per milliliter of sample:
Total cells/mL = (avg cells/square) × (squares/filter) × (dilution factor) / sample volume, mL.
6. Bibliography
HOBBIE, J.E., R.J. DALEY & S. JASPER. 1977. Use of nuclepore filters for counting bacteria by
fluorescence microscopy. Appl. Environ. Microbiol. 33:1225.
SIERACK, M.E., P.W. JOHNSON & J.MCH. SIEBURTH. 1985. Detection, enumeration, and sizing of
planktonic bacteria by image-analyzed epifluorescence microscopy. Appl. Environ.
Microbiol. 49:799.
AMERICAN SOCIETY FOR TESTING AND MATERIALS. 1987. Standard test method for
enumeration of aquatic bacteria by epifluorescence microscopy counting procedure. ASTM
D4455-85, Annual Book of ASTM Standards, Vol. 11.02, Water. American Soc. Testing &
Materials, Philadelphia, Pa.
9217
ASSIMILABLE ORGANIC CARBON*#(30)
9217 A.
Introduction
1. Significance
Growth of bacteria in drinking water distribution and storage systems can lead to the
deterioration of water quality, violation of water quality standards, and increased operating costs.
Growth or regrowth results from viable bacteria surviving the disinfection process and utilizing
nutrients in the water and biofilm to sustain growth.1 Factors other than nutrients that influence
regrowth include temperature,2 residence time in mains and storage units,3 and the efficacy of
disinfection.4 Tests to determine the potential for bacterial regrowth focus on the concentration
of nutrients.5-7
Not all organic compounds are equally susceptible to microbial decomposition; the fraction
that provides energy and carbon for bacterial growth has been called labile dissolved organic
carbon,8,9 biodegradable organic carbon (BDOC),7 or assimilable organic carbon (AOC).5
Easily measured chemical surrogates for AOC are not available now.10,11 As alternatives to
chemical methods, bioassays have been proposed.5-7,12-14
In a bioassay, the growth of a bacterial inoculum to maximum density can be used to
estimate the concentrations of limiting nutrients; the underlying assumptions of the AOC
bioassay are that nitrogen and phosphorus are present in excess, i.e., that organic carbon is
limiting, and that the bioassay organism(s) represent the physiological capabilities of the
distribution system microflora. Various bioassay procedures use an inoculum of one to four
species of bacteria5,12,13,15,16 growing in log phase or present in late stationary phase, or may
use undefined bacteria attached to a sand substratum,7 suspended in the sample,6 or filtered from
the sample and then resuspended.14 Incubation vessels vary as to material,17 size,18,19 closure,18
and cleaning procedure.5,18,19 Water to be tested for nutrient concentrations has been variously
prepared.5,7,14 The AOC bioassay is an indirect or surrogate method, wherein nutrient
concentrations are not measured directly, but colony-forming units (CFU) of the bioassay
organism(s) are the test variable. Nutrient concentrations have been estimated directly from
changes in dissolved organic carbon concentrations within the test vessel7 or indirectly from
epifluorescence microscopic counts of the maximum number of bacterial cells grown,13,14
turbidity,14 or incorporation of tritiated thymidine into bacterial DNA.6,20 CFU densities, total
cell densities, or bacterial production are converted to nutrient concentration by the growth yield
of bacteria, defined as either the ratio between CFU or cells produced and organic carbon used,
or biomass produced and organic carbon used.5,6
The method described below is a two-species bioassay using Pseudomonas fluorescens
strain P-17 and Spirillum strain NOX (van der Kooij)10 that has been modified to reduce
problems of bacterial and carbon contamination.18,19 It uses a defined inoculum and
miniaturized incubation vessels, requires no specialized equipment, and has been related to the
presence of coliforms in a drinking water distribution system.22 The two-species inoculum
probably underestimates the total quantity of AOC, is consistently lower than BDOC estimates,
and does not provide an estimate of refractory organic carbon.23 Critical aspects of the proposed
method, including the preparation of the incubation vessel, test water, and inoculum, and
enumeration of the test organisms, are transferable to alternate AOC assays that use a different
defined inoculum.
With an undefined bacterial inoculum, enumeration by the spread plate technique is not
applicable; alternate response variables, such as changes in dissolved organic carbon (DOC)
concentration, turbidity, epifluorescence microscopic counts, bacterial mortality, or bacterial
growth, have been used.6,7,14
3. Sampling and Storage
Follow precautions outlined in Section 9060A and Section 9060B for collecting and storing
samples. Pasteurized and dechlorinated water samples probably can be held for several days
without deterioration if properly sealed. Initiate the AOC assay as quickly as possible after
pasteurization (see ¶ B.4c).
4. References
1. CHARACKLIS, W.G. 1988. Bacterial Regrowth in Distribution Systems. American Water
Works Assoc. Research Foundation Research Rep., American Water Works Assoc.,
Denver, Colo.
2. FRANSOLET, G., G. VILLERS & W.J. MASSCHELEIN. 1985. Influence of temperature on
bacterial development in waters. Ozone Sci. Eng. 7: 205.
3. MAUL, A., A.H. EL-SHAARAWI & J.C. BLOCK. 1985. Heterotrophic bacteria in water
distribution systems. I. Spatial and temporal variation. Sci. Total Environ. 44:201.
4. LECHEVALLIER, M.W., C.D. CAWTHON & R.G. LEE. 1988. Factors promoting survival of
bacteria in chlorinated water supplies. Appl. Environ. Microbiol. 54:649.
5. VAN DER KOOIJ, D., A. VISSER & W.A.M. HIJNEN. 1982. Determining the concentration of
easily assimilable organic carbon in drinking water. J. Amer. Water Works Assoc.
74:540.
6. SERVAIS, P., G. BILLEN & M.C. HASCOET. 1987. Determination of the biodegradable
fraction of dissolved organic matter in waters. Water Res. 21:445.
7. JORET, J.C., Y. LEVI, T. DUPIN & M. GILBERT. 1988. Rapid method for estimating
bioeliminable organic carbon in water. In Proc. Annu. Conf. American Water Works
Association, June 19–23, 1988, Orlando, Fla., p. 1715. American Water Works Assoc.,
Denver, Colo.
8. WETZEL, R.G. & B.A. MANNY. 1972. Decomposition of dissolved organic carbon and
nitrogen compounds from leaves in an experimental hard-water stream. Limnol.
Oceanogr. 17:927.
9. OGURA, N. 1975. Further studies on decomposition of dissolved organic matter in
coastal seawater. Mar. Biol. 31:101.
10. VAN DER KOOIJ, D. 1988. Assimilable Organic Carbon (AOC) in Water. In The Search
for a Surrogate. AWWA Research Foundation/KIWA Cooperative Research Rep. p.
311. American Water Works Assoc. Research Foundation, Denver, Colo.
11. KAPLAN, L.A. & T.L. BOTT. 1990. Nutrients for Bacterial Growth in Drinking Water:
Bioassay Evaluation. EPA Project Summary, EPA-600/S2-89-030: 1-7. U.S.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
Environmental Protection Agency, Washington, D.C.
KENNY, F.A., J.C. FRY & R.A. BREACH. 1988. Development and Operational
Implementation of Modified and Simplified Method for Determination of Assimilable
Organic Carbon (AOC) in Drinking Water. International Assoc. Water Pollution
Research & Control, Brighton, U.K., pp. 1–5.
NEDWELL, D.B. 1987. Distribution and pool sizes of microbially available carbon in
sediment measured by a microbiological assay. Microbiol. Ecol. 45:47.
WERNER, P. 1984. Investigations on the substrate character of organic substances in
connection with drinking water treatment. Zentralbl. Bakt. Hyg. 180:46.
VAN DER KOOIJ, D. & W.A.M. HIJNEN. 1983. Nutritional versatility of a starch utilizing
Flavobacterium at low substrate concentrations. Appl. Environ. Microbiol. 45:804.
VAN DER KOOIJ, D. & W.A.M. HIJNEN. 1984. Substrate utilization of an
oxalate-consuming Spirillum species in relation to its growth in ozonated water. Appl.
COLBOURNE, J.S., R.M TREW & P.J. DENNIS. 1988. Treatment of water for aquatic
bacterial growth studies. J. Appl. Bacteriol. 65:79.
KAPLAN, L.A. & T.L. BOTT. 1989. Measurement of assimilable organic carbon in water
distribution systems by a simplified bioassay technique. In Advances in Water
Analysis and Treatment, Proc. 16th Annu. AWWA Water Quality Technology Conf.,
Nov. 13–17, 1988, St. Louis, Mo., p. 475. American Water Works Assoc., Denver,
Colo.
KAPLAN, L.A., T.L. BOTT & D.J. REASONER. 1993. Evaluation and simplification of the
assimilable organic carbon nutrient bioassay for bacterial growth in drinking water.
Appl. Environ. Microbiol. 59: 1532.
MORIARTY, D.J.W. 1986. Measurement of bacterial growth rates in aquatic systems
from rates of nucleic acid synthesis. In K.C. Marshall, ed. Advan. Microb. Ecol. 9:245.
VAN DER KOOIJ, D., W.A.M. HIJNEN & J.C. KRUITHOF. 1989. The effects of ozonation,
biological filtration and distribution on the concentration of easily assimilable organic
carbon (AOC) in drinking water. Ozone Sci. Eng. 11:297.
LECHEVALLIER, M.W., W.H. SHULZ & R.G. LEE. 1989. Bacterial nutrients in drinking
water. In M.W. LeChevallier, B.H. Olson & G.A. McFeters, eds. Assessing and
Controlling Bacterial Regrowth in Distribution Systems. American Water Works
Assoc. Research Foundation Research Rep., American Water Works Assoc., Denver,
Colo.
PREVOST, M., D. DUCHESNE, J. COALLIER, R. DESJARDINS & P. LAFRANCE. 1990.
Full-scale evaluation of biological activated carbon filtration for the treatment of
drinking water. In Advances in Water Analysis and Treatment, Proc. 17th Annu.
AWWA Water Quality Technology Conf., Nov. 12–16, 1989, Philadelphia, Pa., p. 147.
American Water Works Assoc., Denver, Colo.
5. Bibliography
VAN DER KOOIJ, D. 1979. Characterization and classification of fluorescent pseudomonads
isolated from tap water and surface water. Antonie van Leeuwenhoek 45:225.
VAN DER KOOIJ, D., A. VISSER & W.A.M. HIJNEN. 1980. Growth of Aeromonas hydrophila at low
concentrations of substrates added to tap water. Appl. Environ. Microbiol. 39:1198.
WERNER, P. 1981. Microbial studies on the chemical and biological treatment of ground water
containing humic acid. Vom Wasser 57:157.
OLSON, B.H. 1982. Assessment and implications of bacterial regrowth in water distribution
systems. EPA Project Summary, EPA-600/S2-82-072:1-10. U.S. Environmental Protection
Agency, Washington, D.C.
RIZET, M., F. FIESSINGER & N. HOUEL. 1982. Bacterial regrowth in a distribution system and its
relationship with the quality of the feed water: case studies. In Proc. Annu. Conf. American
Water Works Association, May 16–20, 1982, Miami Beach, Fla., p. 1199. American Water
Works Assoc., Denver, Colo.
VAN DER KOOIJ, D., J.P. ORANJE & W.A.M. HIJNEN. 1982. Growth of Pseudomonas aeruginosa in
tap water in relation to utilization of substrates at concentrations of a few micrograms per
liter. Appl. Environ. Microbiol. 44:1086.
CAMPER, A.K., M.W. LECHEVALLIER, S.C. BROADAWAY & G.A. MCFETERS. 1986. Bacteria
associated with granular activated carbon particles in drinking water. Appl. Environ.
Microbiol. 52:434.
WENG, C., D.L. HOVEN & B.J. SCHWARTZ. 1986. Ozonation: An economic choice for water
treatment. J. Amer. Water Works Assoc. 78(11):83.
CARLUCCI, A.F., S.L. SHIMP & D.B. CRAVEN. 1987. Bacterial response to labile dissolved organic
matter increases associated with marine discontinuities. Fed. European Microbiological
Societies, Microbiol. Ecol. 45:211.
LECHEVALLIER, M.W., T.M. BABCOCK & R.G. LEE. 1987. Examination and
characterization of distribution system biofilms. Appl. Environ. Microbiol. 53:2714.
THINGSTAD, T.F. 1987. Utilization of N, P, and organic C by heterotrophic bacteria. I. Outline of
a chemostat theory with a consistent concept of maintenance metabolism. Marine Ecol.
Progr. Ser. 35:99.
ANSELME, C., I.H. SUFFET & J. MALLEVIALLE. 1988. Effects of ozonation on tastes and odors. J.
Amer. Water Works Assoc. 80(10):45.
FRANSOLET, G., A. DEPELCHIN, G. VILLERS, R. GOOSSENS & W.J. MASSCHELEIN. 1988. The role
of bicarbonate in bacterial growth in oligotrophic waters. J. Amer. Water Works Assoc.
80(11):57.
9217 B.
Pseudomonas fluorescens Strain P-17, Spirillum Strain NOX Method
a. Principle: The AOC bioassay using Pseudomonas fluorescens strain P-17 and Spirillum
strain NOX involves growth to a maximum density of a small inoculum in a batch culture of
pasteurized test water. Pasteurization inactivates native microflora. The test organisms are
enumerated by the spread plate method for heterotrophic plate counts (Section 9215C) and the
density of viable cells is converted to AOC concentrations by an empirically derived yield factor
for the growth of P-17 on acetate-carbon and NOX on oxalate-carbon as standards. The number
of organisms at stationary phase is assumed to be the maximum number of organisms that can be
supported by the nutrients in the sample and the yields on acetate carbon and oxalate carbon are
assumed to equal the yield on naturally occurring AOC.1,2
b. Interferences: Untreated surface waters, especially those with high concentrations of
suspended solids or high turbidity, can contain large numbers of spore-forming bacteria that may
survive pasteurization, grow, and interfere with the enumeration of P-17 and NOX on spread
plates. Such waters generally have high AOC concentrations and can be diluted with
organic-free water amended with mineral salts or prefiltered through carbon-free filters. Potable
waters that have been disinfected and carry a disinfectant residual will inhibit growth of the test
organism unless the disinfectant is neutralized. Surface waters from reservoirs treated with
copper sulfate also may be inhibitory unless a chelating agent is added to the sample,3 and
lime-softened waters with elevated pH values may require pH adjustment. Any amendment to a
sample requires a control for AOC contamination.
c. Minimum detectable concentration: In theory, concentrations of less than 1 µg C/L can be
detected. In practice, organic carbon contamination during glassware preparation and sample
handling imposes a limit of detection of approximately 5 to 10 µg AOC/L.
2. Apparatus
a. Incubation vessels: Organic-carbon-free borosilicate glass vials (45 mL capacity) with
TFE-lined silicone septa.
b. Incubator, set at 15 ± 0.5°C.
c. Hot water bath capable of achieving and holding 70°C.
d. Continuously adjustable pipet*#(31) capable of delivering between 10 and 100 µL.
e. Erlenmeyer flask, 125-mL, with ground-glass stopper.
f. Apparatus for preparing dilution water and making heterotrophic plate counts: See
Section 9050C and Section 9215C.
3. Reagents
a. Sodium acetate stock solution, 400 mg acetate-C/L: Dissolve 2.267 g CH3COONa⋅3H2O
in 1 L organic-carbon-free, deionized water. Transfer to 45-mL vials, fill to shoulder, cap tightly,
and autoclave. Although standard autoclave practice is to loosen caps, keep vials with septa
capped tightly for autoclaving. Store at 5°C in tightly capped vials. Solution may be held for up
to 6 months.
b. Sodium thiosulfate solution: Dissolve 30 g Na2S2O3 in 1 L deionized water. Transfer to
45-mL vials and autoclave as directed in ¶ 3a.
c. Buffered water: See Section 9050C.
d. R2A agar: See Section 9215A.6c.
e. Sodium persulfate solution, 10% (w/v): Dissolve 100 g Na2S2O8 in 1 L deionized water.
f. Organic-free water: See Section 5710B.3e. Alternatively, use HPLC-grade bottled water.
g. Mineral salts solution: Dissolve 171 mg K2HPO4, 767 mg NH4Cl, and 1.444 g KNO3 in 1
L carbon-free water. Transfer to 45-mL vials and autoclave as directed in ¶ 3a.
h. Cultures of strains P-17 (ATCC 49642) and NOX (ATCC 49643).†#(32)
4. Procedure
a. Preparation of incubation vessels: Wash 45-mL vials with detergent, rinse with hot water,
0.1N HCl two times, and deionized water three times, dry, cap with foil, and heat to 550°C for 6
h. Soak TFE-lined silicone septa in a 10% sodium persulfate solution for 1 h at 60°C; rinse three
times with carbon-free deionized water. Alternatively, use pre-cleaned water sampling vials4 or
an equivalent AOC-free vial.‡#(33) Use same cleaning procedure for all glassware.
b. Preparation of stock inoculum: Prepare individual turbid suspensions of P-17 and NOX by
transferring growth from a slant culture on R2A agar into 2 to 3 mL filtered (0.2 µm), autoclaved
sample. Use slant not older than 6 months. The autoclaved sample can be any water that supports
growth of P-17 and NOX and is organic-carbon-limited. Neutralize chlorinated samples with
sodium thiosulfate (42 µL/50 mL). Transfer 100 µL of suspension to 50 mL filtered, autoclaved
sample in a sterile 125-mL ground-glass-stoppered erlenmeyer flask. Add 125 µL sodium acetate
solution (suspension contains 1 mg acetate-C/L). Incubate at room temperature (≤ 25°C) until the
viable cell count reaches the stationary phase. Organic-carbon limitation will insure complete
utilization of acetate-C so that no AOC is transferred with the inoculum. The stationary phase is
reached when the viable cell count, as measured by spread plates, reaches maximum value. Store
stock cultures for not more than 6 months at 5°C. Before inoculating a bioassay vessel, make a
viable count of the culture (spread plate) to determine the appropriate volume of inoculum to be
added to each bioassay vessel.
c. Preparation of incubation water: Collect samples directly into 10 45-mL vials. Use 9 vials
for AOC measurement and 1 for growth control. Fill each vial to the neck (40 mL) within as
short a time as possible. Place septa on the vials, TFE side down, and secure with open-topped
screw caps. Alternatively, collect 500 mL sample in an organic-carbon-free vessel and pour into
each vial. Neutralize samples containing disinfectant residuals with 33 µL sodium thiosulfate
solution added to each vial or 0.5 mL per 500-mL sample. Preferably, collect an extra vial to
check for residual chlorine after neutralization. In the laboratory, cap vials tightly and pasteurize
in 70°C water bath for 30 min.
d. Inoculation and incubation: Cool, inoculate with 500 colony-forming units (CFU)/mL
each of P-17 and NOX, either by injecting through the septum or by removing cap and using a
carbon-free pipet. Plastic, sterile tips for continuously adjustable pipets are suitable. Use the
following equation to calculate volume of inoculum:
Hold vials at 15°C in the dark for 1 week. If a 15°C incubator is unavailable, incubate at
room temperature not to exceed 25°C. Because incubation temperature influences growth yield,
record and report temperature. Determine yields as directed below if an alternative temperature
is used.
e. Enumeration of test bacterium: On incubation days 7, 8, and 9 remove three vials from the
incubator. Sample an individual vial on only 1 d. Shake vials vigorously for 1 min, remove 1
mL with a sterile pipet, and prepare a dilution series (see Section 9215B). Plate three dilutions
(10–2, 10–3, and 10–4) in duplicate. Incubate plates at 25°C for 3 to 5 d and score the number of
colonies of each strain. P-17 colonies appear on plates first; they are 3 to 4 mm in diameter with
diffuse yellow pigmentation. NOX colonies are small (1- to 2-mm diam) white dots. It may be
necessary to count P-17 and NOX colonies at different dilutions. Sample vials on three separate
days to check whether maximum density has been reached. Day-to-day variations of between 11
and 16% of the mean for batch cultures of P-17 in stationary phase are typical.1 A consistent
increase in cell densities of 20% or more over the 3-d period indicates that the cultures are not in
stationary phase; repeat assay with longer incubation period. Alternatively, collect more samples
(three for each additional sampling day) and prepare as in ¶ c above so that extended incubation
can be used. A sharp population decrease of approximately 0.5 log over the 3 d is unusual, but
may occur. If this happens repeat the assay.
f. Determination of yield of P-17 and NOX: The yields of P-17 and NOX on model carbon
compounds should be constant if organic carbon is limiting and the incubation temperature is
kept constant. It is acceptable to use the previously derived empirical yield values of 4.1 × 106
CFU P-17/µg acetate-C, 1.2 × 107 CFU-NOX/µg acetate-C, and 2.9 × 106 CFU-NOX/µg
oxalate-C at 15°C.5 However, the determination of a yield control provides an important check
on both the bioassay (see also 6. Quality Control, below) and carbon limitation in the sample.
5. Calculation
a. AOC concentration: Average viable count results for the 3 d and calculate concentration
of AOC as the product of the mean of the viable counts and the inverse of the yield:
µg AOC/L = [(mean P-17 CFU/mL)(1/yield)
+(mean NOX CFU/mL)(1/yield)](1000 mL/L)
When the empirical yield factors5 are used, the equation becomes:
µg AOC/L = [(mean P-17 CFU/mL)(µg acetate-C/4.1 × 106 CFU)
+(mean NOX CFU/mL)(µg oxalate-C/2.9 × 106 CFU)]
(1000 mL/L)
or
µg AOC/L = [(mean P-17 CFU/mL)(2.44 × 10–7 µg acetate-C/CFU)
+(mean NOX CFU/mL)(3.45 × 10–7 µg oxalate-C/CFU)] (1000 mL/L)
In practice, the densities of organisms vary during the stationary phase. Using average
density over 3-d period provides a more accurate estimate of the real maximum density.
Reporting AOC as µg C/L assumes that the yields on acetate and oxalate are equal to the
yields on naturally occurring AOC. To permit data comparisons report incubation temperature,
contribution of each species to AOC, and yield factors used.
6. Quality Control
See Section 9020B for general quality control procedures. Quality control specific to the
AOC bioassay includes testing the inoculum for purity and viability by plating a portion on R2A
agar, testing the incubation vessel, inoculum, thiosulfate solution, and any supplemental
procedure such as filtration or dilution for organic carbon contamination, testing the P-17 and
NOX inocula for yield, and testing the sample for carbon limitation or inhibition of assay
organisms. Test all deviations in procedure (see ¶ 6).
To make these tests, use separate controls for blank, yield, and growth. The controls outlined
below use a single vial and are meant as a trouble-shooting guide. Definitive determination, for
example, that the yield is different from a published value or that a sample is inhibitory, requires
replication and statistical analysis.
a. Blank control: Dilute mineral salts solution 10:1 with carbon-free water. Follow
procedures outlined above: Fill a vial to the shoulder with organic-carbon-free water, add 100 µL
mineral salts and 100 µL sodium thiosulfate, pasteurize, inoculate with P-17/NOX, incubate, and
enumerate growth.
b. Yield control: Dilute sodium acetate or sodium oxalate solution 10:1 with carbon-free
water, preparing 40 mg C/L working concentrations. Follow procedures outlined above: Fill a
vial to the shoulder with carbon-free water, add 100 µL mineral salts, 100 µL sodium thiosulfate,
and 100 µL sodium acetate or sodium oxalate working solution, pasteurize, inoculate with
P-17/NOX, incubate, and enumerate growth. P-17, unlike NOX, will not grow with oxalate as
sole carbon source (oxalate is considered a major by-product of ozonation). NOX growth in
HPLC-grade water presumed to be organic carbon-free is to be expected. The yield control is a
quality control measurement and is not intended to provide a conversion factor for the
calculation of AOC.
c. Growth control: Use additional sample of test water collected with the nine AOC vials, (¶
4c above) but amend with 100 µL diluted mineral salts and 100 µL of diluted acetate or oxalate
solution per vial before pasteurization. As with other controls, inoculate with P-17/NOX,
incubate, and enumerate growth.
d. Yield calculations: If previously derived empirical yield values (see ¶ 4 f above) are not
used, a conversion factor can be derived empirically by using pure cultures of P-17 and NOX.
Mixed cultures of the organisms cannot be used and a separate blank control for each species is
required. Convert density units to CFU/L by multiplying CFU/mL by 1000, and divide by 100
µg acetate or oxalate-C/L. Express yield as CFU P-17 or NOX/ µg acetate-C or oxalate-C. For
P-17 and acetate-C, the equation is:
e. Interpretation of growth control: Subtract densities of P-17 and NOX that grew in the
sample amended with only thiosulfate from the densities of P-17 and NOX that grew in the
growth control. Compare difference to the difference between yield and blank controls.
If: (growth control − sample) = (yield control − blank control)
Then: sample is carbon-limited and not inhibitory
If: (growth control − sample) < (yield control − blank control)
Then: sample is inhibitory to bioassay organism
If: (growth control − sample) > (yield control − blank control)
Then: sample is not carbon-limited
f. Supplemental procedure check: When using such supplemental procedures as filtration,
dilution, or chemical amendment check for carbon contribution to the AOC values. To test a
procedure, use carbon-free water and blank control as a base line. Perform the supplemental
procedure on additional carbon-free water and compare to densities of P-17 and NOX that grow
in the blank control.
7. Precision and Bias
The P-17 bioassay performed in a single laboratory using 45-mL vials had a precision of ±
17.5% based on a total of 58 assays with 14 different samples.6
8. References
1. KAPLAN, L.A. & T.L. BOTT. 1989. Measurement of assimilable organic carbon in water
distribution systems by a simplified bioassay technique. In Advances in Water
Analysis and Treatment, Proc. 16th Annu. AWWA Water Quality Technology Conf.,
Nov. 13–17, 1988, St. Louis, Mo., p. 475. American Water Works Assoc., Denver,
Colo.
2. VAN DER KOOIJ, D., A. VISSER & J.P. ORANJE. 1982. Multiplication of fluorescent
pseudomonads at low substrate concentrations in tap water. Antonie van Leeuwenhoek
48:229.
3. LECHEVALLIER, M.W., W.H. SHULZ & R.G. LEE. 1989. Bacterial nutrients in drinking
water. In M.W. LeChevallier, B.H. Olson & G.A. McFeters, eds. Assessing and
Controlling Bacterial Regrowth in Distribution Systems. American Water Works
Assoc. Research Foundation Research Rep., American Water Works Assoc., Denver,
Colo.
4. KAPLAN, L.A. & T.L. BOTT. 1990. Modifications to simplify an AOC bioassay for
routine use by utilities monitoring bacterial regrowth potential in water distribution
systems. In Advances in Water Analysis and Treatment, Proc. 17th Annu. AWWA
Water Quality Technology Conf., Nov. 12–16, 1989, Philadelphia, Pa., p. 1031.
5. VAN DER KOOIJ, D., W.A.M. HIJNEN & J.C. KRUITHOF. 1989. The effects of ozonation,
biological filtration and distribution on the concentration of easily assimilable organic
carbon (AOC) in drinking water. Ozone Sci. Eng. 11:297.
6. KAPLAN, L.A. & T.L. BOTT. 1990. Nutrients for bacterial growth in drinking water.
Bioassay evaluation. EPA Project Summary, EPA-600/S2-89-030: 1-7. U.S.
Environmental Protection Agency, Washington, D.C.
9. Bibliography
KING, E.O., M.K. WARD & D.E. RANEY. 1954. Two simple media for the demonstration of
pyocyanin and fluorescin. J. Lab. Clin. Med. 44: 301.
MASON, J. & D.P. KELLY. 1988. Thiosulfate oxidation by obligately heterotrophic bacteria.
Microbial Ecol. 15:123.
9221
MULTIPLE-TUBE FERMENTATION TECHNIQUE FOR MEMBERS OF THE
COLIFORM GROUP*#(34)
9221 A.
Introduction
The coliform group consists of several genera of bacteria belonging to the family
Enterobacteriaceae. The historical definition of this group has been based on the method used for
detection (lactose fermentation) rather than on the tenets of systematic bacteriology.
Accordingly, when the fermentation technique is used, this group is defined as all facultative
anaerobic, gram-negative, non-spore-forming, rod-shaped bacteria that ferment lactose with gas
and acid formation within 48 h at 35°C.
The standard test for the coliform group may be carried out either by the multiple-tube
fermentation technique or presence-absence procedure (through the presumptive-confirmed
phases or completed test) described herein, by the membrane filter (MF) technique (Section
9222) or by the enzymatic substrate coliform test (Section 9223). Each technique is applicable
within the limitations specified and with due consideration of the purpose of the examination.
Production of valid results requires strict adherence to quality control procedures. Quality
control guidelines are outlined in Section 9020.
When multiple tubes are used in the fermentation technique, results of the examination of
replicate tubes and dilutions are reported in terms of the Most Probable Number (MPN) of
organisms present. This number, based on certain probability formulas, is an estimate of the
mean density of coliforms in the sample. Coliform density, together with other information
obtained by engineering or sanitary surveys, provides the best assessment of water treatment
effectiveness and the sanitary quality of source water.
The precision of each test depends on the number of tubes used. The most satisfactory
information will be obtained when the largest sample inoculum examined shows gas in some or
all of the tubes and the smallest sample inoculum shows no gas in all or a majority of the tubes.
Bacterial density can be estimated by the formula given or from the table using the number of
positive tubes in the multiple dilutions (Section 9221C.2). The number of sample portions
selected will be governed by the desired precision of the result. MPN tables are based on the
assumption of a Poisson distribution (random dispersion). However, if the sample is not
adequately shaken before the portions are removed or if clumping of bacterial cells occurs, the
MPN value will be an underestimate of the actual bacterial density.
1. Water of Drinking Water Quality
When drinking water is analyzed to determine if the quality meets the standards of the U.S.
Environmental Protection Agency (EPA), use the fermentation technique with 10 replicate tubes
each containing 10 mL, 5 replicate tubes each containing 20 mL, or a single bottle containing a
100-mL sample portion. When examining drinking water by the fermentation technique, process
all tubes or bottles demonstrating growth with or without a positive acid or gas reaction to the
confirmed phase (Section 9221B.2). Apply the completed test (Section 9221B.3) to not less than
10% of all coliform-positive samples per quarter. Obtain at least one positive sample per quarter.
A positive EC broth (Section 9221E) or a positive EC MUG broth (Section 9221F) test result is
considered an alternative to the positive completed test phase.
For the routine examination of public water supplies the object of the total coliform test is to
determine the efficiency of treatment plant operation and the integrity of the distribution system.
It is also used as a screen for the presence of fecal contamination. A high proportion of coliform
occurrences in a distribution system may be attributed not to treatment failure at the plant or the
well source, but to bacterial regrowth in the mains. Because it is difficult to distinguish between
coliform regrowth and new contamination, assume all coliform occurrences to be new
contamination unless otherwise demonstrated.
2. Water of Other than Drinking Water Quality
In the examination of nonpotable waters inoculate a series of tubes with appropriate decimal
dilutions of the water (multiples and submultiples of 10 mL), based on the probable coliform
density. Use the presumptive-confirmed phase of the multiple-tube procedure. Use the more
labor-intensive completed test (Section 9221B.3) as a quality control measure on at least 10% of
coliform-positive nonpotable water samples on a seasonal basis. The object of the examination
of nonpotable water generally is to estimate the density of bacterial contamination, determine a
source of pollution, enforce water quality standards, or trace the survival of microorganisms. The
multiple-tube fermentation technique may be used to obtain statistically valid MPN estimates of
coliform density. Examine a sufficient number of samples to yield representative results for the
sampling station. Generally, the geometric mean or median value of the results of a number of
samples will yield a value in which the effect of sample-to-sample variation is minimized.
3. Other Samples
The multiple-tube fermentation technique is applicable to the analysis of salt or brackish
waters as well as muds, sediments, and sludges. Follow the precautions given above on portion
sizes and numbers of tubes per dilution.
To prepare solid or semisolid samples weigh the sample and add diluent to make a 10−1
dilution. For example, place 50 g sample in sterile blender jar, add 450 mL sterile phosphate
buffer or 0.1% peptone dilution water, and blend for 1 to 2 min at low speed (8000 rpm). Prepare
the appropriate decimal dilutions of the homogenized slurry as quickly as possible to minimize
settling.
9221 B.
Standard Total Coliform Fermentation Technique
1. Presumptive Phase
Use lauryl tryptose broth in the presumptive portion of the multiple-tube test. If the medium
has been refrigerated after sterilization, incubate overnight at room temperature (20°C) before
use. Discard tubes showing growth and/or bubbles.
a. Reagents and culture medium:
1) Lauryl tryptose broth:
Tryptose
Lactose
20.0 g
5.0 g
2.75 g
Potassium dihydrogen phosphate, KH2PO4
2.75 g
Reagent-grade water
5.0
0.1
1
g
g
L
Add dehydrated ingredients to water, mix thoroughly, and heat to dissolve. pH should be 6.8
± 0.2 after sterilization. Before sterilization, dispense sufficient medium, in fermentation tubes
with an inverted vial, to cover inverted vial at least one-half to two-thirds after sterilization.
Alternatively, omit inverted vial and add 0.01 g/L bromcresol purple to presumptive medium to
determine acid production, the indicator of a positive result in this part of the coliform test. Close
tubes with metal or heat-resistant plastic caps.
Make lauryl tryptose broth of such strength that adding 100-mL, 20-mL, or 10-mL portions
of sample to medium will not reduce ingredient concentrations below those of the standard
medium. Prepare in accordance with Table 9221:I.
b. Procedure:
1) Arrange fermentation tubes in rows of five or ten tubes each in a test tube rack. The
number of rows and the sample volumes selected depend upon the quality and character of the
water to be examined. For potable water use five 20-mL portions, ten 10-mL portions, or a single
bottle of 100 mL portion; for nonpotable water use five tubes per dilution (of 10, 1, 0.1 mL, etc.).
In making dilutions and measuring diluted sample volumes, follow the precautions given in
Section 9215B.2. Use Figure 9215:1 as a guide to preparing dilutions. Shake sample and
dilutions vigorously about 25 times. Inoculate each tube in a set of five with replicate sample
volumes (in increasing decimal dilutions, if decimal quantities of the sample are used). Mix test
portions in the medium by gentle agitation.
2) Incubate inoculated tubes or bottles at 35 ± 0.5C. After 24 ± 2 h swirl each tube or bottle
gently and examine it for growth, gas, and acidic reaction (shades of yellow color) and, if no gas
or acidic reaction is evident, reincubate and reexamine at the end of 48 ± 3 h. Record presence or
absence of growth, gas, and acid production. If the inner vial is omitted, growth with acidity
signifies a positive presumptive reaction.
c. Interpretation: Production of an acidic reaction or gas in the tubes or bottles within 48 ± 3
h constitutes a positive presumptive reaction. Submit tubes with a positive presumptive reaction
to the confirmed phase (Section 9221B.2).
The absence of acidic reaction or gas formation at the end of 48 ± 3 h of incubation
constitutes a negative test. Submit drinking water samples demonstrating growth without a
positive gas or acid reaction to the confirmed phase (Section 9221B.2). An arbitrary 48-h limit
for observation doubtless excludes occasional members of the coliform group that grow very
slowly (see Section 9212).
2. Confirmed Phase
a. Culture medium: Use brilliant green lactose bile broth fermentation tubes for the
confirmed phase.
Brilliant green lactose bile broth:
Peptone
Lactose
Oxgall
Brilliant green
Reagent-grade water
10.0
10.0
20.0
0.0133
1
g
g
g
g
L
± 0.2 after sterilization. Before sterilization, dispense, in fermentation tubes with an inverted
vial, sufficient medium to cover inverted vial at least one-half to two-thirds after sterilization.
Close tubes with metal or heat-resistant plastic caps.
b. Procedure: Submit all presumptive tubes or bottles showing growth, any amount of gas, or
acidic reaction within 24 ± 2 h of incubation to the confirmed phase. If active fermentation or
acidic reaction appears in the presumptive tube earlier than 24 ± 2 h, transfer to the confirmatory
medium; preferably examine tubes at 18 ± 1 h. If additional presumptive tubes or bottles show
active fermentation or acidic reaction at the end of a 48 ± 3- h incubation period, submit these to
the confirmed phase.
Gently shake or rotate presumptive tubes or bottles showing gas or acidic growth to
resuspend the organisms. With a sterile loop 3.0 to 3.5 mm in diameter, transfer one or more
loopfuls of culture to a fermentation tube containing brilliant green lactose bile broth or insert a
sterile wooden applicator at least 2.5 cm into the culture, promptly remove, and plunge
applicator to bottom of fermentation tube containing brilliant green lactose bile broth. Remove
and discard applicator. Repeat for all other positive presumptive tubes.
Incubate the inoculated brilliant green lactose bile broth tube at 35 ± 0.5°C. Formation of gas
in any amount in the inverted vial of the brilliant green lactose bile broth fermentation tube at
any time (e.g., 6 ± 1 h, 24 ± 2 h) within 48 ± 3 h constitutes a positive confirmed phase.
Calculate the MPN value from the number of positive brilliant green lactose bile tubes as
described in Section 9221C.
c. Alternative procedure: Use this alternative only for polluted water or wastewater known to
produce positive results consistently.
If all presumptive tubes are positive in two or more consecutive dilutions within 24 h,
submit to the confirmed phase only the tubes of the highest dilution (smallest sample inoculum)
in which all tubes are positive and any positive tubes in still higher dilutions. Submit to the
confirmed phase all tubes in which gas or acidic growth is produced only after 48 h.
3. Completed Phase
To establish the presence of coliform bacteria and to provide quality control data, use the
completed test on at least 10% of positive confirmed tubes (see Figure 9221:1). Simultaneous
inoculation into brilliant green lactose bile broth for total coliforms and EC broth for fecal
coliforms (see Section 9221E below) or EC-MUG broth for Escherichia coli may be used.
Consider positive EC and EC-MUG broths elevated temperature (44.5°C) results as a positive
completed test response. Parallel positive brilliant green lactose bile broth cultures with negative
EC or EC-MUG broth cultures indicate the presence of nonfecal coliforms.
a. Culture media and reagents:
1) LES Endo agar: See Section 9222B. Use 100- × 15-mm petri plates.
2) MacConkey agar:
Peptone
Proteose peptone
Lactose
Bile salts
Agar
Neutral red.
Crystal violet
Reagent-grade water
17
3
10
1.5
5
13.5
0.03
0.001
1
g
g
g
g
g
g
g
g
L
Add ingredients to water, mix thoroughly, and heat to boiling to dissolve. Sterilize by
autoclaving for 15 min at 121°C. Temper agar after sterilization and pour into petri plates (100 ×
15 mm). pH should be 7.1 ± 0.2 after sterilization.
3) Nutrient agar:
Peptone
Beef extract
Agar
Reagent-grade water
5.0
3.0
15.0
1
g
g
g
L
Add ingredients to water, mix thoroughly, and heat to dissolve. pH should be 6.8 ± 0.2 after
sterilization. Before sterilization, dispense in screw-capped tubes. After sterilization,
immediately place tubes in an inclined position so that the agar will solidify with a sloped
surface. Tighten screw caps after cooling and store in a protected, cool storage area.
4) Gram-stain reagents:
a) Ammonium oxalate-crystal violet (Hucker’s): Dissolve 2 g crystal violet (90% dye
content) in 20 mL 95% ethyl alcohol; dissolve 0.8 g (NH4)2C2O4⋅H2O in 80 mL reagent-grade
water; mix the two solutions and age for 24 h before use; filter through paper into a staining
bottle.
b) Lugol’s solution, Gram’s modification: Grind 1 g iodine crystals and 2 g KI in a mortar.
Add reagent-grade water, a few milliliters at a time, and grind thoroughly after each addition
until solution is complete. Rinse solution into an amber glass bottle with the remaining water
(using a total of 300 mL).
c) Counterstain: Dissolve 2.5 g safranin dye in 100 mL 95% ethyl alcohol. Add 10 mL to
100 mL reagent-grade water.
d) Acetone alcohol: Mix equal volumes of ethyl alcohol (95%) with acetone.
b. Procedure:
1) Using aseptic technique, streak one LES Endo agar (Section 9222B.2) or MacConkey agar
plate from each tube of brilliant green lactose bile broth showing gas, as soon as possible after
the observation of gas. Streak plates in a manner to insure presence
of some discrete colonies separated by at least 0.5 cm. Observe the following precautions
when streaking plates to obtain a high proportion of successful isolations if coliform organisms
are present: (a) Use a sterile 3-mm-diam loop or an inoculating needle slightly curved at the tip;
(b) tap and incline the fermentation tube to avoid picking up any membrane or scum on the
needle; (c) insert end of loop or needle into the liquid in the tube to a depth of approximately 0.5
cm; and (d) streak plate for isolation with curved section of the needle in contact with the agar to
avoid a scratched or torn surface. Flame loop between second and third quadrants to improve
colony isolation.
Incubate plates (inverted) at 35 ± 0.5°C for 24 ± 2 h.
2) The colonies developing on LES Endo agar are defined as typical (pink to dark red with a
green metallic surface sheen) or atypical (pink, red, white, or colorless colonies without sheen)
after 24 h incubation. Typical lactose-fermenting colonies developing on MacConkey agar are
red and may be surrounded by an opaque zone of precipitated bile. From each plate pick one or
more typical, well-isolated coliform colonies or, if no typical colonies are present, pick two or
more colonies considered most likely to consist of organisms of the coliform group, and transfer
growth from each isolate to a single-strength lauryl tryptose broth fermentation tube and onto a
nutrient agar slant. (The latter is unnecessary for drinking water samples.)
If needed, use a colony magnifying device to provide optimum magnification when colonies
are picked from the LES Endo or MacConkey agar plates. When transferring colonies, choose
well-isolated ones and barely touch the surface of the colony with a flame-sterilized, air-cooled
transfer needle to minimize the danger of transferring a mixed culture.
Incubate secondary broth tubes (lauryl tryptose broth with inverted fermentation vials
inserted) at 35 ± 0.5°C for 24 ± 2 h; if gas is not produced within 24 ± 2 h reincubate and
examine again at 48 ± 3 h. Microscopically examine Gram-stained preparations from those 24-h
nutrient agar slant cultures corresponding to the secondary tubes that show gas.
3) Gram-stain technique—The Gram stain may be omitted from the completed test for
potable water samples only because the occurrences of gram-positive bacteria and spore-forming
organisms surviving this selective screening procedure are infrequent in drinking water.
Various modifications of the Gram stain technique exist. Use the following modification by
Hucker for staining smears of pure culture; include a gram-positive and a gram-negative culture
as controls.
Prepare separate light emulsions of the test bacterial growth and positive and negative
control cultures on the same slide using drops of distilled water on the slide. Air-dry and fix by
passing slide through a flame and stain for 1 min with ammonium oxalate-crystal violet solution.
Rinse slide in tap water and drain off excess; apply Lugol’s solution for 1 min.
Rinse stained slide in tap water. Decolorize for approximately 15 to 30 s with acetone
alcohol by holding slide between the fingers and letting acetone alcohol flow across the stained
smear until the solvent flows colorlessly from the slide. Do not over-decolorize. Counterstain
with safranin for 15 s, rinse with tap water, blot dry with absorbent paper or air dry, and examine
microscopically. Gram-positive organisms are blue; gram-negative organisms are red. Results
are acceptable only when controls have given proper reactions.
c. Interpretation: Formation of gas in the secondary tube of lauryl tryptose broth within 48 ±
3 h and demonstration of gram-negative, nonspore-forming, rod-shaped bacteria from the agar
culture constitute a positive result for the completed test, demonstrating the presence of a
member of the coliform group.
4. Bibliography
MEYER, E.M. 1918. An aerobic spore-forming bacillus giving gas in lactose broth isolated in
routine water examination. J. Bacteriol. 3:9.
HUCKER, G.J. & H.J. CONN. 1923. Methods of Gram Staining. N.Y. State Agr. Exp. Sta. Tech.
Bull. No. 93.
NORTON, J.F. & J.J. WEIGHT. 1924. Aerobic spore-forming lactose fermenting organisms and
their significance in water analysis. Amer. J. Pub. Health 14:1019.
HUCKER, G.J. & H.J. CONN. 1927. Further Studies on the Methods of Gram Staining. N.Y. State
Agr. Exp. Sta. Tech. Bull. No. 128.
PORTER, R., C.S. MCCLESKEY & M. LEVINE. 1937. The facultative sporulating bacteria producing
gas from lactose. J. Bacteriol. 33:163.
COWLES, P.B. 1939. A modified fermentation tube. J. Bacteriol. 38:677.
SHERMAN, V.B.D. 1967. A Guide to the Identification of the Genera of Bacteria. Williams &
Wilkins, Baltimore, Md.
GELDREICH, E.E. 1975. Handbook for Evaluating Water Bacteriological Laboratories, 2nd ed.
EVANS, T.M., C.E. WAARVICK, R.J. SEIDLER & M.W. LECHEVALLIER. 1981. Failure of the
most-probable number technique to detect coliforms in drinking water and raw water
supplies. Appl. Environ. Microbiol. 41:130.
SEIDLER, R.J., T.M. EVANS, J.R. KAUFMAN, C.E. WAARVICK & M.W. LECHEVALLIER. 1981.
Limitations of standard coliform enumeration techniques. J. Amer. Water Works Assoc.
73:538.
GERHARDS, P., ed. 1981. Manual of Methods for General Bacteriology. American Soc.
Microbiology, Washington, D.C.
KRIEG, N.R. & J.G. HOLT, eds. 1984. Bergey’s Manual of Systematic Bacteriology, Vol 1.
Williams & Wilkins, Baltimore, Md.
GREENBERG, A.E. & D.A. HUNT, eds. 1985. Laboratory Procedures for the Examination of
Seawater and Shellfish, 5th ed. American Public Health Assoc., Washington, D.C.
U.S. ENVIRONMENTAL PROTECTION AGENCY. 1989. National primary drinking water
regulations: analytical techniques; coliform bacteria; final rule. Federal Register
54(135):29998 (July 17, 1989).
9221 C.
Estimation of Bacterial Density
1. Precision of Fermentation Tube Test
Unless a large number of sample portions is examined, the precision of the fermentation
tube test is rather low. For example, if only 1 mL is examined in a sample containing 1 coliform
organism/mL, about 37% of 1-mL tubes may be expected to yield negative results because of
random distribution of the bacteria in the sample. When five tubes, each with 1 mL sample, are
used under these conditions, a completely negative result may be expected less than 1% of the
time.
Consequently, exercise great caution when interpreting the sanitary significance of coliform
results obtained from the use of a few tubes with each sample dilution, especially when the
number of samples from a given sampling point is limited.
2. Computing and Recording of MPN
To calculate coliform density, compute in terms of the Most Probable Number (MPN). The
MPN values, for a variety of planting series and results, are given in Table 9221:II, Table
9221:III, and Table 9221:IV. Included in these tables are the 95% confidence limits for each
MPN value determined. If the sample volumes used are those found in the tables, report the
value corresponding to the number of positive and negative results in the series as the MPN/100
mL or report as total or fecal coliform presence or absence.
The sample volumes indicated in Table 9221:II and Table 9221:III relate more specifically
to finished waters. Table 9221:IV illustrates MPN values for combinations of positive and
negative results when five 10-mL, five 1.0-mL, and five 0.1-mL volumes of samples are tested.
When the series of decimal dilutions is different from that in the table, select the MPN value
from Table 9221:IV for the combination of positive tubes and calculate according to the
following formula:
When more than three dilutions are used in a decimal series of dilutions, use the results from
only three of these in computing the MPN. To select the three dilutions to be used in determining
the MPN index, choose the highest dilution that gives positive results in all five portions tested
(no lower dilution giving any negative results) and the two next succeeding higher dilutions. Use
the results at these three volumes in computing the MPN index. In the examples given below, the
significant dilution results are shown in boldface. The number in the numerator represents
positive tubes; that in the denominator, the total tubes planted; the combination of positives
simply represents the total number of positive tubes per dilution:
Example
a
b
c
1
mL
0.1
mL
0.01
mL
0.001
mL
Combination
of positives
MPN Index
/100 mL
5/5
5/5
4/5
1/5
2/5
2/5
0/5
0/5
0/5
0/5
5-2-0
5-4-2
0-1-0
5000
2200
20
5/5
0/5
In c, select the first three dilutions so as to include the positive result in the middle dilution.
When a case such as that shown below in line d arises, where a positive occurs in a dilution
higher than the three chosen according to the rule, incorporate it in the result for the highest
chosen dilution, as in e:
Example
1
mL
0.1
mL
0.01
mL
0.001
mL
Combination
of positives
MPN Index
/100 mL
d
e
5/5
5/5
3/5
3/5
1/5
2/5
1/5
0/5
5-3-2
5-3-2
1400
1400
Example
1
mL
0.1
mL
0.01
mL
0.001
mL
Combination
of positives
MPN Index
/100 mL
When it is desired to summarize with a single MPN value the results from a series of
samples, use the geometric mean or the median.
Table 9221:IV shows the most likely positive tube combinations. If unlikely combinations
occur with a frequency greater than 1% it is an indication that the technique is faulty or that the
statistical assumptions underlying the MPN estimate are not being fulfilled. The MPN for
combinations not appearing in the table, or for other combinations of tubes or dilutions, may be
estimated by Thomas’ simple formula:
While the MPN tables and calculations are described for use in the coliform test, they are
equally applicable to determining the MPN of any other organisms provided that suitable test
media are available.
3. Bibliography
MCCRADY, M.H. 1915. The numerical interpretation of fermentation tube results. J. Infect. Dis.
12:183.
MCCRADY, M.H. 1918. Tables for rapid interpretation of fermentation-tube results. Pub. Health
J. 9:201.
HOSKINS, J.K. 1933. The most probable numbers of B. coli in water analysis. J. Amer. Water
Works Assoc. 25:867.
HOSKINS, J.K. 1934. Most Probable Numbers for evaluation of coli-aerogenes tests by
fermentation tube method. Pub. Health Rep. 49:393.
HOSKINS, J.K. & C.T. BUTTERFIELD. 1935. Determining the bacteriological quality of drinking
water. J. Amer. Water Works Assoc. 27:1101.
HALVORSON, H.O. & N.R. ZIEGLER. 1933–35. Application of statistics to problems in
bacteriology. J. Bacteriol. 25:101; 26:331,559; 29:609.
SWAROOP, S. 1938. Numerical estimation of B. coli by dilution method. Indian J. Med. Res.
26:353.
DALLA VALLE, J.M. 1941. Notes on the most probable number index as used in bacteriology.
Pub. Health Rep. 56:229.
THOMAS, H.A., JR. 1942. Bacterial densities from fermentation tube tests. J. Amer. Water Works
Assoc. 34:572.
WOODWARD, R.L. 1957. How probable is the Most Probable Number? J. Amer. Water Works
Assoc. 49:1060.
MCCARTHY, J.A., H.A. THOMAS, JR. & J.E. DELANEY. 1958. Evaluation of the reliability of
coliform density tests. Amer. J. Pub. Health 48: 1628.
U.S. ENVIRONMENTAL PROTECTION AGENCY. 1989. National primary drinking water
regulations: analytical techniques; coliform bacteria; final rule. Federal Register
54(135):29998 (July 17, 1989).
DE MAN, J.C. 1977. MPN tables for more than one test. European J. Appl. Microbiol. 4:307.
9221 D.
Presence-Absence (P-A) Coliform Test
The presence-absence (P-A) test for the coliform group is a simple modification of the
multiple-tube procedure. Simplification, by use of one large test portion (100 mL) in a single
culture bottle to obtain qualitative information on the presence or absence of coliforms, is
justified on the theory that no coliforms should be present in 100 mL of a drinking water sample.
The P-A test also provides the optional opportunity for further screening of the culture to isolate
other indicators (fecal coliform, Aeromonas, Staphylococcus, Pseudomonas, fecal streptococcus,
and Clostridium) on the same qualitative basis. Additional advantages include the possibility of
examining a larger number of samples per unit of time. Comparative studies with the membrane
filter procedure indicate that the P-A test may maximize coliform detection in samples
containing many organisms that could overgrow coliform colonies and cause problems in
detection.
The P-A test is intended for use on routine samples collected from distribution systems or
water treatment plants. When sample locations produce a positive P-A result for coliforms, it
may be advisable to determine coliform densities in repeat samples. Quantitative information
may indicate the magnitude of a contaminating event.
1. Presumptive Phase
a. Culture media:
1) P-A broth: This medium is commercially available in dehydrated and in sterile
concentrated form.
Beef extract
Peptone
Lactose
Tryptose
3.0
5.0
7.46
9.83
1.35
g
g
g
g
g
1.35
g
Bromcresol purple
Reagent-grade water
2.46
0.05
0.0085
1
g
g
g
L
Make this formulation triple (3×) strength when examining 100-mL samples. Dissolve the
P-A broth medium in water without heating, using a stirring device. Dispense 50 mL prepared
medium into a screw-cap 250-mL milk dilution bottle. A fermentation tube insert is not
necessary. Autoclave for 12 min at 121°C with the total time in the autoclave limited to 30 min
or less. pH should be 6.8 ± 0.2 after sterilization. When the PA medium is sterilized by filtration
a 6× strength medium may be used. Aseptically dispense 20 mL of the 6× medium into a sterile
250-mL dilution bottle or equivalent container.
2) Lauryl tryptose broth: See Section 9221B.1.
b. Procedure: Shake sample vigorously for 5 s (approximately 25 times) and inoculate 100
mL into a P-A culture bottle. Mix thoroughly by inverting bottle once or twice to achieve even
distribution of the triple-strength medium throughout the sample. Incubate at 35 ± 0.5°C and
inspect after 24 and 48 h for acid reactions.
c. Interpretation: A distinct yellow color forms in the medium when acid conditions exist
following lactose fermentation. If gas also is being produced, gently shaking the bottle will result
in a foaming reaction. Any amount of gas and/or acid constitutes a positive presumptive test
requiring confirmation.
2. Confirmed Phase
The confirmed phase is outlined in Figure 9221:1.
a. Culture medium: Use brilliant green lactose bile fermentation tubes (see Section 9221B.2).
b. Procedure: Transfer all cultures that show acid reaction or acid and gas reaction to
brilliant green lactose bile (BGLB) broth for incubation at 35 ± 0.5°C (see Section 9221B.2).
c. Interpretation: Gas production in the BGLB broth culture within 48 ± 3 h confirms the
presence of coliform bacteria. Report result as presence-absence test positive or negative for total
coliforms in 100 mL of sample.
3. Completed Phase
The completed phase is outlined in Section 9221B.3 and Figure 9221:1.
4. Bibliography
WEISS, J.E. & C.A. HUNTER. 1939. Simplified bacteriological examination of water. J. Amer.
Water Works Assoc. 31:707.
CLARK, J.A. 1969. The detection of various bacteria indicative of water pollution by a
presence-absence (P-A) procedure. Can. J. Microbiol. 15:771.
CLARK, J.A. & L.T. VLASSOFF. 1973. Relationships among pollution indicator bacteria isolated
from raw water and distribution systems by the presence-absence (P-A) test. Health Lab. Sci.
10:163.
CLARK, J.A. 1980. The influence of increasing numbers of nonindicator organisms upon the
detection of indicator organisms by the membrane filter and presence-absence tests. Can. J.
Microbiol. 26: 827.
CLARK, J.A., C.A. BURGER & L.E. SABATINOS. 1982. Characterization of indicator bacteria in
municipal raw water, drinking water and new main water samples. Can. J. Microbiol.
28:1002.
JACOBS, N.J., W.L. ZEIGLER, F.C. REED, T.A. STUKEL & E.W. RICE. 1986. Comparison of membrane
filter, multiple-fermentation-tube, and presence-absence techniques for detecting total
coliforms in small community water systems. Appl. Environ. Microbiol. 51:1007.
RICE, E.W., E.E. GELDREICH & E.J. READ. 1989. The presence-absence coliform test for
monitoring drinking water quality. Pub. Health Rep. 104:54.
9221 E.
Fecal Coliform Procedure
Elevated-temperature tests for distinguishing organisms of the total coliform group that also
belong to the fecal coliform group are described herein. Modifications in technical procedures,
standardization of methods, and detailed studies of the fecal coliform group have established the
value of this procedure. The test can be performed by one of the multiple-tube procedures
described here or by membrane filter methods as described in Section 9222. The procedure using
A-1 broth is a single-step method.
The fecal coliform test (using EC medium) is applicable to investigations of drinking water,
stream pollution, raw water sources, wastewater treatment systems, bathing waters, seawaters,
and general water-quality monitoring. Prior enrichment in presumptive media is required for
optimum recovery of fecal coliforms when using EC medium. The test using A-1 medium is
applicable to source water, seawater, and treated wastewater.
1. Fecal Coliform Test (EC Medium)
The fecal coliform test is used to distinguish those total coliform organisms that are fecal
coliforms. Use EC medium or, for a more rapid test of the quality of shellfish waters, treated
wastewaters, or source waters, use A-1 medium in a direct test.
a. EC medium:
Tryptose or trypticase
Lactose
Bile salts mixture or bile salts No. 3
20.0
5.0
1.5
4.0
g
g
g
g
1.5 g
Reagent-grade water
5.0 g
1 L
± 0.2 after sterilization. Before sterilization, dispense in fermentation tubes, each with an
inverted vial, sufficient medium to cover the inverted vial at least partially after sterilization.
Close tubes with metal or heat-resistant plastic caps.
b. Procedure: Submit all presumptive fermentation tubes or bottles showing any amount of
gas, growth, or acidity within 48 h of incubation to the fecal coliform test.
1) Gently shake or rotate presumptive fermentation tubes or bottles showing gas, growth, or
acidity. Using a sterile 3- or 3.5-mm-diam loop or sterile wooden applicator stick, transfer
growth from each presumptive fermentation tube or bottle to EC broth (see Section 9221B.2).
2) Incubate inoculated EC broth tubes in a water bath at 44.5 ± 0.2°C for 24 ± 2 h.
Place all EC tubes in water bath within 30 min after inoculation. Maintain a sufficient water
depth in water bath incubator to immerse tubes to upper level of the medium.
c. Interpretation: Gas production with growth in an EC broth culture within 24 ± 2 h or less
is considered a positive fecal coliform reaction. Failure to produce gas (with little or no growth)
constitutes a negative reaction. If multiple tubes are used, calculate MPN from the number of
positive EC broth tubes as described in Section 9221C. When using only one tube for
subculturing from a single presumptive bottle, report as presence or absence of fecal coliforms.
2. Fecal Coliform Direct Test (A-1 Medium)
a. A-1 broth: This medium may be used for the direct isolation of fecal coliforms from water.
Prior enrichment in a presumptive medium is not required.
Lactose
Tryptone
Salicin
Polyethylene glycol p-isooctylphenyl ether*#(35)
Reagent-grade water
5.0
20.0
5.0
0.5
1.0
1
g
g
g
g
mL
L
Heat to dissolve solid ingredients, add polyethylene glycol p-isooctylphenyl ether, and adjust
to pH 6.9 ± 0.1. Before sterilization dispense in fermentation tubes with an inverted vial
sufficient medium to cover the inverted vial at least partially after sterilization. Close with metal
or heat-resistant plastic caps. Sterilize by autoclaving at 121°C for 10 min. Store in dark at room
temperature for not longer than 7 d. Ignore formation of precipitate.
Make A-1 broth of such strength that adding 10-mL sample portions to medium will not
reduce ingredient concentrations below those of the standard medium. For 10-mL samples
prepare double-strength medium.
b. Procedure: Inoculate tubes of A-1 broth as directed in Section 9221B.1b1). Incubate for 3
h at 35 ± 0.5°C. Transfer tubes to a water bath at 44.5 ± 0.2°C and incubate for an additional 21
± 2 h.
c. Interpretation: Gas production in any A-1 broth culture within 24 h or less is a positive
reaction indicating the presence of fecal coliforms. Calculate MPN from the number of positive
A-1 broth tubes as described in Section 9221C.
3. Bibliography
PERRY, C.A. & A.A. HAJNA. 1933. A modified Eijkman medium. J. Bacteriol. 26:419.
PERRY, C.A. & A.A. HAJNA. 1944. Further evaluation of EC medium for the isolation of coliform
bacteria and Escherichia coli. Amer. J. Pub. Health 34:735.
GELDREICH, E.E., H.F. CLARK, P.W. KABLER, C.B. HUFF & R.H. BORDNER. 1958. The coliform
group. II. Reactions in EC medium at 45°C. Appl. Microbiol. 6:347.
GELDREICH, E.E., R.H. BORDNER, C.B. HUFF, H.F. CLARK & P.W. KABLER. 1962. Type distribution
of coliform bacteria in the feces of warm-blooded animals. J. Water Pollut. Control Fed.
34:295.
GELDREICH, E.E. 1966. Sanitary significance of fecal coliforms in the environment. FWPCA
Publ. WP-20-3 (Nov.). U.S. Dep. Interior, Washington, D.C.
ANDREWS, W.H. & M.W. PRESNELL. 1972. Rapid recovery of Escherichia coli from estuarine
water. Appl. Microbiol. 23:521.
OLSON, B.H. 1978. Enhanced accuracy of coliform testing in seawater by a modification of the
most-probable-number method. Appl. Microbiol. 36:438.
STRANDRIDGE, J.H. & J.J. DELFINO. 1981. A-1 Medium: Alternative technique for fecal coliform
organism enumeration in chlorinated wastewaters. Appl. Environ. Microbiol. 42:918.
9221 F.
Escherichia coli Procedure (PROPOSED)
Escherichia coli is a member of the fecal coliform group of bacteria. This organism in water
indicates fecal contamination. Enzymatic assays have been developed that allow for the
identification of this organism. In this method E. coli are defined as coliform bacteria that
possess the enzyme β-glucuronidase and are capable of cleaving the fluorogenic substrate
4-methylumbelliferyl-β-D-glucuronide (MUG) with the corresponding release of the fluorogen
when grown in EC-MUG medium at 44.5°C within 24 ± 2 h or less. The procedure is used as a
confirmatory test after prior enrichment in a presumptive medium for total coliform bacteria.
This test is performed as a tube procedure as described here or by the membrane filter method as
described in Section 9222. The chromogenic substrate procedure (Section 9223) can be used for
direct detection of E. coli.
Tests for E. coli (using EC-MUG medium) are applicable for the analysis of drinking water,
surface and ground water, and wastewater. E. coli is a member of the indigenous fecal flora of
warm-blooded animals. The occurrence of E. coli is considered a specific indicator of fecal
contamination and the possible presence of enteric pathogens.
1. Escherichia coli Test (EC-MUG medium)
Use EC-MUG medium for the confirmation of E. coli.
a. EC-MUG medium:
Lactose
20.0
5.0
1.5
4.0
g
g
g
g
1.5 g
5.0 g
0.05 g
4-methylumbelliferyl-β-D-glucuronide (MUG)
Reagent-grade water
1
L
± 0.2 after sterilization. Before sterilization, dispense in tubes that do not fluoresce under
long-wavelength (366 nm) ultraviolet (UV) light. An inverted tube is not necessary. Close tubes
with metal or heat-resistant plastic caps.
b. Procedure: Submit all presumptive fermentation tubes or bottles showing growth, gas, or
acidity within 48 ± 3 h of incubation to the E. coli test.
1) Gently shake or rotate presumptive fermentation tubes or bottles showing growth, gas, or
acidity. Using a sterile 3- or 3.5-mm-diam metal loop or sterile wooden applicator stick, transfer
growth from presumptive fermentation tube or bottle to EC-MUG broth.
2) Incubate inoculated EC-MUG tubes in a water bath or incubator maintained at 44.5 ±
0.2°C for 24 ± 2 h. Place all EC-MUG tubes in water bath within 30 min after inoculation.
Maintain a sufficient water depth in the water-bath incubator to immerse tubes to upper level of
medium.
c. Interpretation: Examine all tubes exhibiting growth for fluorescence using a
long-wavelength UV lamp (preferably 6 W). The presence of bright blue fluorescence is
considered a positive response for E. coli. A positive control consisting of a known E. coli
(MUG-positive) culture, a negative control consisting of a thermotolerant Klebsiella pneumoniae
(MUG-negative) culture, and an uninoculated medium control may be necessary to interpret the
results and to avoid confusion of weak auto-fluorescence of the medium as a positive response.
If multiple tubes are used, calculate MPN from the number of positive EC-MUG broth tubes as
described in Section 9221C. When using only one tube or subculturing from a single
presumptive bottle, report as presence or absence of E. coli.
2. Bibliography
FENG, P.C.S. & P.A. HARTMAN. 1982. Fluorogenic assays for immediate confirmation of
Escherichia coli. Appl. Environ. Microbiol. 43:1320.
HARTMAN, P.A. 1989. The MUG (glucuronidase) test for E. coli in food and water. In A. Balows
et al., eds., Rapid Methods and Automation in Microbiology and Immunology. Proc. 5th Intl.
Symp. on Rapid Methods and Automation in Microbiology & Immunology, Florence, Italy,
Nov. 4–6, 1987.
SHADIX, L.C. & E.W. RICE. 1991. Evaluation of β-glucuronidase assay for the detection of
Escherichia coli from environmental waters. Can. J. Microbiol. 37:908.
9222
MEMBRANE FILTER TECHNIQUE FOR MEMBERS OF THE COLIFORM
GROUP*#(36)
9222 A.
Introduction
The membrane filter (MF) technique is highly reproducible, can be used to test relatively
large sample volumes, and usually yields numerical results more rapidly than the multiple-tube
fermentation procedure. The MF technique is extremely useful in monitoring drinking water and
a variety of natural waters. However, the MF technique has limitations, particularly when testing
waters with high turbidity or large numbers of noncoliform (background) bacteria. When the MF
technique has not been used previously, it is desirable to conduct parallel tests with the method
the laboratory is using currently to demonstrate applicability and comparability.
1. Definition
As related to the MF technique, the coliform group is defined as those facultative anaerobic,
gram-negative, non-spore-forming, rod-shaped bacteria that develop red colonies with a metallic
(golden) sheen within 24 h at 35°C on an Endo-type medium containing lactose. Some members
of the total coliform group may produce dark red, mucoid, or nucleated colonies without a
metallic sheen. When verified these are classified as atypical coliform colonies. When purified
cultures of coliform bacteria are tested, they produce negative cytochrome oxidase and positive
β-galactosidase test reactions.†#(37) Generally, pink (non-mucoid), blue, white, or colorless
colonies lacking sheen are considered noncoliforms by this technique.
2. Applications
Turbidity caused by the presence of algae, particulates, or other interfering material may not
permit testing of a sample volume sufficient to yield significant results. Low coliform estimates
may be caused by the presence of high numbers of noncoliforms or of toxic substances. The MF
technique is applicable to the examination of saline waters, but not wastewaters that have
received only primary treatment followed by chlorination because of turbidity in high volume
samples or wastewaters containing toxic metals or toxic organic compounds such as phenols. For
the detection of stressed total coliforms in treated drinking water and chlorinated secondary or
tertiary wastewater effluents, use a method designed for stressed organism recovery (see Section
9212B.1). A modified MF technique for fecal coliforms (Section 9212) in chlorinated
wastewater may be used if parallel testing over a 3-month period with the multiple-tube
fermentation technique shows comparability for each site-specific type of sample.
The standard volume to be filtered for drinking water samples is 100 mL. This may be
distributed among multiple membranes if necessary. However, for special monitoring purposes,
such as troubleshooting water quality problems or identification of coliform breakthrough in low
concentrations from treatment barriers, it may be desirable to test 1-L samples. If particulates
prevent filtering a 1-L sample through a single filter, divide sample into four portions of 250 mL
for analysis. Total the coliform counts on each membrane to report the number of coliforms per
liter. Smaller sample volumes will be necessary for source or recreational waters and wastewater
effluents that have much higher coliform densities.
Statistical comparisons of results obtained by the multiple-tube method and the MF
technique show that the MF is more precise (compare Table 9221:II and Table 9221:III with
Table 9222:II). Data from each test yield approximately the same water quality information,
although numerical results are not identical.
3. Bibliography
KABLER, P.W. 1954. Water examinations by membrane filter and MPN procedures. Amer. J.
Pub. Health 44:379.
THOMAS, H.A. & R.L. WOODWARD. 1956. Use of molecular filter membranes for water potability
control. J. Amer. Water Works Assoc. 48: 1391.
MCCARTHY, J.A., J.E. DELANEY & R.J. GRASSO. 1961. Measuring coliforms in water. Water
Sewage Works 108:238.
LIN, S. 1973. Evaluation of coliform test for chlorinated secondary effluents. J. Water Pollut.
Control Fed. 45:498.
MANDEL, J. & L.F. NANNI. 1978. Measurement evaluation. In S.L. Inhorn, ed. Quality Assurance
Practices for Health Laboratories, p. 209. American Public Health Assoc., Washington, D.C.
9222 B.
Standard Total Coliform Membrane Filter Procedure
For MF analyses use glassware and other apparatus composed of material free from agents
that may affect bacterial growth.
a. Sample bottles: See Section 9030B.18.
b. Dilution bottles: See Section 9030B.13.
c. Pipets and graduated cylinders: See Section 9030B.9. Before sterilization, loosely cover
opening of graduated cylinders with metal foil or a suitable heavy wrapping-paper substitute.
Immediately after sterilization secure cover to prevent contamination.
d. Containers for culture medium: Use clean borosilicate glass flasks. Any size or shape of
flask may be used, but erlenmeyer flasks with metal caps, metal foil covers, or screw caps
provide for adequate mixing of the medium contained and are convenient for storage.
e. Culture dishes: Use sterile borosilicate glass or disposable, presterilized plastic petri
dishes, 60 × 15 mm, 50 × 9 mm, or other appropriate size. Wrap convenient numbers of clean,
glass culture dishes in metal foil if sterilized by dry heat, or suitable heavy wrapping paper when
autoclaved. Incubate loose-lidded glass and disposable plastic culture dishes in tightly closed
containers with wet paper or cloth to prevent moisture evaporation with resultant drying of
medium and to maintain a humid environment for optimum colony development.
Presterilized disposable plastic dishes with tight-fitting lids that meet the specifications
above are available commercially and are used widely. Reseal opened packages of disposable
dish supplies for storage.
f. Filtration units: The filter-holding assembly (constructed of glass, autoclavable plastic,
porcelain, or stainless steel) consists of a seamless funnel fastened to a base by a locking device
or by magnetic force. The design should permit the membrane filter to be held securely on the
porous plate of the receptacle without mechanical damage and allow all fluid to pass through the
membrane during filtration. Discard plastic funnels with deep scratches on inner surface or glass
funnels with chipped surfaces.
Wrap the assembly (as a whole or separate parts) in heavy wrapping paper or aluminum foil,
sterilize by autoclaving, and store until use. Alternatively expose all surfaces of the previously
cleaned assembly to ultraviolet radiation (2 min exposure) for the initial sanitization before use
in the test procedure, or before reusing units between successive filtration series. Field units may
be sanitized by dipping or spraying with alcohol and then igniting or immersing in boiling water
for 2 min. After submerging unit in boiling water, cool it to room temperature before reuse. Do
not ignite plastic parts. Sterile, disposable field units may be used.
For filtration, mount receptacle of filter-holding assembly on a 1-L filtering flask with a side
tube or other suitable device (manifold to hold three to six filter assemblies) such that a pressure
differential (34 to 51 kPa) can be exerted on the filter membrane. Connect flask to a vacuum line,
an electric vacuum pump, a filter pump operating on water pressure, a hand aspirator, or other
means of securing a pressure differential (138 to 207 kPa). Connect a flask of approximately the
same capacity between filtering flask and vacuum source to trap carry-over water.
g. Membrane filter: Use membrane filters (for additional specifications, see Section 9020)
with a rated pore diameter such that there is complete retention of coliform bacteria. Use only
those filter membranes that have been found, through adequate quality control testing and
certification by the manufacturer, to exhibit: full retention of the organisms to be cultivated,
stability in use, freedom from chemical extractables that may inhibit bacterial growth and
development, a satisfactory speed of filtration (within 5 min), no significant influence on
medium pH (beyond ± 0.2 units), and no increase in number of confluent colonies or spreaders
compared to control membrane filters. Use membranes grid-marked in such a manner that
bacterial growth is neither inhibited nor stimulated along the grid lines when the membranes
with entrapped bacteria are incubated on a suitable medium. Preferably use fresh stocks of
membrane filters and if necessary store them in an environment without extremes of temperature
and humidity. Obtain no more than a year’s supply at any one time.
Preferably use presterilized membrane filters for which the manufacturer has certified that
the sterilization technique has neither induced toxicity nor altered the chemical or physical
properties of the membrane. If membranes are sterilized in the laboratory, autoclave for 10 min
at 121°C. At the end of the sterilization period, let the steam escape rapidly to minimize
accumulation of water of condensation on filters.
h. Absorbent pads consist of disks of filter paper or other material certified for each lot by
the manufacturer to be of high quality and free of sulfites or other substances of a concentration
that could inhibit bacterial growth. Use pads approximately 48 mm in diameter and of sufficient
thickness to absorb 1.8 to 2.2 mL of medium. Presterilized absorbent pads or pads subsequently
sterilized in the laboratory should release less than 1 mg total acidity (calculated as CaCO3)
when titrated to the phenolphthalein end point, pH 8.3, using 0.02N NaOH and produce pH
levels of 7 ± 0.2. Sterilize pads simultaneously with membrane filters available in resealable
kraft envelopes, or separately in other suitable containers. Dry pads so they are free of visible
moisture before use. See sterilization procedure described for membrane filters above and
Section 9020 for additional specifications on absorbent pads.
i. Forceps: Smooth flat forceps, without corrugations on the inner sides of the tips. Sterilize
before use by dipping in 95% ethyl or absolute methyl alcohol and flaming.
j. Incubators: Use incubators to provide a temperature of 35 ± 0.5°C and to maintain a
humid environment (60% relative humidity).
k. Microscope and light source: To determine colony counts on membrane filters, use a
magnification of 10 to 15 diameters and a cool white fluorescent light source adjusted to give
maximum sheen discernment. Optimally use a binocular wide-field dissecting microscope. Do
not use a microscope illuminator with optical system for light concentration from an
incandescent light source for discerning coliform colonies on Endo-type media.
The need for uniformity dictates the use of commercial dehydrated media. Never prepare
media from basic ingredients when suitable dehydrated media are available. Follow
manufacturer’s directions for rehydration. Store opened supplies of dehydrated media in a
desiccator. Commercially prepared media in liquid form (sterile ampule or other) may be used if
known to give equivalent results. See Section 9020 for media quality control specifications.
Test each new medium lot against a previously acceptable lot for satisfactory performance as
described in Section 9020B. With each new lot of Endo-type medium, verify a minimum 10% of
coliform colonies, obtained from natural samples or samples with known additions, to establish
the comparative recovery of the medium lot.
Before use, test each batch of laboratory-prepared MF medium for performance with positive
and negative culture controls. Check for coliform contamination at the beginning and end of each
filtration series by filtering 20 to 30 mL of dilution or rinse water through the filter. If controls
indicate contamination, reject all data from affected samples and request resample.
a. LES Endo agar:*#(38)
Yeast extract
Casitone or trypticase
Thiopeptone or thiotone
Tryptose
Lactose
1.2
3.7
3.7
7.5
9.4
3.3
g
g
g
g
g
g
1.0
g
Sodium sulfite, Na2SO3
3.7
0.1
0.05
1.6
g
g
g
g
Basic fuchsin
Agar
Reagent-grade water
0.8
15.0
1
g
g
L
Rehydrate product in 1 L water containing 20 mL 95% ethanol. Do not use denatured
ethanol, which reduces background growth and coliform colony size. Bring to a near boil to
dissolve agar, then promptly remove from heat and cool to 45 to 50°C. Do not sterilize by
autoclaving. Final pH 7.2 ± 0.2. Dispense in 5- to 7-mL quantities into lower section of 60-mm
glass or plastic petri dishes. If dishes of any other size are used, adjust quantity to give an
equivalent depth of 4 to 5 m. Do not expose poured plates to direct sunlight; refrigerate in the
dark, preferably in sealed plastic bags or other containers to reduce moisture loss. Discard
unused medium after 2 weeks or sooner if there is evidence of moisture loss, medium
contamination, or medium deterioration (darkening of the medium).
b. M-Endo medium:†#(39)
Tryptose or polypeptone
Thiopeptone or thiotone
Casitone or trypticase
Yeast extract
Lactose
10.0
5.0
5.0
1.5
12.5
5.0
4.375
g
g
g
g
g
g
g
1.375 g
Sodium sulfite, Na2SO3
0.05
0.10
2.10
g
g
g
1.05
15.0
1
g
g
L
Basic fuchsin
Agar (optional)
Reagent-grade water
1) Agar preparation—Rehydrate product in 1 L water containing 20 mL 95% ethanol. Heat
to near boiling to dissolve agar, then promptly remove from heat and cool to between 45 and
50°C. Dispense 5- to 7-mL quantities into 60-mm sterile glass or plastic petri dishes. If dishes of
any other size are used, adjust quantity to give an equivalent depth. Do not sterilize by
autoclaving. Final pH should be 7.2 ± 0.2. A precipitate is normal in Endo-type media.
Refrigerate finished medium in the dark and discard unused agar after 2 weeks.
2) Broth preparation—Prepare as above, omitting agar. Dispense liquid medium (at least 2.0
mL per plate) onto absorbent pads (see absorbent pad specifications, Section 9222B.1) and
carefully remove excess medium by decanting the plate. The broth may have a precipitate but
this does not interfere with medium performance if pads are certified free of sulfite or other toxic
agents at a concentration that could inhibit bacterial growth. Refrigerated broth may be stored for
up to 4 d.
c. Buffered dilution rinse water: See Section 9050C.1.
3. Samples
Collect samples as directed in Section 9060A and Section 9060B.
4. Coliform Definition
Bacteria that produce a red colony with a metallic (golden) sheen within 24 h incubation at
35°C on an Endo-type medium are considered members of the coliform group. The sheen may
cover the entire colony or may appear only in a central area or on the periphery. The coliform
group thus defined is based on the production of aldehydes from fermentation of lactose. While
this biochemical characteristic is part of the metabolic pathway of gas production in the
multiple-tube test, some variations in degree of metallic sheen development may be observed
among coliform strains. However, this slight difference in indicator definition is not considered
critical to change its public health significance, particularly if suitable studies have been
conducted to establish the relationship between results obtained by the MF and those obtained by
the standard multiple-tube fermentation procedure.
5. Procedures
a. Selection of sample size: Size of sample will be governed by expected bacterial density. In
drinking water analyses, sample size will be limited only by the degree of turbidity or by the
noncoliform growth on the medium (Table 9222:I). For regulation purposes, 100 mL is the
official sample size.
An ideal sample volume will yield 20 to 80 coliform colonies and not more than 200
colonies of all types on a membrane-filter surface. Analyze drinking waters by filtering 100 to
1000 mL, or by filtering replicate smaller sample volumes such as duplicate 50-mL or four
replicates of 25-mL portions. Analyze other waters by filtering three different volumes (diluted
or undiluted), depending on the expected bacterial density. See Section 9215B.2 for preparation
of dilutions. When less than 10 mL of sample (diluted or undiluted) is to be filtered, add
approximately 10 mL sterile dilution water to the funnel before filtration or pipet the sample
volume into a sterile dilution bottle, then filter the entire dilution. This increase in water volume
aids in uniform dispersion of the bacterial suspension over the entire effective filtering surface.
b. Sterile filtration units: Use sterile filtration units at the beginning of each filtration series
as a minimum precaution to avoid accidental contamination. A filtration series is considered to
be interrupted when an interval of 30 min or longer elapses between sample filtrations. After
such interruption, treat any further sample filtration as a new filtration series and sterilize all
membrane filter holders in use. See Section 9222B.1 f for sterilization procedures and Section
9020B.3m and n for UV cleaning and safety guidelines.
c. Filtration of sample: Using sterile forceps, place a sterile membrane filter (grid side up)
over porous plate of receptacle. Carefully place matched funnel unit over receptacle and lock it
in place. Filter sample under partial vacuum. With filter still in place, rinse the interior surface of
the funnel by filtering three 20- to 30-mL portions of sterile dilution water. Alternatively, rinse
funnel by a flow of sterile dilution water from a squeeze bottle. This is satisfactory only if the
squeeze bottle and its contents do not become contaminated during use. Rinsing between
samples prevents carryover contamination. Upon completion of final rinse and the filtration
process disengage vacuum, unlock and remove funnel, immediately remove membrane filter
with sterile forceps, and place it on selected medium with a rolling motion to avoid entrapment
of air. If the agar-based medium is used, place prepared filter directly on agar, invert dish, and
incubate for 22 to 24 h at 35 ± 0.5°C.
If liquid medium is used, place a pad in the culture dish and saturate with at least 2.0 mL
M-Endo medium and carefully remove excess medium by decanting the plate. Place prepared
filter directly on pad, invert dish, and incubate for 22 to 24 h at 35 ± 0.5°C.
Differentiation of some colonies from either agar or liquid medium substrates may be lost if
cultures are incubated beyond 24 h.
Insert a sterile rinse water sample (100 mL) after filtration of a series of 10 samples to check
for possible cross-contamination or contaminated rinse water. Incubate the rinse water control
membrane culture under the same conditions as the sample.
For nonpotable water samples, preferably decontaminate filter unit after each sample (as
described above) because of the high number of coliform bacteria present in these samples.
Alternatively, use an additional buffer rinse of the filter unit after the filter is removed to prevent
carryover between samples.
d. Alternative enrichment technique: Place a sterile absorbent pad in the lid of a sterile
culture dish and pipet at least 2.0 mL lauryl tryptose broth, prepared as directed in Section
9221B.1.a1), to saturate pad. Carefully remove any excess liquid from absorbent pad by
decanting plate. Aseptically place filter through which the sample has been passed on pad.
Incubate filter, without inverting dish, for 1.5 to 2 h at 35 ± 0.5°C in an atmosphere of at least
60% relative humidity.
If the agar-based Endo-type medium is used, remove enrichment culture from incubator, lift
filter from enrichment pad, and roll it onto the agar surface, which has been allowed to
equilibrate to room temperature. Incorrect filter placement is at once obvious, because patches of
unstained membrane indicate entrapment of air. Where such patches occur, carefully reseat filter
on agar surface. If the liquid medium is used, prepare final culture by removing enrichment
culture from incubator and separating the dish halves. Place a fresh sterile pad in bottom half of
dish and saturate with at least 2.0 mL of M-Endo medium and carefully remove excess liquid
from absorbent pad by decanting plate. Transfer filter, with same precautions as above, to new
pad. Discard used enrichment pad.
With either the agar or the liquid medium, invert dish and incubate for 20 to 22 h at 35 ±
0.5°C. Proceed to ¶ e below.
e. Counting: To determine colony counts on membrane filters, use a low-power (10 to 15
magnifications) binocular wide-field dissecting microscope or other optical device, with a cool
white fluorescent light source directed to provide optimal viewing of sheen. The typical coliform
colony has a pink to dark-red color with a metallic surface sheen. Count both typical and
atypical coliform colonies. The sheen area may vary in size from a small pinhead to complete
coverage of the colony surface. Atypical coliform colonies can be dark red, mucoid, or nucleated
without sheen. Generally pink, blue, white, or colorless colonies lacking sheen are considered
noncoliforms. The total count of colonies (coliform and noncoliform) on Endo-type medium has
no consistent relationship to the total number of bacteria present in the original sample. A high
count of noncoliform colonies may interfere with the maximum development of coliforms.
Refrigerating cultures (after 22 h incubation) with high densities of noncoliform colonies for 0.5
to 1 h before counting may deter spread of confluence while aiding sheen discernment.
Samples of disinfected water or wastewater effluent may include stressed organisms that
grow relatively slowly and produce maximum sheen in 22 to 24 h. Organisms from undisinfected
sources may produce sheen at 16 to 18 h, and the sheen subsequently may fade after 24 to 30 h.
f. Coliform verification: Occasionally, typical sheen colonies may be produced by
noncoliform organisms and atypical colonies (dark red or nucleated colonies without sheen) may
be coliforms. Preferably verify all typical and atypical colony types. For drinking water, verify
all suspect colonies by swabbing the entire membrane or pick at least five typical colonies and
five atypical colonies from a given membrane filter culture. For waters other than drinking
water, at a minimum, verify at least 10 sheen colonies (and representative atypical colonies of
different morphological types) from a positive water sample monthly. See Section 9020B.8.
Based on need and sample type, laboratories may incorporate more stringent quality control
measures (e.g., verify at least one colony from each typical or atypical colony type from a given
membrane filter culture, verify 10% of the positive samples). Adjust counts on the basis of
verification results. Verification tests are listed below.
1) Lactose fermentation—Transfer growth from each colony or swab the entire membrane
with a sterile cotton swab (for presence-absence results in drinking water samples) and place in
lauryl tryptose broth; incubate the lauryl tryptose broth at 35 ± 0.5°C for 48 h. Gas formed in
lauryl tryptose broth and confirmed in brilliant green lactose broth (Section 9221B.2 for medium
preparation) within 48 h verifies the colony as a coliform. Simultaneous inoculation of both
media for gas production is acceptable. Inclusion of EC broth inoculation for 44.5 ± 0.2°C
incubation will provide information on the presence of fecal coliforms. Use of EC-MUG with
incubation at 44.5 ± 0.2°C for 24 h will provide information on presence of E. coli. See Section
9222G for MF partition procedures.
2) Alternative coliform verifications—Apply this alternative coliform verification procedure
to isolated colonies on the membrane filter culture. If a mixed culture is suspected or if colony
separation is less than 2 mm, streak the growth to M-Endo medium or MacConkey agar to assure
culture purity or submit the mixed growth to the fermentation tube method.
a) Rapid test—A rapid verification of colonies utilizes test reactions for cytochrome oxidase
(CO) and β-galactosidase. Coliform reactions are CO negative and β-galactosidase positive
within 4 h incubation of tube culture or micro (spot) test procedure.
b) Commercial multi-test systems—Verify the colony by streaking it for purification,
selecting a well-isolated colony, and inoculating into a multi-test identification system for
Enterobacteriaceae that includes lactose fermentation and/or β-galactosidase and CO test
reactions.
6. Calculation of Coliform Density
Compute the count, using membrane filters with 20 to 80 coliform colonies and not more
than 200 colonies of all types per membrane, by the following equation:
If no coliform colonies are observed, report the coliform colonies counted as ‘‘<1
coliform/100 mL.’’
For verified coliform counts, adjust the initial count based upon the positive verification
percentage and report as ‘‘verified coliform count/100 mL.’’
a. Water of drinking water quality: While the EPA Total Coliform Rule for public water
supply samples requires only a record of coliform presence or absence in 100-mL samples, it
may be advisable to determine coliform densities in repeat sampling situations. This is of
particular importance when a coliform biofilm problem is suspected in the distribution system.
Quantitative information may provide an indication of the magnitude of a contaminating event.
With water of good quality, the occurrence of coliforms generally will be minimal.
Therefore, count all coliform colonies (disregarding the lower limit of 20 cited above) and use
the formula given above to obtain coliform density.
If confluent growth occurs, covering either the entire filtration area of the membrane or a
portion thereof, and colonies are not discrete, report results as ‘‘confluent growth with (or
without) coliforms.’’ If the total number of bacterial colonies, coliforms plus noncoliforms,
exceeds 200 per membrane, or if the colonies are not distinct enough for accurate counting,
report results as ‘‘too numerous to count’’ (TNTC) or ‘‘confluent,’’ respectively. For drinking
water, the presence of coliforms in such cultures showing no sheen may be confirmed by either
transferring a few colonies or placing the entire membrane filter culture into a sterile tube of
brilliant green lactose bile broth. As an alternative, brush the entire filter surface with a sterile
loop, applicator stick, or cotton swab and inoculate this growth to the tube of brilliant green
lactose bile broth. If gas is produced from the brilliant green bile broth tube within 48 h at 35 ±
0.5°C, coliforms are present. For compliance with the EPA Total Coliform Rule, report confluent
growth or TNTC with at least one detectable coliform colony (which is verified) as a total
coliform positive sample. Report confluent growth or TNTC without detectable coliforms as
invalid. For invalid samples, request a new sample from the same location within 24 h and select
more appropriate volumes to be filtered per membrane, observing the requirement that the
standard drinking water portion is 100 mL, or choose another coliform method that is less
subject to heterotrophic bacterial interferences. Thus, to reduce interference from overcrowding,
instead of filtering 100 mL per membrane, filter 50-mL portions through two separate
membranes, 25-mL portions through each of four membranes, etc. Total the coliform counts
observed on all membranes and report as number per 100 mL.
b. Water of other than drinking water quality: As with potable water samples, if no filter has
a coliform count falling in the ideal range, total the coliform counts on all filters and report as
number per 100 mL. For example, if duplicate 50-mL portions were examined and the two
membranes had five and three coliform colonies, respectively, report the count as eight coliform
colonies per 100 mL, i.e.,
Similarly, if 50-, 25-, and 10-mL portions were examined and the counts were 15, 6, and <1
coliform colonies, respectively, report the count as 25/100 mL, i.e.,
On the other hand, if 10-, 1.0-, and 0.1-mL portions were examined with counts of 40, 9, and
<1 coliform colonies, respectively, select the 10-mL portion only for calculating the coliform
density because this filter had a coliform count falling in the ideal range. The result is 400/100
mL, i.e.,
In this last example, if the membrane with 40 coliform colonies also had a total bacterial
colony count greater than 200, report the coliform count as ≥400/100 mL.
Report confluent growth or membranes with colonies too numerous to count as described in
a above. Request a new sample and select more appropriate volumes for filtration or utilize the
multiple-tube fermentation technique.
c. Statistical reliability of membrane filter results: Although the precision of the MF
technique is greater than that of the MPN procedure, membrane counts may underestimate the
number of viable coliform bacteria. Table 9222:II illustrates some 95% confidence limits. These
values are based on the assumption that bacteria are distributed randomly and follow a Poisson
distribution. For results with counts, c, greater than 20 organisms, calculate the approximate 95%
confidence limits using the following normal distribution equations:
7. Bibliography
FIFIELD, C.W. & C.P. SCHAUFUS. 1958. Improved membrane filter medium for the detection of
coliform organisms. J. Amer. Water Works Assoc. 50:193.
MCCARTHY, J.A. & J.E. DELANEY. 1958. Membrane filter media studies. Water Sewage Works
105:292.
RHINES, C.E. & W.P. CHEEVERS. 1965. Decontamination of membrane filter holders by ultraviolet
light. J. Amer. Water Works Assoc. 57: 500.
GELDREICH, E.E., H.L. JETER & J.A. WINTER. 1967. Technical considerations in applying the
membrane filter procedure. Health Lab. Sci. 4:113.
WATLING, H.R. & R.J. WATLING. 1975. Note on the trace metal content of membrane filters.
Water SA 1:28.
LIN, S.D. 1976. Evaluation of Millipore HA and HC membrane filters for the enumeration of
indicator bacteria. Appl. Environ. Microbiol. 32:300.
STANDRIDGE, J.H. 1976. Comparison of surface pore morphology of two brands of membrane
filters. Appl. Environ. Microbiol. 31:316.
GELDREICH, E.E. 1976. Performance variability of membrane filter procedure. Pub. Health Lab.
34:100.
GRABOW, W.O.K. & M. DU PREEZ. 1979. Comparison of m-Endo LES, MacConkey and Teepol
media for membrane filtration counting of total coliform bacteria in water. Appl. Environ.
Microbiol. 38:351.
DUTKA, B.D., ed. 1981. Membrane Filtration Applications, Techniques and Problems. Marcel
Dekker, Inc., New York, N.Y.
EVANS, T.M., R.J. SEIDLER & M.W. LECHEVALLIER. 1981. Impact of verification media and
resuscitation on accuracy of the membrane filter total coliform enumeration technique. Appl.
Environ. Microbiol. 41: 1144.
FRANZBLAU, S.G., B.J. HINNEBUSCH, T.M. KELLEY & N.A. SINCLAIR. 1984. Effect of noncoliforms
on coliform detection in potable groundwater: improved recovery with an anaerobic
membrane filter technique. Appl. Environ. Microbiol. 48:142.
MCFETERS, G.A., J.S. KIPPIN & M.W. LECHEVALLIER. 1986. Injured coliforms in drinking water.
9222 C.
Delayed-Incubation Total Coliform Procedure
Modification of the standard MF technique permits membrane shipment or transport after
filtration to a distant laboratory for transfer to another substrate, incubation, and completion of
the test. This delayed-incubation test may be used where it is impractical to apply conventional
procedures. It also may be used: (a) where it is not possible to maintain the desired sample
temperature during transport; (b) when the elapsed time between sample collection and analysis
would exceed the approved time limit; or (c) where the sampling location is remote from
laboratory services.
Independent studies using both fresh- and salt-water samples have shown consistent results
between the delayed incubation and standard direct test. Determine the applicability of the
delayed-incubation test for a specific water source by comparing with results of conventional
MF methods.
To conduct the delayed-incubation test, filter sample in the field immediately after collection,
place filter on the transport medium, and ship to the laboratory. Complete the coliform
determination in the laboratory by transferring the membrane to standard M-Endo or LES Endo
medium, incubating at 35 ± 0.5°C for 20 to 22 h, and counting typical and atypical coliform
colonies that develop. For drinking water samples collected for compliance with the EPA Total
Coliform Rule, report the presence or absence of verified coliforms in 100-mL samples. Verify
colonies as outlined previously in Section 9222B.5 f.
Transport media are designed to keep coliform organisms viable and generally do not permit
visible growth during transit time. Bacteriostatic agents in holding/preservative media suppress
growth of microorganisms en route but allow normal coliform growth after transfer to a fresh
medium.
The delayed-incubation test follows the methods outlined for the total coliform MF
procedure, except as indicated below. Two alternative methods are given, one using the M-Endo
preservative medium and the other the M-ST holding medium.
1. Apparatus
a. Culture dishes: Use disposable, sterile, plastic petri dishes (50 × 12 mm) with tight-fitting
lids. Such containers are light in weight and are less likely to break in transit. In an emergency or
when plastic dishes are unavailable, use sterile glass petri dishes wrapped in plastic film or
similar material. See Section 9222B.1e for specifications.
b. Field filtration units: See Section 9222B.1 f for specifications. Disinfect by adding methyl
alcohol to the filtering chamber, igniting the alcohol, and covering unit to produce formaldehyde.
Ultraviolet light disinfection also may be used in the field if an appropriate power source is
available (115 V, 60 Hz). Glass or metal filtration units may be sterilized by immersing in
boiling water for 2 min. Use a hand aspirator to obtain necessary vacuum.
2. Materials and Transport Media
a. M-Endo methods:
1) M-Endo preservative medium: Prepare M-Endo medium as described in Section
9222B.2b. After cooling to below 45°C, aseptically add 3.84 g sodium benzoate (USP grade)/L
or 3.2 mL 12% sodium benzoate solution to 100 mL medium. Mix ingredients and dispense in 5to 7-mL quantities to 50- × 9-mm petri plates. Refrigerate poured plates. Discard unused medium
after 96 h.
2) Sodium benzoate solution: Dissolve 12 g NaC7H5O2 in sufficient reagent water to make
100 mL. Sterilize by autoclaving or by filtering through a 0.22-µm pore size membrane filter.
Discard after 6 months.
3) Cycloheximide:*#(40) Optionally add cycloheximide to M-Endo preservative medium. It
may be used for samples that previously have shown overgrowth by fungi, including yeasts.
Prepare by aseptically adding 50 mg cycloheximide/100 mL to M-Endo preservative medium.
Store cycloheximide solution in refrigerator and discard after 6 months. Cycloheximide is a
powerful skin irritant; handle with caution according to the manufacturer’s directions.
b. M-ST method:
M-ST holding medium:
Sodium phosphate, monobasic, NaH2PO4⋅H2O
0.1 g
Dipotassium hydrogen phosphate, KH2PO4
3.0 g
Sulfanilamide
Ethanol (95%)
Tris (hydroxymethyl) aminomethane
Reagent-grade water
1.5
10
3.0
1
g
mL
g
L
Dissolve ingredients by rehydrating in water. Sterilize by autoclaving at 121°C for 15 min.
Final pH should be 8.6 ± 0.2. Dispense at least 2.0 mL to tight-lidded plastic culture dishes
containing an absorbent pad and carefully remove excess liquid from pad by decanting plate.
Store in refrigerator for use within 96 h.
3. Procedure
a. Sample preservation and shipment: Place absorbent pad in bottom of sterile petri dish and
saturate with selected coliform holding medium (see Section 9222C.2 above). Remove
membrane filter from filtration unit with sterile forceps and roll it, grid side up, onto surface of
medium-saturated pad. Protect membrane from moisture loss by tightly closing plastic petri dish.
Seal loose-fitting dishes with an appropriate sealing tape to prevent membrane dehydration
during transit. Place culture dish containing membrane in an appropriate shipping container and
send to the laboratory for test completion. The sample can be held without visible growth for a
maximum of 72 h on the holding/preservative medium. This usually allows use of the mail or a
common carrier. Visible growth occasionally begins on the transport medium when high
temperatures are encountered during transit.
b. Transfer and incubation: At the laboratory, transfer membrane from holding medium on
which it was shipped to a second sterile petri dish containing M-Endo or LES Endo medium and
incubate at 35 ± 0.5°C for 20 to 22 h.
4. Estimation of Coliform Density
Proceed as in Section 9222B.6 above. Record times of collection, filtration, and laboratory
examination, and calculate the elapsed time. Report elapsed time with coliform results.
5. Bibliography
GELDREICH, E.E., P.W. KABLER, H.L. JETER & H.F. CLARK. 1955. A delayed incubation membrane
filter test for coliform bacteria in water. Amer. J. Pub. Health 45:1462.
PANEZAI, A.K., T.J. MACKLIN & H.G. COLES. 1965. Coli-aerogenes and Escherichia coli counts on
water samples by means of transported membranes. Proc. Soc. Water Treat. Exam. 14:179.
BREZENSKI, F.T. & J.A. WINTER. 1969. Use of the delayed incubation membrane filter test for
determining coliform bacteria in sea water. Water Res. 3:583.
CHEN, M. & P.J. HICKEY. 1986. Elimination of overgrowth in delayed-incubation membrane filter
test for total coliforms by M-ST holding medium. Appl. Environ. Microbiol. 52:778.
9222 D.
Fecal Coliform Membrane Filter Procedure
Fecal coliform bacterial densities may be determined either by the multiple-tube procedure
or by the MF technique. See Section 9225 for differentiation of Escherichia coli, the
predominant fecal coliform. If the MF procedure is used for chlorinated effluents, demonstrate
that it gives comparable information to that obtainable by the multiple-tube test before accepting
it as an alternative. The fecal coliform MF procedure uses an enriched lactose medium and
incubation temperature of 44.5 ± 0.2°C for selectivity. Because incubation temperature is
critical, submerge waterproofed (plastic bag enclosures) MF cultures in a water bath for
incubation at the elevated temperature or use an appropriate solid heat sink incubator or other
incubator that is documented to hold the 44.5°C temperature within 0.2°C throughout the
chamber, over a 24-h period. Areas of application for the fecal coliform method in general are
stated in the introduction to the multiple-tube fecal coliform procedures, Section 9221E.
1. Materials and Culture Medium
a. M-FC medium: The need for uniformity dictates the use of dehydrated media. Never
prepare media from basic ingredients when suitable dehydrated media are available. Follow
manufacturer’s directions for rehydration. Commercially prepared media in liquid form (sterile
ampule or other) also may be used if known to give equivalent results. See Section 9020 for
quality control specifications.
M-FC medium:
Tryptose or biosate
10.0 g
Yeast extract
Lactose
Bile salts No. 3 or bile salts mixture
Aniline blue
Agar (optional)
Reagent-grade water
5.0
3.0
5.0
12.5
1.5
0.1
15.0
1
g
g
g
g
g
g
g
L
Rehydrate product in 1 L water containing 10 mL 1% rosolic acid in 0.2N NaOH.*#(41)
Heat to near boiling, promptly remove from heat, and cool to below 50°C. Do not sterilize by
autoclaving. If agar is used, dispense 5- to 7-mL quantities to 50- × 12-mm petri plates and let
solidify. Final pH should be 7.4 ± 0.2. Refrigerate finished medium, preferably in sealed plastic
bags or other containers to reduce moisture loss, and discard unused broth after 96 h or unused
agar after 2 weeks.
Test each medium lot against a previously acceptable lot for satisfactory performance as
described in Section 9020B, by making dilutions of a culture of E. coli (Section 9020) and
filtering appropriate volumes to give 20 to 60 colonies per filter. With each new lot of medium
verify 10 or more colonies obtained from several natural samples, to establish the absence of
false positives. For most samples M-FC medium may be used without the 1% rosolic acid
addition, provided there is no interference with background growth. Such interference may be
expected in stormwater samples collected during the first runoff (initial flushing) after a long dry
period.
Before use, test each batch of laboratory-prepared MF medium for performance with positive
and negative culture controls. Check for coliform contamination at the beginning and end of each
filtration series by filtering 20 to 30 mL of dilution or rinse water through filter. If controls
indicate contamination, reject all data from affected samples and request resample.
b. Culture dishes: Tight-fitting plastic dishes are preferred because the membrane filter
cultures are submerged in a water bath during incubation. Place fecal coliform cultures in plastic
bags or seal individual dishes with waterproof (freezer) tape to prevent leakage during
submersion. Specifications for plastic culture dishes are given in Section 9222B.1e.
c. Incubator: The specificity of the fecal coliform test is related directly to the incubation
temperature. Static air incubation may be a problem in some types of incubators because of
potential heat layering within the chamber, slower heat transfer from air to the medium, and the
slow recovery of temperature each time the incubator is opened during daily operations. To meet
the need for greater temperature control use a water bath, a heat-sink incubator, or a properly
designed and constructed incubator shown to give equivalent results. A temperature tolerance of
44.5 ± 0.2°C can be obtained with most types of water baths that also are equipped with a gable
top for the reduction of water and heat losses.
2. Procedure
a. Selection of sample size: Select volume of water sample to be examined in accordance
with the information in Table 9222:III. Use sample volumes that will yield counts between 20
and 60 fecal coliform colonies per membrane.
When the bacterial density of the sample is unknown, filter several volumes or dilutions to
achieve a countable density. Estimate volume and/or dilution expected to yield a countable
membrane and select two additional quantities representing one-tenth and ten times this volume,
respectively.
b. Filtration of sample: Follow the same procedure and precautions as prescribed under
Section 9222B.5b above.
c. Preparation of culture dish: Place a sterile absorbent pad in each culture dish and pipet at
least 2.0 mL M-FC medium, prepared as directed above, to saturate pad. Carefully remove any
excess liquid from culture dish by decanting the plate. Aseptically, place prepared filter on
medium-impregnated pad as described in Section 9222B above.
As a substrate substitution for the nutrient-saturated absorbent pad, add 1.5% agar to M-FC
broth as described in Section 9222B above.
d. Incubation: Place prepared dishes in waterproof plastic bags or seal, invert, and submerge
petri dishes in water bath, and incubate for 24 ± 2 h at 44.5 ± 0.2°C. Anchor dishes below water
surface to maintain critical temperature requirements. Place all prepared cultures in the water
bath within 30 min after filtration. Alternatively, use an appropriate, accurate solid heat sink or
equivalent incubator.
e. Counting: Colonies produced by fecal coliform bacteria on M-FC medium are various
shades of blue. Nonfecal coliform colonies are gray to cream-colored. Normally, few nonfecal
coliform colonies will be observed on M-FC medium because of selective action of the elevated
temperature and addition of rosolic acid salt reagent. Count colonies with a low-power (10 to 15
magnifications) binocular wide-field dissecting microscope or other optical device.
f. Verification: Verify typical blue colonies and any atypical grey to green colonies as
described in Section 9020 for fecal coliform analysis. Simultaneous inoculation at both
temperatures is acceptable.
3. Calculation of Fecal Coliform Density
a. General: Compute the density from the sample quantities that produced MF counts within
the desired range of 20 to 60 fecal coliform colonies. This colony density range is more
restrictive than the 20 to 80 total coliform range because of larger colony size on M-FC medium.
Calculate fecal coliform density as directed in Section 9222B.6 above. Record densities as fecal
coliforms per l00 mL.
b. Sediment and biosolid samples: For total solid (dry weight basis) see Section 2540G.
Calculate fecal coliforms per gram dry weight for biosolid analysis as follows:
where dilution and % dry solids are expressed in decimal form.
Example 1: There were 22 colonies observed on the 1:10 000 dilution plate of a biosolid with
4% dry solids.
If no filter has a coliform count falling in the ideal range (20 to 60), total the coliform counts
on all countable filters and report as fecal coliforms per gram dry weight.
Example 2: There were 18 colonies observed on the 1:10 000 dilution plate and 2 colonies
observed on the 1:100 000 dilution plate of a biosolid sample with 4% dry solids.
To compute a geometric mean of samples, convert coliform densities of each sample to log10
values. Determine the geometric mean for the given number of samples (usually seven) by
averaging the log10 values of the coliform densities and taking the antilog of that value.
4. Bibliography
GELDREICH, E.E., H.F. CLARK, C.B. HUFF & L.C. BEST. 1965. Fecal-coliform-organism medium for
the membrane filter technique. J. Amer. Water Works Assoc. 57:208.
ROSE, R.E., E.E. GELDREICH & W. LITSKY. 1975. Improved membrane filter method for fecal
coliform analysis. Appl. Microbiol. 29:532.
LIN, S.D. 1976. Membrane filter method for recovery of fecal coliforms in chlorinated sewage
effluents. Appl. Environ. Microbiol. 32:547.
PRESSWOOD, W.G. & D.K. STRONG. 1978. Modification of M-FC medium by eliminating rosolic
acid. Appl. Environ. Microbiol. 36:90.
GREEN, B.L., W. LITSKY & K.J. SLADEK. 1980. Evaluation of membrane filter methods for
enumeration of faecal coliforms from marine waters. Mar. Environ. Res. 67:267.
SARTORY, D.P. 1980. Membrane filtration faecal coliform determinations with unmodified and
modified M-FC medium. Water SA 6:113.
GRABOW, W.O.K., C.A. HILNER & P. COUBROUGH. 1981. Evaluation of standard and modified
M-FC, MacConkey, and Teepol media for membrane filter counting of fecal coliform in
RYCHERT, R.C. & G.R. STEPHENSON. 1981. Atypical Escherichia coli in streams. Appl. Environ.
Microbiol. 41:1276.
PAGEL, J.E., A.A. QURESHI, D.M. YOUNG & L.T. VLASSOFF. 1982. Comparison of four membrane
filter methods for fecal coliform enumeration. Appl. Environ. Microbiol. 43:787.
U.S. ENVIRONMENTAL PROTECTION AGENCY. 1992. Environmental Regulations and
Technology. Control of Pathogens and Vector Attraction in Sewage Sludge.
EPA-626/R-92-013, Washington, D.C.
U.S. ENVIRONMENTAL PROTECTION AGENCY. 1993. Standards for the Use or Disposal of
Sewage Sludge: Final Rule. 40 CFR Part 257; Federal Register 58:9248, Feb. 19, 1993.
9222 E.
Delayed-Incubation Fecal Coliform Procedure
This delayed-incubation procedure is similar to the delayed-incubation total coliform
procedure (Section 9222C). Use the delayed-incubation test only when the standard immediate
fecal coliform test cannot be performed (i.e., where the appropriate field incubator is not
available, or where, under certain circumstances, a specialized laboratory service is advisable to
examine, confirm, or speciate the suspect colonies).
Results obtained by this delayed method have been consistent with results from the standard
fecal coliform MF test under various laboratory and field use conditions. However, determine
test applicability for a specific water source by comparison with the standard MF test, especially
for saline waters, chlorinated wastewaters, and waters containing toxic substances.
To conduct the delayed-incubation test filter sample in the field immediately after collection,
place filter on M-ST holding medium (see Section 9222C.2b below), and ship to the laboratory.
Complete fecal coliform test by transferring filter to M-FC medium, incubating at 44.5°C for 24
± 2 h, and counting fecal coliform colonies.
The M-ST medium keeps fecal coliform organisms viable but prevents visible growth during
transit. Membrane filters can be held for up to 3 d on M-ST holding medium with little effect on
the fecal coliform counts.
1. Apparatus
a. Culture dishes: See Section 9222C.1a for specifications.
b. Field filtration units: See Section 9222C.1b.
2. Materials and Transport Medium
a. M-ST medium: Prepare as described in Section 9222C.2b.
b. M-FC medium: Prepare as described in Section 9222D.1a.
3. Procedure
a. Membrane filter transport: Place an absorbent pad in a tight-lid plastic petri dish and
saturate with M-ST holding medium. After filtering sample remove membrane filter from
filtration unit and place it on medium-saturated pad. Use only tight-lid dishes to prevent moisture
loss; however, avoid having excess liquid in the dish. Place culture dish containing membrane in
an appropriate shipping container and send to laboratory. Membranes can be held on the
transport medium at ambient temperature for a maximum of 72 h with little effect on fecal
coliform counts.
b. Transfer: At the laboratory remove membrane from holding medium and place it in
another dish containing M-FC medium.
c. Incubation: After transfer of filter to M-FC medium, place tight-lid dishes in waterproof
plastic bags, invert, and submerge in a water bath at 44.5°C ± 0.2°C for 24 ± 2 h or use a solid
heat sink or equivalent incubator.
d. Counting: Colonies produced by fecal coliform bacteria are various shades of blue.
Nonfecal coliform colonies are gray to cream-colored. Count colonies with a binocular
wide-field dissecting microscope at 10 to 15 magnifications.
e. Verification: Verify typical blue colonies and any atypical (grey to green) colonies as
described in Section 9020 for fecal coliform analysis.
4. Estimation of Fecal Coliform Density
Count as directed in Section 9222D.2e above and compute fecal coliform density as
described in Section 9222D.3. Record time of collection, filtration, and laboratory examination,
and calculate and report elapsed time.
5. Bibliography
CHEN, M. & P.J. HICKEY. 1983. Modification of delayed-incubation procedure for detection of
fecal coliforms in water. Appl. Environ. Microbiol. 46:889.
9222 F.
Klebsiella Membrane Filter Procedure
Klebsiella bacteria belong to the family Enterobacteriaceae and are included in the total
coliform group. The outermost layer of Klebsiella bacteria consists of a large polysaccharide
capsule, a characteristic that distinguishes this genus from most other bacteria in this family; this
capsule provides some measure of protection from disinfectants. Klebsiella bacteria are
commonly associated with coliform regrowth in large water supply distribution systems.
Klebsiellae may be opportunistic pathogens that can give rise to bacteremia, pneumonia,
urinary tract, and several other types of human infection. Approximately 60 to 80% of all
Klebsiella from feces and from clinical specimens are positive in the fecal coliform test and are
Klebsiella pneumoniae.
Klebsiella bacteria also are widely distributed in nature, occurring in soil, water, grain,
vegetation, etc. Wood pulp, paper mills, textile finishing plants, and sugar-cane processing
operations contain large numbers of klebsiellae in their effluents (104 to 106), and Klebsiella sp.
are often the predominant coliform in such effluents.
Rapid quantitation may be achieved in the MF procedure by modifying M-FC agar base
through substitution of inositol for lactose and adding carbenicillin or by using M-Kleb agar.
These methods reduce the necessity for biochemical testing of pure strains. Preliminary
verification of differentiated colonies is recommended.
1. Apparatus
a. Culture dishes: See Section 9222B.1e for specifications.
b. Filtration units: See Section 9222B.1 f.
2. Materials and Culture Medium
a. Modified M-FC agar (M-FCIC agar): This medium may not be available in dehydrated
form and may require preparation from the basic ingredients:
Tryptose or biosate
Yeast extract
Inositol
Bile salts No. 3 or bile salts mixture
Aniline blue
Agar
Reagent-grade water
10.0
5.0
3.0
5.0
10.0
1.5
0.1
15.0
1
g
g
g
g
g
g
g
g
L
Heat medium to boiling and add 10 mL 1% rosolic acid dissolved in 0.2N NaOH. Cool to
below 45°C and add 50 mg carbenicillin.*#(42) Dispense aseptically in 5- to 7-mL quantities
into 50- × 9-mm plastic petri dishes. Refrigerate until needed. Discard unused agar medium
after 2 weeks. Do not sterilize by autoclaving. Final pH should be 7.4 ± 0.2.
b. M-Kleb agar:
Phenol red agar
Adonitol
Aniline blue
31.0 g
5.0 g
0.1 g
Reagent-grade water
0.1 g
1 L
Sterilize by autoclaving for 15 min at 121°C. After autoclaving, cool to 50°C in a water
bath; add 20 mL 95% ethyl alcohol (not denatured) and 0.05 g filter sterilized carbenicillin/L.
Shake thoroughly and dispense aseptically into 50- × 9-mm plastic culture plates. The final pH
should be 7.4 ± 0.2. Refrigerated medium can be held for 20 d at 4 to 8°C.
3. Procedure
a. See Section 9222B.5 for selection of sample size and filtration procedure. Select sample
volumes that will yield counts between 20 and 60 Klebsiella colonies per membrane. Place
membrane filter on agar surface; incubate for 24 ± 2 h at 35 ± 0.5°C. Klebsiella colonies on
M-FCIC agar are blue or bluish-gray. Most atypical colonies are brown or brownish. Occasional
false positive occurrences are caused by Enterobacter species. Klebsiella colonies on M-Kleb
agar are deep blue to blue gray, whereas other colonies most often are pink or occasionally pale
yellow. Count colonies with a low-power (10 to 15 magnifications) binocular wide field
dissecting microscope or other optical device.
b. Verification: Verify Klebsiella colonies from the first set of samples from ambient waters
and effluents and when Klebsiella is suspect in water supply distribution systems. Verify a
minimum of five typical colonies by transferring growth from a colony or pure culture to a
commercial multi-test system for gram-negative speciation. Key tests for Klebsiella are citrate
(positive), motility (negative), lysine decarboxylase (positive), ornithine decarboxylase
(negative), and urease (positive). A Klebsiella strain that is indole-positive, liquefies pectin, and
demonstrates a negative fecal coliform response is most likely of nonfecal origin.
4. Bibliography
DUNCAN, D.W. & W.E. RAZELL. 1972. Klebsiella biotypes among coliforms isolated from forest
environments and farm produce. Appl. Microbiol. 24:933.
STRAMER, S.L. 1976. Presumptive identification of Klebsiella pneumoniae on M-FC medium.
BAGLEY, S.T. & R.J. SEIDLER. 1977. Significance of fecal coliform-positive Klebsiella. Appl.
KNITTEL, M.D., R.J. SEIDLER, C. EBY & L.M. CABE. 1977. Colonization of the botanical
environment by Klebsiella isolates of pathogenic origin. Appl. Environ. Microbiol. 34:557.
EDMONSON, A.S., E.M. COOK, A.P.D. WILCOCK & R. SHINEBAUM. 1980. A comparison of the
properties of Klebsiella isolated from different sources. J. Med. Microbiol. (U.K.) 13:541.
SMITH, R.B. 1981. A Critical Evaluation of Media for the Selective Identification and
Enumeration of Klebsiella. M.S. thesis, Dep. Civil & Environmental Engineering, Univ.
Cincinnati, Ohio.
NIEMELA, S.I. & P. VAATANEN. 1982. Survival in lake water of Klebsiella pneumoniae
discharged by a paper mill. Appl. Environ. Microbiol. 44:264.
GELDREICH, E.E. & E.W. RICE. 1987. Occurrence, significance, and detection of Klebsiella in
water systems. J. Amer. Water Works Assoc. 79:74.
DUNCAN, I.B.R. 1988. Waterborne Klebsiella and human disease. Toxicity Assess. 3:581.
9222 G.
MF Partition Procedures
1. Escherichia coli Partition Methods
a. Applications: Escherichia coli is a member of the fecal coliform group of bacteria; its
presence is indicative of fecal contamination. Rapid quantitation and verification may be
achieved with the MF procedure by transferring the membrane from a total-coliform- or
fecal-coliform-positive sample to a nutrient agar substrate containing
4-methylumbelliferyl-β-D-glucuronide (MUG). In this method E. coli is defined as any coliform
that produces the enzyme β-glucuronidase and hydrolyzes the MUG substrate to produce a blue
fluorescence around the periphery of the colony.
In the examination of drinking water samples, use this method to verify the presence of E.
coli from a total-coliform-positive MF on Endo-type media. In the examination of wastewater
and other nonpotable water samples, use this procedure to verify positive filters from mFC
medium used in the fecal coliform MF procedure.
b. Apparatus:
1) Culture dishes: See Section 9222B.1e.
2) Filtration units: See Section 9222B.1 f.
3) Forceps: See Section 9222B.1i.
4) Incubator: See Section 9222B.1 j.
5) Ultraviolet lamp, long wave (366 nm), preferably 6 W.
6) Microscope and light source: See Section 9222B.1k.
c. Materials and culture medium:
1) Nutrient agar with MUG (NA-MUG):
Peptone
Beef extract
Agar
4-methylumbelliferyl-β-D-glucuronide
Reagent-grade water
5.0
3.0
15.0
0.1
g
g
g
g
1
L
Add dehydrated ingredients to reagent-grade water, mix thoroughly, and heat to dissolve.
Sterilize by autoclaving for 15 min at 121°C. Dispense aseptically into 50-mm plastic culture
plates. The final pH should be 6.8 ± 0.2. Refrigerated prepared medium may be held for 2 weeks.
2) EC broth with MUG (EC-MUG):
Lactose
20.0
5.0
1.5
4.0
g
g
g
g
1.4 g
5.0 g
0.1 g
4-methylumbelliferyl-β-D-glucuronide
Reagent-grade water
1
L
Add dehydrated ingredients to reagent-grade water, mix thoroughly and heat to dissolve. pH
should be 6.9 ± 0.2 after sterilization. Before sterilization, dispense into culture tubes and cap
with metal or heat-resistant plastic caps.
d. Procedure: See Section 9222B.5 for selection of sample size and filtration procedure. For
drinking water samples using Endo-type medium, count and record the metallic golden sheen
colonies. Before transfer of the membrane, transfer a small portion of each target colony to the
appropriate total coliform verification medium, using a sterile needle. See Section 9222B.5 for
total coliform verification procedures.
Alternatively, after transfer and incubation on NA-MUG, swab the surface growth on the
filter and transfer to the appropriate total coliform verification medium. Aseptically transfer the
membrane from the Endo-type medium to NA-MUG or EC-MUG medium. If differentiation of
the total coliforms is desired using NA-MUG medium, mark each sheen colony with a
fine-tipped marker or by puncturing a hole in the membrane adjacent to the colony with a sterile
needle. Incubate NA-MUG at 35 ± 0.5°C for 4 h or EC-MUG at 44.5 ± 0.2 for 24 ±2 h. Observe
individual colonies or tubes using a long-wave-length (366-nm) ultraviolet light source,
preferably containing a 6-W bulb. The presence of a blue fluorescence in the tube, on the
periphery (outer edge) of a colony, or observed from the back of the plate is considered a
positive response for E. coli. Count and record the number of target colonies, if quantification is
desired, or just record presence or absence of fluorescence.
For nonpotable water samples, use mFC medium for initial isolation before transfer to
NA-MUG or EC-MUG medium. The procedure is the same as the above, with the exception of
the total coliform verification process.
For the EC-MUG method, a positive control consisting of a known E. coli (MUG-positive)
culture, a negative control consisting of a thermotolerant Klebsiella pneumoniae
(MUG-negative) culture, and an uninoculated medium control may be necessary to interpret the
results and to avoid confusion of weak autofluorescence of the medium as a positive response.
See Section 9221F.
2. Fecal Coliform Partition Method
a. Applications: Further partitioning of total coliforms from the original MF
coliform-positive culture in a presence/absence search for fecal coliform in a drinking water
sample may be achieved within 24 h. This procedure provides additional information from the
original sample.
b. Materials and culture medium: EC broth. See Section 9221E.1a.
c. Procedure: See Section 9222B.5 for selection of sample size and filtration procedure. For
drinking water samples using Endo-type media, count and record the metallic (golden) sheen
colonies. Before transfer of membrane or swabbing of plate, transfer a small portion of each
target colony to the appropriate total coliform verification media using a sterile needle (see
Section 9222B.5 f). Use a sterile cotton swab to collect bacteria from the membrane surface, or
pick discrete colonies with a 3-mm loop or sterile applicator stick, or transfer the entire
membrane to inoculate a tube of EC medium. Incubate inoculated EC broth in a water bath at
44.5 ± 0.2°C for 24 ± 2 h. Place all EC tubes in water bath within 30 min after inoculation.
Maintain a sufficient water depth in water bath incubator to immerse tubes to upper level of the
medium. Gas production in an EC broth culture in 24 h or less is considered a positive response
for fecal coliform bacteria.
3. Bibliography
U.S. ENVIRONMENTAL PROTECTION AGENCY. 1989. Drinking Water; National Primary Drinking
Water Regulations; Total Coliforms (Including Fecal Coliforms and E. coli); Final Rule. 40
CFR Parts 141 and 142. Federal Register 54:27544, June 29, 1989.
MATES, A. & M. SHAFFER. 1989. Membrane filtration differentiation of E. coli from coliforms in
the examination of water. J. Appl. Bacteriol. 67:343.
U.S. ENVIRONMENTAL PROTECTION AGENCY. 1991. National Primary Drinking Water
Regulations; Analytical Techniques; Coliform Bacteria. 40 CFR Part 141, Federal Register
56:636, Jan. 8, 1991.
MATES, A. & M. SHAFFER. 1992. Quantitative determination of Escherichia coli from coliforms
and fecal coliforms in sea water. Microbios 71:27.
SARTORY, D. & L. HOWARD. 1992. A medium detecting beta-glucuronidase for the simultaneous
membrane filtration enumeration of Escherichia coli and coliforms from drinking water. Lett.
Appl. Microbiol. 15:273.
SHADIX, L.C., M.E. DUNNIGAN & E.W. RICE. 1993. Detection of Escherichia coli by the nutrient
agar plus 4-methylumbelliferyl-β-D-glucuronide (MUG) membrane filter method. Can. J.
Microbiol. 39: 1066.
9223
ENZYME SUBSTRATE COLIFORM TEST*#(43)
9223 A.
Introduction
The enzyme substrate test utilizes hydrolyzable substrates for the simultaneous detection of
total coliform bacteria and Escherichia coli enzymes. When the enzyme technique is used, the
total coliform group is defined as all bacteria possessing the enzyme β-D-galactosidase, which
cleaves the chromogenic substrate, resulting in release of the chromogen. Escherichia coli are
defined as bacteria giving a positive total coliform response and possessing the enzyme
β-glucuronidase, which cleaves a fluorogenic substrate, resulting in the release of the fluorogen.
The test can be used in either a multiple-tube, multi-well, or a presence-absence (single 100-mL
sample) format.
1. Principle
a. Total coliform bacteria: Chromogenic substrates, such as
ortho-nitrophenyl-β-D-galactopyranoside (ONPG) or chlorophenol red-β-D-galactopyranoside
(CPRG), are used to detect the enzyme β-D-galactosidase, which is produced by total coliform
bacteria. The β-D-galactosidase enzyme hydrolyzes the substrate and produces a color change,
which indicates a positive test for total coliforms at 24 h (ONPG) or 28 h (CPRG) without
additional procedures. Noncoliform bacteria, such as Aeromonas and Pseudomonas species, may
produce small amounts of the enzyme β-D-galactosidase, but are suppressed and generally will
not produce a positive response within the incubation time unless more than 104 colony-forming
units (CFU)/mL (106 CFU/100 mL) are present.
b. Escherichia coli: A fluorogenic substrate, such as 4-methylumbelliferyl-β-D-glucuronide
(MUG), is used to detect the enzyme β-glucuronidase, which is produced by E. coli. The
β-glucuronidase enzyme hydrolyzes the substrate and produces a fluorescent product when
viewed under long-wavelength (366-nm) ultraviolet (UV) light. The presence of fluorescence
indicates a positive test for E. coli. Some strains of Shigella spp. also may produce a positive
fluorescence response. Because Shigella spp. are overt human pathogens, this is not considered a
detriment for testing the sanitary quality of water.
2. Applications
The enzyme substrate coliform test is recommended for the analysis of drinking and source
water samples. Formulations also are available for the analysis of marine waters. Initially,
laboratories planning to use this procedure should conduct parallel quantitative testing (including
seasonal variations) with one of the standard coliform tests to assess the effectiveness of the test
for the specific water type being analyzed and to determine the comparability of the two
techniques. This is particularly important when testing source waters.
Water samples containing humic or other material may be colored. If there is background
color, compare inoculated tubes to a control tube containing only water sample. In certain
waters, high calcium salt content can cause precipitation but this should not affect the reaction.
Do not use the enzyme substrate test to verify presumptive coliform cultures or membrane
filter colonies, because the substrate may be overloaded by the heavy inoculum of weak
β-D-galactosidase-producing noncoliforms, causing false-positive results.
9223 B.
Enzyme Substrate Test
1. Substrate Media
Formulations are available commercially*#(44) in disposable tubes for the multiple-tube
procedure, in disposable multi-wells†#(45) for the multi-well procedure, or in containers that
will hold 100-mL samples for the presence-absence approach.* Appropriate preweighed portions
of the reagent for mixing and dispensing into multiple tubes for 10-mL test portions or other
containers for 100-mL samples also are available. The need for good quality assurance and
uniformity requires the use of a commercial substrate medium. Avoid prolonged exposure of the
substrate to direct sunlight. Store media according to directions and use before expiration date.
Discard discolored media.
2. Procedure
a. Multiple-tube procedure: Select the appropriate number of tubes per sample with
predispensed media for the multiple-tube test and label. Follow manufacturer’s instructions for
preparing serial dilutions for various formulations. Aseptically add 10 mL sample to each tube,
cap tightly, and mix vigorously to dissolve. The mixture remains colorless with ONPG-based
tests and turns yellow with the CPRG format. Some particles may remain undissolved
throughout the test; this will not affect test performance. Incubate at 35 ± 0.5°C for period
specified by substrate manufacturer.
The procedure also can be performed by adding appropriate amounts of the substrate media
to the sample, mixing thoroughly, and dispensing into five or ten sterile tubes. Incubate as stated
for multiple-tube procedure.
b. Multi-well procedure: The multi-well procedure is performed with sterilized disposable
packets. Add sample to 100-mL container with substrate, shake vigorously, and pour into tray.
The tray sealer dispenses the sample into the wells and seals the package. Incubate at 35 ± 0.5°C
for period specified by substrate manufacturer. The MPN value is obtained from the table
provided by the manufacturer.
c. Presence-absence procedure (P/A): Aseptically add preweighed enzymatic medium to
100-mL sample in a sterile, transparent, nonfluorescent borosilicate glass or equivalent bottle or
container. Optionally, add 100-mL sample to the enzymatic substrate in a sterile container
provided by the manufacturer. Aseptically cap and mix thoroughly to dissolve. Incubate as
specified in manufacturer’s instructions.
3. Interpretation
a. Total coliform bacteria: After the minimum proper incubation period, examine tubes or
containers for the appropriate color change (Table 9223:I). ONPG is hydrolyzed by the bacterial
enzyme to yield a yellow color. CPRG is hydrolyzed by the bacterial enzyme to yield a red or
magenta color. If the color response is not uniform throughout the tube, mix by inversion before
reading. Read manufacturer’s instructions for interpretation guidelines. Some manufacturers
suggest comparing sample tubes against a color comparator available through the manufacturer.
Samples are negative for total coliforms if no color is observed in ONPG tests or if the tube is
yellow when CPRG is used. If a chromogenic response is questionable after 18 or 24 h for
ONPG, incubate up to an additional 4 h. If response is negative after 28 h for CPRG, incubate up
to an additional 20 h. If the chromogen intensifies, the sample is total-coliform positive; if it does
not, the sample is negative.
b. Escherichia coli: Examine positive total coliform tubes or containers for fluorescence
using a long-wavelength (366-nm) ultraviolet lamp (preferably 6-W bulb). Compare each tube
against the reference comparator available from a commercial source of the substrate. The
presence of fluorescence is a positive test for E. coli. If fluorescence is questionable, incubate for
an additional 4 h for ONPG tests and up to an additional 20 h for CPRG tests; intensified
fluorescence is a positive test result.
4. Reporting
If performing an MPN procedure, calculate the MPN value for total coliforms and E. coli
from the number of positive tubes as described in Section 9221C. If using the presence-absence
procedure, report results as total coliform and E. coli present or absent in 100-mL sample.
5. Quality Control
Test each lot of media purchased for performance by inoculation with three control bacteria:
Escherichia coli, a total coliform other than E. coli (e.g., Enterobacter cloacae), and a
noncoliform. Also add a sterile water control. If the sterile water control exhibits faint
fluorescence or faint positive coliform result, discard and use a new batch of substrate. Avoid
using a heavy inoculum. If Pseudomonas is used as the representative noncoliform, select a
nonfluorescent species. Incubate these controls at 35 ± 0.5°C as indicated above. Read and
record results. Other quality-control guidelines are included in Section 9020.
6. Bibliography
EDBERG, S.C., M.J. ALLEN, D.B. SMITH & THE NATIONAL COLLABORATIVE STUDY. 1988.
National field evaluation of a defined substrate method for the simultaneous enumeration of
total coliforms and Escherichia coli from drinking water: Comparison with the standard
multiple tube fermentation method. Appl. Environ. Microbiol. 54:1595.
EDBERG, S.C. & M.M. EDBERG. 1988. A defined substrate technology for the enumeration of
microbial indicators of environmental pollution. Yale J. Biol. Med. 61:389.
COVERT, T.C., L.C. SHADIX, E.W. RICE, J.R. HAINES & R.W. FREYBERG. 1989. Evaluation of the
Autoanalysis Colilert test for detection and enumeration of total coliforms. Appl. Environ.
Microbiol. 55:2443.
EDBERG, S.C. & D.B. SMITH. 1989. Absence of association between total heterotrophic and total
coliform bacteria from a public water supply. Appl. Environ. Microbiol. 55:380.
EDBERG, S.C., M.J. ALLEN, D.B. SMITH & THE NATIONAL COLLABORATIVE STUDY. 1989.
National field evaluation of a defined substrate method for the simultaneous detection of
total coliforms and Escherichia coli from drinking water: Comparison with presenceabsence techniques. Appl. Environ. Microbiol. 55:1003.
EDBERG, S.C., M.J. ALLEN, D.B. SMITH & N.J. KRIZ. 1990. Enumeration of total coliforms and
Escherichia coli from source water by the defined substrate technology. Appl. Environ.
Microbiol. 56:366.
RICE, E.W., M.J. ALLEN & S.C. EDBERG. 1990. Efficacy of β-glucuronidase assay for identification
of Escherichia coli by the defined-substrate technology. Appl. Environ. Microbiol. 56:1203.
RICE, E.W., M.J. ALLEN, D.J. BRENNER & S.C. EDBERG. 1991. Assay for β-glucuronidase in species
of the genus Escherichia and its application for drinking water analysis. Appl. Environ.
Microbiol. 57:592.
SHADIX, L.C. & E.W. RICE. 1991. Evaluation of β-glucuronidase assay for the detection of
Escherichia coli from environmental waters. Can. J. Microbiol. 37:908.
EDBERG, S.C., M.J. ALLEN & D.B. SMITH. 1991. Defined substrate technology method for rapid
and simultaneous enumeration of total coliforms and Escherichia coli from water:
Collaborative study. J. Assoc. Offic. Anal. Chem. 74:526.
EDBERG, S.C., F. LUDWIG & D.B. SMITH. 1991. The Colilert® System for Total Coliforms and
Escherichia coli. American Water Works Association Research Foundation, Denver, Colo.
COVERT, T.C., E.W. RICE, S.A. JOHNSON, D. BERMAN, C.H. JOHNSON & P.M. MASON. 1992.
Comparing defined-substrate coliform tests for the detection of Escherichia coli in water. J.
Amer. Water Works Assoc. 84(5):98.
MCCARTY, S.C., J.H. STANDRIDGE & M.C. STASIAK. 1992. Evaluating a commercially available
defined-substrate test for recovery of chlorine-treated Escherichia coli. J. Amer. Water
Works Assoc. 84(5): 91.
PALMER, C.J., Y. TSAI, A.L. LANG & L.R. SANGERMANO. 1993. Evaluation of Colilert-marine
water for detection of total coliforms and Escherichia coli in the marine environment. Appl.
CLARK, J.A. & A.H. SHAARAWI. 1993. Evaluation of commercial presence-absence test kits for
detection of total coliforms, Escherichia coli, and other indicator bacteria. Appl. Environ.
Microbiol. 59:380.
U.S. ENVIRONMENTAL PROTECTION AGENCY. 1994. National Primary and Secondary Drinking
Water Regulation: Analytical methods for regulated drinking water contaminants; Final
Rule. 40 CFR Parts 141 & 143; Federal Register 59:62456.
MCFETERS, G.A., S.C. BROADWAY, B.H. PYLE, M. PICKETT & Y. EGOZY. 1995. Comparative
performance of ColisureTM and accepted methods in the detection of chlorine-injured total
coliforms and E. coli. Water Sci. Technol. 31:259.
9225
DIFFERENTIATION OF THE COLIFORM BACTERIA*#(46)
9225 A.
Introduction
Identification of bacteria that constitute the coliform group sometimes is necessary to
determine the nature of pollution. It is of particular importance in reference to distinguishing the
presence of Escherichia coli. Special procedures for detection of E. coli are given in Section
9221F, Section 9222G, and Section 9223. Differential tests for identification must be used with
the knowledge that all strains taxonomically assigned to the coliform group do not conform
necessarily to the coliform definition stated in this manual because they may not ferment lactose,
or if they do, they may not produce gas. Furthermore, gram-negative bacteria other than
coliforms ferment lactose and produce sheen (e.g., Aeromonas spp.) and not all strains of a
species will react uniformly in media. Unusual strains (such as E. coli, inactive, Table 9225:I),
mutants, and injured organisms may not give classical responses. The traditional ‘‘IMViC’’ tests
(i.e., indole, methyl red, Voges-Proskauer, and citrate utilization) are useful for coliform
differentiation, but do not provide complete identification. Additional biochemical tests often
are necessary. Commercial kits for identification are available and may serve as economical
alternatives to traditional differential media. Automated systems of identifying large numbers of
isolates also are available.
The significance of various coliform organisms in water has been and is a subject of
considerable study. Collectively, the coliforms are referred to as indicator organisms. The genera
Enterobacter, Klebsiella, Citrobacter, and Escherichia usually are represented in the majority of
isolations made from raw and treated municipal water supplies.
9225 B.
Culture Purification
1. Procedure
A pure culture is essential for accurate identification. Obtain a pure culture by carefully
picking a well-isolated colony that gives typical responses on an appropriate solid medium or
membrane filter, and streaking on a tryptic soy or nutrient agar plate. Better distribution of
colonies in the subculture is obtained if a portion of the picked colony is emulsified in peptone
broth or physiological saline (0.85% w/v) and then streaked. When picking a colony from a
primary culture on a selective medium, be aware that viable cells, which have not formed
colonies themselves, may surround the picked colony. Incubate the subculture at 35 ± 0.5°C for
24 h and test a single well-isolated colony by the Gram stain to confirm the sole presence of
gram-negative, non-spore-forming rods (Section 9221B). Also determine that the culture is
oxidase-negative (Section 9225D). Oxidase-positive, gram-negative, non-spore-forming rods are
not coliform bacteria, but may be organisms such as Aeromonas, which is not regarded as an
indicator of fecal pollution.
Variation in organisms of the coliform group occurs occasionally and mixed reactions in
differential media may indicate a pure culture undergoing variation. Persistent variations of
reactions in differential media indicate a mixed culture caused by inadequate purification.
2. Bibliography
PTAK, D.J., W. GINSBURG & B.F. WILLEY. 1974. Aeromonas, the great masquerader. Proc.
AWWA Water Quality Technology Conf., Dallas, Tex., p. V-1. American Water Works
Assoc., Denver, Colo.
VAN DER KOOJ, D. 1988. Properties of aeromonads and their occurrence and hygienic
significance in drinking water. Zentralbl. Bacteriol. Hyg. B 187:1.
HARTMAN, P.A., B. SWAMINATHAN, M.S. CURIALE, R. FIRSTENBERG-EDEN, A.N. SHARPE, N.A.
COX, D.Y.C. FUNG & M.C. GOLDSCHMIDT. 1992. Rapid methods and automation. In: C.
Vanderzant & D.F. Splittstoesser, eds., Compendium of Methods for the Microbiological
Examination of Foods, 3rd. ed. p.665. American Public Health Assoc., Washington, D.C.
STAGER, C.E. & J.R. DAVIS. 1992. Automated systems for identification of microorganisms. Clin.
Microbiol. Rev. 5:302.
RICE, E.W., M.J. ALLEN, T.C. COVERT, J. LANGEWIS & J. STANDRIDGE. 1993. Identifying
Escherichia species with biochemical test kits and standard bacteriological tests. J. Amer.
Water Works Assoc. 85(2): 74.
9225 C.
Identification
1. Definition
Coliforms are defined here as facultative anaerobic, gram-negative non-spore-forming rods
that ferment lactose with gas formation within 48 h at 35 °C or, as applied to the membrane filter
method, produce a dark red colony with a metallic sheen within 24 h on an Endo-type medium
containing lactose. However, anaerogenic (non-gas-producing) lactose-fermenting strains of
Escherichia coli and coliforms that do not produce metallic sheen on Endo medium may be
encountered. These organisms, as well as typical coliforms, can be considered indicator
organisms, but they are excluded from the current definition of coliforms. More extensive testing
may be required for proper identification.
2. Characteristics and Tests
Coliforms belong to the bacterial taxonomic family Enterobacteriaceae. Table 9225:I
provides data on some of the biochemical reactions used for differentiating these organisms.
Preparing differential media and reagents may not be as economical for many laboratories as
using commercially prepared and prepackaged multiple-test kits, which reduce quality-control
work. These commercial kits are simple to store and use, and give reproducible and generally
accurate results. Periodically test reactions with known stock cultures of bacteria to assure
accuracy and reproducibility of results. Make further tests if the kit provides equivocal results.
3. Bibliography
KRIEG, N.R., ed. 1984. Bergey’s Manual of Systematic Bacteriology, Vol. I. Williams & Wilkins
Co., Baltimore, Md.
EDWARDS, P.R. & W.H. EWING. 1986. Identification of Enterobacteriaceae, 4th ed. Burgess Publ.
Co., Minneapolis, Minn.
9225 D.
Media, Reagents, and Procedures
Commercially available media and reagents can reduce work and cost; however, include
negative and positive controls with known stock cultures to assure accuracy and reliability.
Detailed methods are available. Expected test results are shown in Table 9225:I.
1. Lactose, Sorbitol, and Cellobiose Fermentation Tests
Suspend 16 g phenol red broth base and 5 g of the desired carbohydrate in 1 L reagent-grade
water and stir to dissolve completely. Dispense in tubes to a depth of one-third tube length. To
determine gas production place a small inverted vial (Durham tube) in the tubes of media at the
time of preparation. Close tubes and sterilize at 121°C for 15 min. Store tubes in the dark
(refrigeration preferred) and discard if evaporation exceeds 10% of the volume.
To conduct a test, inoculate with a loopful of growth from a well-isolated colony or slant and
incubate for 24 to 48 h at 35 ± 0.5°C. Carbohydrate fermentation (acid production) is indicated
by a decrease in pH, resulting in a change in color of the pH indicator, phenol red, from
red-orange to yellow (pH <6.6). Alternatively, for lactose fermentation, lauryl tryptose broth
(Section 9221B) may be used.
2. ONPG Hydrolysis
Numerous commercial test kits and disks for determining ONPG hydrolysis are available, or
an ONPG-containing medium (Section 9222) can be used. Alternatively, prepare peptone water
by dissolving 1 g peptone and 0.5 g NaCl in 100 mL reagent-grade water. Sterilize at 121°C for
15 min. Also prepare ONPG solution by dissolving 0.6 g o-nitrophenyl-β-D-galactopyranoside
(ONPG) in 100 mL 0.01M Na2HPO4, sterilize by filtration, and store in the dark at 4 to 10°C. To
prepare ONPG broth, aseptically combine 25 mL ONPG solution and 75 mL peptone water,
dispense aseptically in 2.5-mL amounts in sterile 13- × 100-mm tubes, and store in the dark for
up to 1 month at 4 to 10°C. Do not use the ONPG solution if it becomes yellow.
To conduct the test, inoculate 0.5 mL ONPG broth with a heavy loopful of growth from a
slant and incubate at 35 ± 0.5°C for up to 24 h. A yellow color, compared with an uninoculated
tube or (preferably) a tube inoculated with an ONPG-negative culture, is a positive test. Interpret
tests of yellow-pigmented organisms with caution. Do not use the enzyme substrate method
(Section 9223) to test ONPG hydrolysis.
3. Indole Test
Indole is a product of the metabolism of tryptophane.
a. Reagents:
1) Medium: Use tryptophane broth. Dissolve 10.0 g tryptone or trypticase/L reagent-grade
water. Dispense in 5-mL portions in test tubes and sterilize.
2) Test reagent: Dissolve 5 g p-dimethylaminobenzaldehyde in 75 mL isoamyl (or normal
amyl) alcohol, ACS grade, and add 25 mL conc HCl. The reagent should be yellow. Some
brands of p-dimethylaminobenzaldehyde are not satisfactory and some good brands become
unsatisfactory on aging.
The amyl alcohol solution should have a pH value of less than 6.0. Purchase both amyl
alcohol and benzaldehyde in as small amounts as will be consistent with the volume of work to
be done.
b. Procedure: Inoculate 5-mL portions of medium from a pure culture and incubate at 35 ±
0.5°C for 24 ± 2 h. Add 0.2 to 0.3 mL test reagent and gently shake. Let stand for about 10 min
and observe results.
A dark red color in the amyl alcohol surface layer constitutes a positive indole test; the
original color of the reagent, a negative test. An orange color probably indicates the presence of
skatole, a breakdown product of indole.
4. Methyl Red Test
The methyl red test measures the ability of organisms to produce stable acid end products
from glucose fermentation.
a. Reagents:
1) Medium: Use buffered glucose broth. Dissolve 7.0 g proteose peptone or equivalent
peptone, 5.0 g glucose, and 5.0 g dipotassium hydrogen phosphate (K2HPO4) in 1 L
reagent-grade water. Dispense in 5-mL portions in test tubes and sterilize by autoclaving at
121°C for 12 to 15 min, making sure that total time of exposure to heat is not longer than 30 min.
2) Indicator solution: Dissolve 0.1 g methyl red in 300 mL 95% ethyl alcohol and dilute to
500 mL with reagent-grade water.
b. Procedure: Inoculate 10-mL portions of medium from a pure culture. Incubate at 35 ±
0.5°C for 5 d. To 5 mL of the culture add 5 drops methyl red indicator solution.
Incubation for 48 h is adequate for most cultures, but do not incubate for less than 48 h. If
test results are equivocal at 48 h repeat with cultures incubated for 4 or 5 d. In such cases
incubate duplicate cultures at 22 to 25°C. Testing of culture portions at 2, 3, 4, and 5 d may
provide positive results sooner.
Record a distinct red color as methyl-red-positive and a distinct yellow color as
methyl-red-negative. Record a mixed shade as questionable and possibly indicative of
incomplete culture purification.
5. Voges-Proskauer Test
The Voges-Proskauer test measures the ability of organisms to produce a neutral end
product (acetoin) from glucose fermentation.
a. Reagents:
1) Medium: See ¶ 4a1) above.
2) Naphthol solution: Dissolve 5 g purified α-naphthol (melting point 92.5°C or higher) in
100 mL absolute ethyl alcohol. When stored at 5 to 10°C, this solution is stable for 2 weeks.
3) Potassium hydroxide, 7N: Dissolve 40 g KOH in 100 mL reagent-grade water.
b. Procedure: Inoculate 5 mL medium and incubate for 48 h at 35 ± 0.5°C. To 1 mL of
culture add 0.6 mL naphthol solution and 0.2 mL KOH solution. Shake well after the addition of
each reagent. Development of a pink to crimson color at the surface within 5 min constitutes a
positive test. Do not read after 10 min. Disregard tubes developing a copper color.
6. Simmons’ Citrate Test
The citrate test measures the ability of bacteria to utilize citrate as the sole source of carbon.
a. Medium: Use Simmons’ citrate agar. To make Simmons’ citrate agar, add 0.2 g
MgSO4⋅7H2O, 1.0 g ammonium dihydrogen phosphate (NH4H2PO4), 1.0 g K2HPO4, 2.0 g
sodium citrate dihydrate, 5.0 g NaCl, 15.0 g agar, and 0.08 g bromthymol blue to
1 L reagent-grade water. Tube for long slants.
b. Procedure: Inoculate agar medium by the streak technique using a light inoculum.
Incubate 48 h at 35 ± 0.5°C. Record growth on the medium with a blue color as a positive
reaction; record absence of growth or color change as negative.
7. Motility Test
The motility test measures whether an organism is motile in a semi-solid medium.
a. Medium: Use motility test medium made by adding 3.0 g beef extract, 10.0 g peptone, 5.0
g NaCl, and 4.0 g agar to 1 L reagent-grade water. Adjust pH to 7.4, dispense in 3-mL portions
in 13- × 100-mm tubes or 8-mL portions in 16- × 125-mm tubes, and sterilize.
b. Procedure: Inoculate by stabbing into the center of the medium, using an inoculating
needle, to a depth of 5 mm. Incubate for 1 to 2 d at 35°C. If negative, incubate an additional 5 d
at 22 to 25°C.
Diffuse growth through the medium from the point of inoculation is positive. In a negative
test, growth is visible only along the stab line and the surrounding medium stays clear.
Alternatively, prepare the medium without agar and examine a young culture using the hanging
drop slide technique for motile organisms.
8. Lysine and Ornithine Decarboxylase Tests
This procedure tests the ability of bacteria to metabolize the amino acids lysine and
ornithine.
a. Reagents:
1) Media: Use a basal medium made according to the Moeller or Falkow methods. For the
Moeller method, dissolve 5.0 g peptone (Orthana special, thiotone, or equivalent), 5.0 g beef
extract, 0.625 mL bromcresol purple (1.6%), 2.5 mL cresol red (0.2%), 0.5 g glucose, and 5.0
mg pyridoxal in 1 L reagent-grade water and adjust to pH 6.0 to 6.5. For the Falkow method,
dissolve 5.0 g peptone, 3.0 g yeast extract, 1.0 g glucose, and 1.0 mL bromcresol purple (1.6%)
in 1 L reagent-grade water and adjust to pH 6.7 to 6.8. For either decarboxylase test divide into
three portions: make no addition to the first portion, add enough L-lysine dihydrochloride to the
second portion to make a 1% solution, and add L-ornithine dihydrochloride to the third to make
1% (for the Falkow method, add only 0.5% of the L-amino acid). After adding ornithine readjust
pH of the medium to 6.0 ± 0.2. Dispense in 3- to 4-mL portions in screw-capped test tubes and
sterilize by autoclaving at 121°C for 10 min. A floccular precipitate in the ornithine medium
does not interfere with its use.
2) Mineral oil: Use mineral oil sterilized by autoclaving at 121°C for 30 to 60 min depending
on the size of the container.
b. Procedure: Lightly inoculate each of the three media, add a layer of about 10 mm
thickness of mineral oil, and incubate at 37°C for up to 4 d. Examine tubes daily. A color change
from yellow to violet or reddish-violet constitutes a positive decarboxylase test; a change to
bluish gray indicates a weak positive; no color change or a yellow color represents a negative
test. See Table 9225:I.
9. Oxidase Test
The oxidase test determines the presence of oxidase enzymes. Coliform bacteria are
oxidase-negative.
a. Reagents:
1) Media: Use either nutrient agar or tryptic soy agar plates to streak cultures and produce
isolated colonies. From these obtain the inoculum for oxidase testing on impregnated filter
paper. Do not use any medium that includes a carbohydrate in its formulation. Use only tryptic
soy agar if reagent is dropped on colonies. Tryptic soy agar:
Tryptone
Soytone
15.0 g
5.0 g
5.0 g
Agar
Reagent-grade water
15.0 g
1.0 L
pH should be 7.3 ± 0.2 after sterilization.
2) Tetramethyl p-phenylenediamine dihydrochloride, 1% aqueous solution, freshly prepared
or refrigerated for no longer than 1 week. Impregnate a filter paper strip*#(47) with this solution.
Alternatively, prepare a 1% solution of dimethyl p-phenylenediamine hydrochloride. Single-use
reagent ampules, commercially available, are convenient and economical, but use them with
caution. When the reagent is to be dropped directly on colonies, use tryptic soy agar plates
because nutrient agar plates give inconsistent results; when smearing a portion of a picked
colony on reagent-impregnated filter paper, do not transfer any medium with the culture
material.
b. Procedure: Remove some of a colony from agar plate with a platinum wire, a wooden or
plastic applicator stick, or a glass rod and smear on the test strip. Do not use iron or other
reactive wire because it will cause false positive reactions. A dark purple color that develops
within 10 s is a positive oxidase test. Test positive and negative cultures concurrently. If the
liquid reagent is used, drop it on colonies on the culture plate. Oxidase-positive colonies develop
a pink color that successively becomes maroon, dark red, and finally, black.
10. Yellow Pigment
Observe isolated colonies on nutrient agar slants and plates or plates of tryptic soy agar
incubated at 35 ± 0.5°C for up to 48 h. Pigmentation often intensifies as time of incubation
proceeds.
11. Bibliography
MACFADDIN, J.F. 1976. Biochemical Tests for Identification of Medical Bacteria. Williams &
Wilkins Co., Baltimore, Md.
FARMER, J.J., III. 1985. Biochemical identification of new species and biogroups of
Enterobacteriaceae. J. Clin. Microbiol. 21:46.
WASHINGTON, J.A., ed. 1985. Laboratory Procedures in Clinical Microbiology, 2nd ed.
Springer-Verlag, New York, N.Y.
BALOWS, A.H., W.J. HAUSLER, JR., K.L. HERMANN, H.P. ISENBERG & H.J. SHADOMY, eds. 1991.
Manual of Clinical Microbiology, 5th ed. American Soc. for Microbiology, Washington,
D.C.
9230
FECAL STREPTOCOCCUS AND ENTEROCOCCUS GROUPS*#(48)
9230 A.
Introduction
1. Fecal Streptococcus Group
The fecal streptococcus group consists of a number of species of the genus Streptococcus,
such as S. faecalis, S. faecium, S. avium, S. bovis, S. equinus, and S. gallinarum. They all give a
positive reaction with Lancefield’s Group D antisera1 and have been isolated from the feces of
warm-blooded animals. In addition, S. avium sometimes reacts with Lancefield’s Group Q
antisera. S. faecalis subsp. liquefaciens and S. faecalis subsp. zymogenes are differentiated based
on the ability of these strains to liquefy gelatin and hemolyze red cells. However, the validity of
these subspecies is questionable.2,3
The normal habitat of fecal streptococci is the gastrointestinal tract of warm-blooded
animals. S. faecalis and S. faecium once were thought to be more human-specific than other
Streptococcus species. Other species have been observed in human feces but less frequently.4
Similarly, S. bovis, S. equinus, and S. avium are not exclusive to animals, although they usually
occur at higher densities in animal feces.5 Certain streptococcal species predominate in some
animal species and not in others, but it is not possible to differentiate the source of fecal
contamination based on speciation of fecal streptococci.
The fecal streptococci have been used with fecal coliforms to differentiate human fecal
contamination from that of other warm-blooded animals. Editions of Standard Methods previous
to the 17th suggested that the ratio of fecal coliforms (FC) to fecal streptococci (FS) could
provide information about the source of contamination. A ratio greater than four was considered
indicative of human fecal contamination, whereas a ratio of less than 0.7 was suggestive of
contamination by nonhuman sources. The value of this ratio has been questioned because of
variable survival rates of fecal streptococcus group species. S. bovis and S. equinus die off
rapidly, once exposed to aquatic environments, whereas S. faecalis and S. faecium tend to
survive longer.6 Furthermore, disinfection of wastewaters appears to have a significant effect on
the ratio of these indicators, which may result in misleading conclusions regarding the source of
contaminants.7 The ratio is affected also by the methods for enumerating fecal streptococci. The
KF membrane filter procedure has a false-positive rate ranging from 10 to 90% in marine and
fresh waters.8-10 For these reasons, the FC/FS ratio cannot be recommended, and should not be
used as a means of differentiating human and animal sources of pollution.
2. Enterococcus Group
The enterococcus group is a subgroup of the fecal streptococci that includes S. faecalis, S.
faecium, S. gallinarum, and S. avium. The enterococci are differentiated from other streptococci
by their ability to grow in 6.5% sodium chloride, at pH 9.6, and at 10°C and 45°C.
The enterococci portion of the fecal streptococcus group is a valuable bacterial indicator for
determining the extent of fecal contamination of recreational surface waters. Studies at marine
and fresh water bathing beaches indicated that swimming-associated gastroenteritis is related
directly to the quality of the bathing water and that enterococci are the most efficient bacterial
indicator of water quality.11,12 Water quality guidelines based on enterococcal density have been
proposed for recreational waters.13 For recreational fresh waters the guideline is 33
enterococci/100 mL while for marine waters it is 35/100 mL. Each guideline is based on the
geometric mean of at least five samples per 30-d period during the swimming season.
The multiple-tube technique is applicable primarily to raw and chlorinated wastewater and
sediments, and can be used for fresh and marine waters. The membrane filter technique also may
be used for fresh and saline water samples, but is unsuitable for highly turbid waters.
4. References
1. SNEATH, P.H.A., N.S. MAIR, M.E. SHARPE & J.G. HOLT. eds. 1986. Bergey’s Manual of
Systematic Bacteriology. Vol. 2, Williams & Wilkins, Baltimore, Md.
2. JACOB, A.E., G.J. DOUGLAS & S.J. HOBBS. 1973. Self-transferable plasmids determining
the haemolysin and bacteriocin of Streptococcus faecalis var. zymogenes. J. Bacteriol.
121:863.
3. OLIVER, D.R., B.L. BROWN & D.B. CLEWELL. 1977. Characterization of plasmids
determining haemolysin and bacteriocin production in Streptococcus faecalis 5952. J.
Bacteriol. 130:948.
4. WATANABE, T., H. SHIMOHASHI, Y. KAWAI, & M. MUTAI. 1981. Studies on streptococci.
I. Distribution of fecal streptococci in man. Microbiol. Immunol. 25:257.
5. THOMAS, C.D. & M.A. LEVIN. 1978. Quantitative analysis of group D streptococci. Abs.
Annual Meeting, American Soc. Microbiology, p. 210.
6. FEACHAM, R. 1975. An improved role for faecal coliform to faecal streptococci ratios
in the differentiation between human and non-human pollution sources. Water Res.
9:689.
7. ROSSER, P.A.E. & D.P. SARTORY. 1982. A note on the effect of chlorination of sewage
effluents on faecal coliform to faecal streptococci ratios in the differentiation of faecal
pollution sources. Water S.A. 8: 66.
8. FUJIOKA, R.S., A.A. UENO & O.T. NARIKAWA. 1984. Recovery of False Positive Fecal
Streptococcus on KF Agar from Marine Recreational Waters. Tech. Rep. No. 168,
Water Resources Research Center, Univ. Hawaii at Manoa, Honolulu.
9. OLIVIERI, V.P., C.W. KRUSE, K. KAWATA & J.E. SMITH. 1977. Microorganisms in Urban
Stormwater. EPA-600/2-77-087, U.S. Environmental Protection Agency, Edison, N.J.
10. ERICKSEN, T.H., C. THOMAS & A. DUFOUR. 1983. Comparison of two selective
membrane filter methods for enumerating fecal streptococci in freshwater samples.
Abs. Annual Meeting, American Soc. Microbiology, p. 279.
11. CABELLI, V.J. 1983. Health Effects Criteria for Marine Waters. EPA-600/1-80-031,
U.S. Environmental Protection Agency, Cincinnati, Ohio.
12. DUFOUR, A.P. 1984. Health Effects Criteria for Fresh Recreational Waters.
13. U.S. ENVIRONMENTAL PROTECTION AGENCY. 1986. Ambient Water Quality Criteria
for Bacteria—1986. EPA-440/5-84-002, U.S. Environmental Protection Agency,
Washington, D.C.
9230 B.
Multiple-Tube Technique
a. Azide dextrose broth:1
Beef extract
Tryptone or polypeptone
Glucose
Sodium azide, NaN3
4.5
15.0
7.5
7.5
0.2
g
g
g
g
g
1
L
Peptone C
Peptone B
Yeast extract
Bacteriological bile
Sodium citrate
Esculin
Ferric ammonium citrate
Sodium azide, NaN3
17.0
3.0
5.0
10.0
5.0
1.0
1.0
0.5
0.25
g
g
g
g
g
g
g
g
g
Agar
Reagent-grade water
15.0
1
g
L
Reagent-grade water
pH should be 7.2 ± 0.2 at 25°C after sterilization.
b. Pfizer selective enterococcus (PSE) agar:2
pH should be 7.1 ± 0.2 after sterilization. Hold medium for not more than 4 h at 45 to 50°C
before plates are poured.
2. Presumptive Test Procedure
Inoculate a series of tubes of azide dextrose broth with appropriate graduated quantities of
sample. Use sample of 10 mL portions or less. Use double-strength broth for 10-mL inocula. The
portions used will vary in size and number with the sample character. Use only decimal
multiples of 1 mL (see Section 9221 for suggested sample sizes).
Incubate inoculated tubes at 35 ± 0.5°C. Examine each tube for turbidity at the end of 24 ± 2
h. If no definite turbidity is present, reincubate, and read again at the end of 48 ± 3 h.
3. Confirmed Test Procedure
Subject all azide dextrose broth tubes showing turbidity after 24- or 48-h incubation to the
confirmed test.
Streak a portion of growth from each positive azide dextrose broth tube on PSE agar.
Incubate the inverted dish at 35 ± 0.5°C for 24 ± 2 h. Brownish-black colonies with brown halos
confirm the presence of fecal streptococci.
Brownish-black colonies with brown halos may be transferred to a tube of brain-heart
infusion broth containing 6.5% NaCl. Growth in 6.5% NaCl broth and at 45°C indicates that the
colony belongs to the enterococcus group.
4. Computing and Recording of MPN
Estimate fecal streptococci densities from the number of tubes in each dilution series that are
positive on PSE agar. Similarly, estimate enterococci densities from the number of tubes in each
dilution series containing streptococci that can grow in 6.5% NaCl broth. Compute the
combination of positives and record as the most probable number (MPN). Refer to Section
9221D.
5. References
1. MALLMAN, W.L. & E.B. SELIGMANN. 1950. A comparative study of media for the
detection of streptococci in water and sewage. Amer. J. Pub. Health 40:286.
2. ISENBERG, H.D., D. GOLDBERT & J. SAMPSON. 1972. Laboratory studies with a selective
enterococcus medium. Health Lab. Sci. 9:289.
9230 C.
Membrane Filter Techniques
a. mE agar for enterococci:1
Peptone
Yeast extract
Esculin
Actidione (cycloheximide)
Sodium azide, NaN3
10.0
15.0
30.0
1.0
0.05
0.15
g
g
g
g
g
g
Agar
Reagent-grade water
15.0
1
g
L
Heat to dissolve ingredients, sterilize, and cool in a water bath at 44 to 46°C. Mix 0.25 g
nalidixic acid in 5 mL reagent-grade water, add a few drops of 0.1N NaOH to dissolve the
antibiotic, and add to the basal medium. Add 0.15 g 2,3,5-triphenyl tetrazolium chloride and mix
well to dissolve. Pour the agar into 9- × 50-mm petri dishes to a depth of 4 to 5 mm
(approximately 4 to 6 mL), and let solidify. The final pH should be 7.1 ± 0.2. Store poured plates
in the dark at 2 to 10°C. Discard after 30 d. (NOTE: This medium is recommended for culturing
enterococci in fresh and marine recreational waters.)
b. EIA substrate:1
Esculin
Ferric citrate
Agar
Reagent-grade water
1.0
0.5
15.0
1
g
g
g
L
The pH should be 7.1 ± 0.2 before autoclaving. Heat to dissolve ingredients, sterilize, and
cool in a water bath at 44 to 46°. Pour medium into 50-mm petri dishes to a depth of 4 to 5 mm
(approximately 4 to 6 mL) and let solidify. Store poured plates in the dark at 2 to 10°C. Discard
after 30 d.
c. m Enterococcus agar for fecal streptococci:2
Tryptose
Yeast extract
Glucose
Dipotassium phosphate, K2HPO4
Sodium azide, NaN3
20.0
5.0
2.0
4.0
g
g
g
g
0.4 g
2,3,5-Triphenyl tetrazolium chloride
Agar
Reagent-grade water
0.1 g
10.0 g
1 L
Heat to dissolve ingredients. Do not autoclave. Dispense into 9- × 50-mm petri plates to a
depth of 4 to 5 mm (approximately 4 to 6 mL), and let solidify. Prepare fresh medium for each
set of samples. (NOTE: This medium is recommended for Group D streptococci in fresh and
marine waters.)
d. Brain-heart infusion broth:
Infusion of calf brain
Infusion of beef heart
Proteose peptone
Glucose
Disodium hydrogen phosphate, Na2HPO4
200
250
10.0
2.0
5.0
2.5
g
g
g
g
g
g
1
L
Reagent-grade water
The pH should be 7.4 after sterilization.
e. Brain-heart infusion agar: Add 15.0 g agar to the ingredients for brain-heart infusion
broth. The pH should be 7.4 after sterilization. Tube for slants.
f. Bile esculin agar:3
Beef extract
Peptone
Oxgall
Esculin
Ferric citrate
Agar
Reagent-grade water
3.0
5.0
40.0
1.0
0.5
15.0
1
g
g
g
g
g
g
L
Heat to dissolve ingredients. Dispense 8 to 10 mL into tubes for slants or an appropriate
volume into a flask for subsequent pouring into plates. Autoclave at 121°C for 15 min. Do not
overheat because this may cause darkening of the medium. Cool to 44 to 46°C and slant the
tubes or dispense 15 mL into 15- × 100-mm petri dishes. The final pH should be 6.6 ± 0.2 after
sterilization. Store at 4 to 10°C.
3. Procedures
a. mE Method:1
1) Selection of sample size and filtration—Filter appropriate sample volumes through a
0.45-µm, gridded, sterile membrane to give 20 to 60 colonies on the membrane surface. Transfer
filter to agar medium in petri dish, avoiding air bubbles beneath the membrane.
2) Incubation—Invert culture plates and incubate at 41°C ± 0.5°C for 48 h.
3) Substrate test—After 48 h incubation, carefully transfer filter to EIA medium. Incubate at
41°C ± 0.5°C for 20 min.
4) Counting—Pink to red enterococci colonies develop a black or reddish-brown precipitate
on the underside of the filter. Count colonies using a fluorescent lamp and a magnifying lens.
b. m Enterococcus method:2
1) Selection of sample size and filtration—See ¶ 3a.
2) Incubation—Let plates stand for 30 min, then invert and incubate at 35 ± 0.5°C for 48 h.
3) Counting—Count all light and dark red colonies as enterococci. Count colonies using a
fluorescent lamp and a magnifying lens.
4. Calculation of Fecal Streptococci or Enterococci Density
Compute density from sample quantities producing membrane filter counts within the
desired 20- to 60-fecal streptococcus or enterococcus colony range. Calculate as in Section
9222B.6. Record densities as fecal streptococci or enterococci per 100 mL.
5. Verification Tests
Pick selected typical colonies from a membrane and streak for isolation onto the surface of a
brain-heart infusion agar plate. Incubate at 35°C ± 0.5°C for 24 to 48 h.
Transfer a loopful of growth from a well-isolated colony on brain-heart infusion agar into a
brain-heart infusion broth tube and to each of two clean glass slides. Incubate the brain-heart
infusion broth at 35 ± 0.5°C for 24 h. Add a few drops of freshly prepared 3% hydrogen
peroxide to the smear on a slide. The appearance of bubbles constitutes a positive catalase test
and indicates that the colony is not a member of the fecal streptococcus group. If the catalase test
is negative, i.e., no bubbles, make a Gram stain of the second slide. Fecal streptococci and
enterococci are gram-positive, ovoid cells, 0.5 to 1.0 µm in diameter, mostly in pairs or short
chains.
Transfer a loopful of growth from the brain-heart infusion broth to each of the following
media: bile esculin agar (incubate at 35 ± 0.5°C for 48 h); brain-heart infusion broth (incubate at
45 ± 0.5°C for 48 h); brain-heart infusion broth with 6.5% NaCl (incubate at 35 ± 0.5°C for 48
h).
Growth of catalase-negative, gram-positive cocci on bile esculin agar and at 45°C in
brain-heart infusion broth verifies that the colony is of the fecal streptococcus group. Growth at
45°C and in 6.5% NaCl broth indicates that the colony belongs to the enterococcus group.
6. Serological Verification of Group D Fecal Streptococci
An alternate verification test for Group D streptococci can be performed using the precipitin
method of Lancefield.4 This test is highly specific for S. faecalis, S. faecium, S. avium, S.
gallinarum, S. bovis, and S. equinus.
a. Antigen preparation: Pick typical single colonies from the membrane filter and streak for
isolation on brain-heart infusion agar or blood agar plates. Pick a well-isolated colony and
inoculate into 30 to 50 mL of Todd-Hewitt broth.5 Incubate at 35°C for 24 h under aerobic
conditions. Concentrate bacterial suspension by centrifuging (3000 × g for 5 min). Draw
supernatant off and resuspend cells in 0.5 mL saline solution. Autoclave resuspended cells for 15
min at 121°C. Centrifuge the bacteria and decant clear supernatant fluid containing the group
antigen.
b. Capillary precipitin test: Antisera for this test may be obtained from commercial sources.
Dip a 1.2- to 1.5-mm-OD capillary tube into antiserum and draw up about 1 cm of serum.
Place a finger over upper end of tube so that no air will be drawn up and carefully wipe off
excess antiserum. Dip tube into streptococcal antigen extract solution and draw up an equal
volume of antigen. Carefully wipe off excess extract. Place a finger over upper end of tube and
force lower end into plasticine to plug lower opening. Invert tube and place it in plasticine
groove of a capillary holding rack.
A positive test for Group D antigen is characterized by a white precipitate that appears at the
antigen-antiserum interface within 15 min and usually by 5 min. If no reaction has occurred by
30 min, the test is negative. Examination of the tubes is more effective if they are read in a bright
light against a dark background.
Serological verification of Group D streptococci also can be done using commercially
available agglutination tests.*#(49) The slide agglutination tests are simple and appear to be
reliable. Group D streptococci are verified directly from isolated colonies on membrane filter
plates or from broth culture tubes. To verify presumptive enterococci cultures, test also for salt
tolerance (growth in 6.5% NaCl broth).
7. Identification of Individual Species within Fecal Streptococcus and Enterococcus
Groups
Table 9230:I shows some of the key biochemical reactions for identifying fecal streptococci,
enterococci, and species within these two groups.
8. References
1. LEVIN, M.A., J.R. FISCHER & V.J. CABELLI. 1975. Membrane filter technique for
enumeration of enterococci in marine waters. Appl. Microbiol 30:66.
2. SLANETZ, L.W. & C.H. BARTLEY. 1957. Numbers of enterococci in water, sewage and
3.
4.
5.
6.
7.
8.
9.
feces determined by the membrane filter technique with an improved medium. J.
Bacteriol. 74:591.
PHILLIPS, E. & P. NASH. 1985. Culture media. In E.H. Lennette, A. Ballows, W.J.
Hausler, Jr. & H.J. Shadomy, eds. Manual of Clinical Microbiology, 5th ed. American
Soc. Microbiology, Washington, D.C.
LANCEFIELD, R.C. 1933. A serological differentiation of human and other groups of
hemolytic streptococci. J. Exp. Med. 57:571.
DIFCO LABORATORIES. 1984. Difco Manual, 10th ed.
FACKLAM, R.R. & R.B. CAREY. 1985. Streptococci and aerococci. In E.H. Lennette, A.
Ballows, W.J.Hausler, Jr. & H.J. Shadomy, eds. Manual of Clinical Microbiology, 5th
ed. American Soc. Microbiology, Washington, D.C.
GROSS, J.C., M.P. HOUGHTON & L.B. SENTERFIT. 1975. Presumptive speciation of
Streptococcus bovis and other Group D streptococci from human sources by using
arginine and pyruvate tests. J. Clin. Microbiol. 1:54.
COWAN, S.T. & K.J. STEEL. 1965. Manual for the Identification of Medical Bacteria.
Cambridge Univ. Press, Cambridge, England.
BRIDGE, P.D. & P.H.A. SNEATH. 1983. Numerical taxonomy of streptococcus. J. Gen.
Microbiol. 129:565.
9240
IRON AND SULFUR BACTERIA*#(50)
9240 A.
Introduction
The group of nuisance organisms collectively designated ‘‘iron and sulfur bacteria’’ is
morphologically and physiologically heterogeneous, having in common the ability to transform
or deposit significant amounts of iron or sulfur, usually in the form of objectionable slimes.
However, iron and sulfur bacteria are not the sole producers of bacterial slimes and in some
cases may be associated with slimes of other bacteria.
The iron and sulfur bacteria may be filamentous or single-celled, autotrophic or
heterotrophic, aerobic or anaerobic. The taxonomic position of these bacteria is very diverse.
They are studied as iron and sulfur bacteria, because these elements and their transformations
may be important in water treatment and distribution systems and may be especially bothersome
in waters for industrial use, such as cooling and boiler waters. Iron bacteria may cause, or be
associated with, fouling and plugging of wells. Also, the growth of these bacteria may result in
consumer complaints of red water in distribution systems for potable water and sulfate-reducing
bacteria may cause rusty water and tuberculation of pipes. These organisms also may cause odor,
taste, frothing, color, and increases in turbidity in waters.
The nutrient supply for iron and sulfur bacteria may be wholly or partly inorganic. Many of
these bacteria can grow under oligotrophic conditions when attached to a substrate in flowing
water. This seems quite important in the case of certain sulfur bacteria utilizing small amounts of
hydrogen sulfide or in the case of organisms such as Gallionella, which obtain their energy from
the oxidation of ferrous iron. Thiobacillus ferrooxidans contributes to the problem of acid mine
drainage and can be identified by tests for transformation of ferrous to ferric iron or oxidation of
reduced sulfur compounds under conditions of low pH. Temperature, light, pH, and oxygen
supply are critical to the growth of iron and sulfur bacteria. Under different environmental
conditions some bacteria may appear either as iron or as sulfur bacteria.
9240 B.
Iron Bacteria
1. General Characteristics
‘‘Iron bacteria’’ are considered to be capable of metabolizing reduced iron present in their
aqueous habitat and of depositing it in the form of hydrated ferric oxide on or in their
mucilaginous secretions. A somewhat similar mechanism is used by bacteria that utilize
manganese. The large amount of brown slime so produced will impart a reddish tinge and an
unpleasant odor to drinking water and may render the supply unsuitable for domestic or
industrial purposes. These bacteria obtain energy by the oxidation of iron from the ferrous to the
ferric state; the ferric form is precipitated as ferric hydroxide [Fe(OH)3]. Iron may be obtained
from the pipe itself if it is made of iron or from the water within it. Because of the energetics
involved, the amount of ferric hydroxide deposited is very large in comparison with the enclosed
cells.
Some bacteria that do not oxidize ferrous iron may cause it to be dissolved or deposited
indirectly. In their growth, either they liberate iron by utilizing organic radicals to which the iron
is attached or they alter environmental conditions to permit the solution or deposition of iron.
Under these conditions, less ferric hydroxide may be produced, but taste, odor, and fouling may
be engendered.
2. Collection of Samples and Identification
Examples of iron bacteria are shown in Figure 9240:1, Figure 9240:2, Figure 9240:3, Figure
9240:4, and Figure 9240:5. Identification of nuisance iron bacteria usually has been made on the
basis of microscopic examination of the suspected material. Directly examine bulked activated
sludge, masses of microbial growth in lakes, rivers, and streams, and slime growths in
cooling-tower waters. Suspected development of iron bacteria in water wells or in distribution
systems may require special efforts to secure samples useful for identification.
Settle or centrifuge samples drawn directly from wells and examine sediment
microscopically. Place a portion of sediment on a microscope slide, cover with a cover slip, and
examine under a low-power microscope for filaments and iron-encrusted filaments. The material
trapped by filters placed in front of back-surge valves often has yielded excellent specimens of
iron bacteria. Water pumped from wells may be passed through a 0.45-µm membrane filter and
the filter examined microscopically after drying and clearing with immersion oil applied directly
to the membrane. Phase-contrast microscopes have made possible the examination of unstained
culture material. Use india ink or lactophenol blue for staining when conventional light
microscopy is used. Also, iron bacteria have been observed when the epifluorescence
microscopic method (Section 9216B) was used.
Continued heavy deposition of iron caused by the oxidation of ferrous iron by air or by other
environmental changes often hides the sheaths or stalks of iron bacteria. The cells within the
filaments often die and disintegrate and the filaments tend to be fragmented or crushed by the
mass of the iron precipitate.
To dissolve iron deposits place several drops of 1N HCl at one edge of cover slip and draw it
under the cover slip by applying filter or blotting paper to the opposite edge. Reducing
compounds such as sodium ascorbate also may be used to dissolve deposits and permit
observation of cellular structure. To verify that the material is iron, add a solution of potassium
ferrocyanide to a sample on a slide, cover, and draw 1N HCl under cover slip. A blue precipitate
of Prussian blue will form as iron around cells or filaments is dissolved.
Floating or hanging slide methods may be used to determine the presence of filamentous and
other periphytic iron bacteria. In the floating method fix a glass microscope slide in a cork and
let it float for 1 to 2 d on the surface of water taken from the source. Slides may be hung by
means of string at various depths in the water for various times before being removed and
observed microscopically for the presence of iron bacteria.1 A cultural method called the iron
bacteria presence test has been developed for the detection of iron bacteria.2
Identify organisms by comparing with available drawings or photographs of iron
bacteria.1-17
3. References
1. WOJCIK, W. & M. WOJCIK. 1986. Monitoring biofouling. In D.R. Cullimore, ed. Proc.
International Symposium on Biofouled Aquifers: Prevention and Restoration.
American Water Resources Assoc., Bethesda, Md.
2. CULLIMORE, D.R. & A.E. MCCANN. 1977. The identification, cultivation and control of
iron bacteria in ground water. In F.A. Skinner & J.M. Shewan, eds. Aquatic
Microbiology. Academic Press, New York, N.Y.
3. LUESCHOW, L.A. & K.M. MACKENTHUN. 1962. Detection and enumeration of iron
bacteria in municipal water supplies. J. Amer. Water Works Assoc. 54:751.
4. STARKEY, R.L. 1945. Transformations of iron by bacteria in water. J. Amer. Water
5. STOKES, J.L. 1954. Studies on the filamentous sheathed iron bacterium Sphaerotilus
natans. J. Bacteriol. 67:278.
6. KUCERA, S. & R.S. WOLFE. 1957. A selective enrichment method for Gallionella
7.
8.
9.
10.
11.
ferruginea. J. Bacteriol. 74:344.
WAITZ, S. & J.B. LACKEY. 1958. Morphological and biochemical studies on the
organism Sphaerotilus natans. Quart. J. Fla. Acad. Sci. 21:335.
WOLFE, R.S. 1958. Cultivation, morphology, and classification of the iron bacteria. J.
WOLFE, R.S. 1960. Observations and studies of Crenothrix polyspora. J. Amer. Water
WOLFE, R.S. 1960. Microbial concentration of iron and manganese in water with low
concentrations of these elements. J. Amer. Water Works Assoc. 52:1335.
DONDERO, N.C., R.A. PHILIPS & H. HEUKELEKIAN. 1961. Isolation and preservation of
cultures of Sphaerotilus. Appl. Microbiol. 9:219.
12. MULDER, E.G. 1964. IRON BACTERIA, PARTICULARLY THOSE OF THE
Sphaerotilus-Leptothrix group, and industrial problems. J. Appl. Bacteriol. 27:151.
13. DRAKE, C.H. 1965. Occurrence of Siderocapsa treubii in certain waters of the
Niederrhein. Gewasser Abwasser 39/40:41.
14. STALEY, J.T., M.P. BRYANT, N. PFENNING & J.G. HOLT, eds. 1989. Bergey’s Manual of
Systematic Bacteriology, Volume 3. Williams & Wilkins, Baltimore, Md.
15. EDMONDSON, W.T., ed. 1959. Ward & Whipple’s Fresh Water Biology, 2nd ed. John
Wiley & Sons, New York, N.Y.
16. SKERMAN, V.B.D. 1967. A Guide to the Identification of the Genera of Bacteria, 2nd ed.
Williams & Wilkins, Baltimore, Md.
17. GHIORSE, W.C. 1984. Biology of iron and manganese depositing bacteria. Annu. Rev.
Microbiol. 38:515.
9240 C.
Sulfur Bacteria
1. General Characteristics
The bacteria that oxidize or reduce significant amounts of inorganic sulfur compounds
exhibit a wide diversity of morphological and biochemical characteristics. One group, the
sulfate-reducing bacteria, consists mainly of single-celled forms that grow anaerobically and
reduce sulfate, SO42−, to hydrogen sulfide, H2S. One member of this group, Desulfonema, is
multicellular and exhibits gliding motility. A second group, the photosynthetic green and purple
sulfur bacteria, grows anaerobically in the light and uses H2S as a hydrogen donor for
photosynthesis. Members of a third colorless filamentous group are myxotrophic and utilize
organic sources of carbon but may get their energy from the oxidation of reduced sulfur
compounds. The sulfide is oxidized to sulfur or sulfate. A fourth group, the aerobic
sulfur-oxidizers, oxidizes reduced sulfur compounds aerobically to obtain energy for
chemoautotrophic growth.
The sulfur bacteria of most importance in the water and wastewater field are the
sulfate-reducing bacteria, which include Desulfovibrio, and the single-celled aerobic
sulfur-oxidizers of the genus Thiobacillus. The sulfate-reducing bacteria contribute greatly to
tuberculations and galvanic corrosion of water mains and to taste and odor problems in water.
Thiobacillus, by its production of sulfuric acid, has contributed to the destruction of concrete
sewers and the acid corrosion of metals.
2. Collection of Samples and Identification
Identification of nuisance sulfur bacteria usually has been made on the basis of microscopic
examination of the suspected material. Examine samples of slimes suspended in waters,
scrapings from exposed surfaces, or sediments directly.
Three groups of sulfur bacteria may be recognized microscopically; green and purple sulfur
bacteria; large, colorless filamentous sulfur bacteria; and large, colorless, nonfilamentous sulfur
bacteria. The fourth group, consisting of sulfate-reducing bacteria and sulfur-oxidizing bacteria
of the genus Thiobacillus, cannot be identified by appearance alone.
a. Green and purple sulfur bacteria:
1) Green sulfur bacteria most frequently occur in waters containing H2S. They are small,
ovoid to rod-shaped, nonmotile organisms, generally less than 1 µm in diameter, and with a
yellowish-green color in masses. Sulfur globules are seldom if ever deposited within the cells.
2) Purple sulfur bacteria (Figure 9240:6) occur in waters containing H2S. They are large,
generally stuffed with sulfur globules, and often so intensely pigmented as to make individual
cells appear red. Large, dense, highly colored masses are detected easily by the naked eye. The
presence of photosynthetic bacteria in concentrated masses can be confirmed by extracting the
mass with ether and scanning the extract absorbance in the infrared region. Bacterial chlorophyll
will absorb strongly in the range of 660 to 870 nm.
b. Colorless filamentous sulfur bacteria: Colorless filamentous sulfur bacteria (Figure
9240:7, Figure 9240:8, and Figure 9240:9) occur in waters where both oxygen and H2S are
present. They may form mats with a slightly yellowish-white appearance due to deposition of
internal sulfur globules. They generally are large and may be motile with a characteristic gliding
movement. Identify by comparing organisms with available photographs.1-4
c. Colorless nonfilamentous sulfur bacteria: Colorless, nonfilamentous sulfur bacteria (see
Figure 9240:10, for example) usually are associated with decaying algae. They are extremely
motile, ovoid to rod-shaped with sulfur globules and possible calcium carbonate deposits. They
generally are very large.
d. Colorless small sulfur bacteria and sulfate-reducing bacteria: The small single-celled
bacteria, Thiobacillus spp., and the sulfate-reducing bacteria, such as Desulfovibrio, cannot be
identified by direct microscopic examination. Thiobacillus types are small, colorless, motile, and
rod-shaped and are found in an environment containing H2S. Sulfur globules are absent. Identify
Thiobacillus types, Desulfovibrio, or other sulfate-reducing bacteria physiologically.
3. References
1. LACKEY, J.B. & E.W. LACKEY. 1961. The habitat and description of a new genus of
sulfur bacterium. J. Gen. Microbiol. 26:28.
2. FAUST, L. & R.S. WOLFE. 1961. Enrichment and cultivation of Beggiatoa alba. J.
Bacteriol. 81:99.
3. MORGAN, G.B. & J.B. LACKEY. 1965. Ecology of a sulfuretum in a semitropical
environment. Z. Allg. Mikrobiol. 5:237.
4. STALEY, J.T., M.P. BRYANT, N. PFENNING & J.G. HOLT, eds. 1989. Bergey’s Manual of
9240 D.
Enumeration, Enrichment, and Isolation of Iron and Sulfur Bacteria
There are no good means of enumerating iron and sulfur bacteria other than the
sulfate-reducing bacteria and the thiobacilli. Laboratory cultivation and isolation of pure cultures
is difficult and successful isolation is uncertain. This is especially true of attempts to isolate
filamentous bacteria from activated sludge or other sources where many different bacterial types
are present.
1. Media
a. Casitone-glycerol-yeast autolysate broth (CGY), for the Sphaerotilus group: This medium
may not be available in dehydrated form and may require preparation from the basic ingredients.
It may be solidified by adding 1.5% agar.
Casitone
Glycerol
Yeast autolysate
Reagent-grade water
5.0
10.0
1.0
1
g
mL
g
L
b. Isolation medium (iron bacteria): This medium may not be available in dehydrated form
and may require preparation from the basic ingredients.
Glucose
Ammonium sulfate, (NH4)2SO4
0.15 g
0.5 g
Calcium nitrate, Ca(NO3)2
0.01 g
0.05 g
0.05 g
Potassium chloride, KCl
Calcium carbonate, CaCO3
0.05 g
0.1 g
Agar
Cyanocobalamin
Thiamine
Reagent-grade water
10.0
0.01
0.4
1
g
mg
mg
L
c. Maintenance (SCY) medium (iron bacteria): This medium may not be available in
dehydrated form and may require preparation from the basic ingredients.
Sucrose
Casitone
Yeast extract
Trypticase soy broth without dextrose
Agar
Cyanocobalamin
Thiamine
Reagent-grade water
1.0
0.75
0.25
0.25
10.0
0.01
0.4
1
g
g
g
g
g
mg
mg
L
d. Mn agar No. 1: This medium may not be available in dehydrated form and may require
Manganous carbonate, MnCO3
2.0
g
Beef extract
Ferrous ammonium sulfate, Fe(NH4)2(SO4)2
1.0
150
g
mg
Sodium citrate
Yeast extract
Cyanocobalamin
Agar
Reagent-grade water
150
75
0.005
10.0
1
mg
mg
mg
g
L
Prepare and sterilize the medium without cyanocobalamin. Separately sterilize cobalamin by
filtration and add aseptically just before medium solidifies.
e. Mn agar No. 2:1 Prepare fresh each time from basic ingredients:
Manganous sulfate, MnSO4⋅H2O
10
Agar
Natural water
15.0 g
1 L
mg
f. Iron oxidizing medium (Thiobacillus ferrooxidans): This medium may not be available in
Basal salts:
3.0 g
Potassium chloride, KCl
0.10 g
0.50 g
0.50 g
Calcium nitrate Ca(NO3)2
0.01 g
H2SO4, 10N
1.0 mL
Reagent-grade water
700
mL
Ferrous sulfate, FeSO4⋅7H2O, 14.74% solution (w/v) 300
mL
Energy source:
Separately sterilize basal salts and energy source and combine when cool. Store in the
refrigerator and discard after 2 weeks. A precipitate will form and the medium will be opalescent
and green. The pH should be 3.0 to 3.6.
g. Ferrous sulfide agar (Gallionella ferruginea): This medium may not be available in
Agar layer:
Ferrous sulfide, FeS (washed precipitate and liquid)
Sodium sulfide, Na2S
500
mL
15.6 g
Ferrous ammonium sulfate, Fe(NH4)2(SO4)2⋅6H2O
78.4 g
Boiling reagent-grade water
Agar (liquid) (30 g/L)
1
500
L
mL
Liquid overlay:
Ammonium chloride, NH2Cl
1.0 g
0.5 g
0.2 g
Calcium chloride, CaCl2
0.1 g
Reagent-grade water
1
L
Prepare FeS by reacting equal molar quantities of Na2S and Fe(NH4)2(SO4)2 in boiling
reagent-grade water. Let precipitate settle from the hot solution in a completely filled and
stoppered bottle. Wash precipitate four times by decanting supernatant and replacing with
boiling water. Store FeS in a glass stoppered bottle completely filled with additional boiling
water.
Add equal volumes of FeS and 3% agar at 45°C. Prepare slants in screw-capped tubes.
Prepare liquid overlay, bubble CO2 through it for 10 to 15 s, and add several milliliters to agar
slant. Modifications to this medium and procedure are available.2-4
A variation of the basic medium requires adding 0.5 mL formalin (40% formaldehyde
solution) to a screw-capped dilution bottle containing 10 mL FeS agar and 100 mL liquid
overlay. This variation is said not to work in all cases.3,4 Add 0.001% bromthymol blue and
0.004% bromcresol purple to liquid overlay.
h. Sulfate-reducing medium: This medium may not be available in dehydrated form and may
require preparation from the basic ingredients.
Sodium lactate
Beef extract
Peptone
3.5
1.0
2.0
2.0
g
g
g
g
Sodium sulfate, Na2SO4
1.5
g
0.5
g
Ferrous ammonium sulfate, Fe(NH4)2(SO4)2⋅6H2O
0.392 g
Calcium chloride, CaCl2
0.10
g
Sodium ascorbate
Reagent-grade water
0.10
1
g
L
pH should be 7.5 ± 0.3 after sterilization. Prepare medium excluding ferrous ammonium
sulfate and sodium ascorbate, dispense in screw-capped test tubes, and sterilize. For use, the
tubes must be completely filled; therefore, in a flask sterilize extra medium to be added to tubes
for filling. On day of use, prepare separate solutions of ferrous ammonium sulfate (3.92 g/100
mL) and sodium ascorbate (1.00 g/100 mL), sterilize by filtration through a 0.45-µm membrane
filter, and aseptically add 0.1 mL each solution/10 mL basal medium.
i. Thiosulfate oxidizing medium (Thiobacillus thioparus): This medium may not be available
in dehydrated form and may require preparation from the basic ingredients.
Sodium thiosulfate, Na2S2O3⋅5H2O
10.0 g
2.0 g
0.1 g
Calcium chloride, CaCl2⋅2H2O
0.1 g
0.1 g
Ferric chloride, FeCl3⋅6H2O
0.02g
Reagent-grade water
1
L
pH should be 7.8 after sterilization. Separately sterilize Na2S2O3 and (NH4)2SO4 and add
before use. If this medium is used to isolate Thiobacillus thioparus, check isolates to ensure they
are autotrophs.
j. Sulfur medium (Thiobacillus thiooxidans): This medium may not be available in
Sulfur, elemental
10.0
3.0
g
g
0.5
g
0.3
g
0.25 g
Ferric chloride, FeCl3⋅6H2O
0.02 g
Reagent-grade water
1
L
pH should be 4.8 after sterilization. Weigh sulfur into 250-mL flasks using 1 g/flask. Add
100 mL medium to each flask and sterilize with intermittent steam (30 min for each of 3
consecutive d).
k. Media for Beggiatoa and myxotrophic strains of Thiothrix:
1) Extracted hay: Extract hay or grass at least five times by boiling in water for 30 min, with
two rinses in cold water between each extraction. Dry the extracted hay, place 10 to 30 blades in
a large test tube, and sterilize by autoclaving.
2) Basal medium
Ammonium chloride, 4% solution
Dipotassium hydrogen phosphate, K2HPO4, 1% solution
5 mL
1 mL
Magnesium sulfate, MgSO4⋅7H2O, 1% solution
1 mL
Calcium sulfate, CaSO4⋅2H2O, saturated solution
20 mL
Trace elements [see ¶ 3) below]
Reagent-grade water
5 mL
968 mL
3) Trace elements
Reagent-grade water
920 mL
10 mL
Zinc sulfate, ZnSO4⋅7H2O, 0.1% solution
Manganous sulfate, MnSO4⋅4H2O, 0.02% solution
10 mL
Copper sulfate, CuSO4⋅5H2O, 0.00005% solution
10 mL
Boric acid, H3BO3, 0.1% solution
10 mL
Cobalt nitrate, Co(NO3)2 or CoCl2⋅6H2O, 0.01% solution
10 mL
Sodium molybdate, Na2MoO4⋅2H2O, 0.01% solution
10 mL
EDTA solution, 2% EDTA with 7% ferrous sulfate,
FeSO4⋅7H2O, with 1 mL conc HCl/100 mL
20 mL
4) MP agar: This medium may not be available in dehydrated form and should be made fresh
before use.
Basal medium
Sodium acetate
Sodium sulfide solution [see ¶ 5) below]
Agar
1
0.1
3
15
L
g
mL
g
Adjust pH to between 7.0 and 7.5 and heat to dissolve agar. Sterilize by autoclaving for no
more than 15 min and cool in a 45 to 50°C water bath. Add sodium sulfide solution immediately
before pouring plates. If the medium is to be used for screw-capped tubes the sulfide may be
added before autoclaving.
5) Sodium sulfide solution: Make up and separately autoclave a 10% solution of Na2S⋅9H2O.
6) MY agar: This medium may not be available in dehydrated form and should be made fresh
before use.
Basal medium
Sodium acetate
Nutrient broth powder (Difco)
Yeast extract (Difco)
Sodium sulfide solution [see ¶ 5) above]
Agar
1
0.1
0.1
0.1
3
15
L
g
g
g
mL
g
Adjust pH to between 7.0 and 7.5 and heat to dissolve agar. Sterilize by autoclaving for no
more than 15 min and cool in a 45 to 50°C water bath. Add sodium sulfide solution immediately
before pouring plates. If the medium is to be used for screw-capped tubes the sulfide may be
added before autoclaving.
l. Media for heterotrophic strains of Thiothrix: Make the media 3) through 9) with solutions
1) and 2), plus indicated additives. After adding all ingredients, adjust pH to 7.2 to 7.5. For solid
media add 12 g agar/L.
1) MSV
0.5
g
0.11
g
0.085 g
0.1
g
0.05
g
Ferric chloride, FeCl3⋅H2O
0.002 g
EDTA
Vitamin mix
Reagent-grade water
0.003 g
1
mL
1
L
2) Vitamin mix
Calcium pantothenate
Niacin
Biotin
Cyanocobalamin
Folic acid
Pyridoxine
p-aminobenzoic acid
Cocarboxylase
Inositol
Thiamine
Riboflavin
Reagent-grade water
0.01
0.01
0.0005
0.0005
0.0005
0.01
0.01
0.01
0.01
0.01
0.01
100
g
g
g
g
g
g
g
g
g
g
g
mL
3) AcS
Sodium acetate
Sodium sulfide, Na2S⋅9H2O
0.15 g
0.187 g
MSV
1
Sucrose
0.15 g
0.187 g
MSV
1
Glucose
0.15 g
0.187 g
MSV
1
L
4) SS
L
5) GS
L
6) SUC
Sodium succinate
MSV
0.15 g
1
L
7) I
Glucose
MSV
0.15 g
1
L
8) S
0.187 g
MSV
1
L
9) LT
Sodium lactate
Sodium thiosulfate, Na2S2O3
0.5 g
0.5 g
MSV
1
L
2. Iron Bacteria
a. Sphaerotilus-Leptothrix:
1) Iron bacteria, especially those belonging to the Sphaerotilus-Leptothrix group, thrive in
media too dilute to support the proliferation of more rapidly growing organisms. One medium5 is
partially selective for Sphaerotilus (BOD dilution water, Section 5210B, supplemented with 100
mg/L sodium lactate). Dispense 50 mL of this medium into French square bottles and autoclave
at 69 kPa for 15 min. To inoculate sample add 25-mL portions of stream water or 1-, 5-, and
10-mL portions of settled wastewater or process liquor to duplicate bottles of medium. Incubate
at 22 to 25°C for 5 d and observe for filamentous growth. Isolate pure cultures by picking a
filament from the BOD-lactate broth and streaking on 0.05% meat extract agar. After incubating
for 24 h at 25°C, pick typical curling filaments with the aid of a dissecting microscope and
transfer to casitone-glycerol-yeast autolysate (CGY) broth. If a pellicle with no underlying
turbidity develops in 2 to 3 d, transfer a filament to a CGY agar slant, incubate at 25°C until
growth is visible, and store in a refrigerator. In addition, extracted alfalfa straw or pea straw may
be used for enrichments.4
A detailed key for identifying filamentous microorganisms in complex mixtures such as
wastewater and activated sludge is available.6
2) Isolation and maintenance media have proven quite successful for identifying various
groups of filamentous organisms, including iron bacteria.7 Prepare agar slants of these media and
aseptically pipet 3 mL sterile, dechlorinated tap water onto surface of slants. Inoculate tubes and
incubate at room temperature until turbid growth has developed in liquid layer. The cells will
remain viable for 3 months in the refrigerator.
3) Another good maintenance medium for cultivating the Sphaerotilus group is CGY.8
4) Leptothrix (Sphaerotilus discophorous) can be distinguished from Sphaerotilus natans by
its ability to oxidize manganous ion. Use Mn agar No. 1 as the differential medium.9
Alternatively, Leptothrix may be grown by direct plating on Mn agar No. 2.1
b. Thiobacillus ferrooxidans: Although this organism also is a sulfur-oxidizing
bacterium,10,11 its main importance has been in acid mine drainage. A medium suitable for
enumeration of the MPN is available.12 Some oxidation of iron occurs during sterilization but
the loss of ferrous iron is not appreciable. The medium has a precipitate (probably ferrous and
ferric phosphates), is opalescent and green, has a pH of 3.0 to 3.6, and contains 9000 mg/L
ferrous iron. Growth of the organism is manifested by a decrease in pH and an increase in
concentration of oxidized iron. With practice and use of uninoculated controls, an increase of
deep orange-brown color can be seen in positive enrichment tubes or flasks as compared to
negative ones. Shake test-tube dilutions daily because these organisms are highly aerobic.
c. Gallionella ferruginea: For cultivation of this organism use ferrous sulfide agar.13,14
Inoculate tubes with a drop of suspension of a suspected Gallionella deposit. Growth at room
temperature usually occurs in 18 to 36 h and appears as a white deposit on sides of test tube. The
ring of colonies occurring at a certain level reflects a balance between upward diffusion of
ferrous ions and downward diffusion of oxygen molecules. Supplementation of ferrous sulfide
agar with formalin to isolate pure cultures may be successful.1,4,15
d. Other iron bacteria: An acid-tolerant (pH 3.5 to 5.0) filamentous iron-oxidizing
Metallogenium has been isolated with a medium16 containing (NH4)2SO4, 0.1%; CaCO3,
0.01%; MgSO4, 0.02%; K2HPO4, 0.001%; potassium acid phthalate, 0.4%; and 250 mg/L
ferrous iron from an acidified FeSO4⋅7H2O, solution. Add 0.4% formalin to 100 mL of the
isolating medium in a 250-mL erlenmeyer flask.
For heterotrophic iron-precipitating bacteria17 use a ferric ammonium citrate medium
consisting of: (NH4)2SO4, 0.5 g/L; NaNO3, 0.5 g/L; K2HPO4, 0.5 g/L; MgSO2⋅7H2O, 0.5 g/L;
and ferric ammonium citrate, 10.0 g/L. Adjust pH to 6.6 to 6.8 and sterilize. To make the
medium solid add 15 g agar/L.
Alternatively grow iron bacteria by combining 20 mL liquid broth medium, 10 mL raw water
or inoculum, and 3 g iron oxide. Incubate 48 to 72 h at 25°C on a wrist-action shaker to produce
good, visible growth.
3. Sulfur Bacteria
a. Sulfate-reducing bacteria:
1) To enumerate sulfate-reducing bacteria such as Desulfovibrio, use a sulfate-reducing
medium.18 Inoculate tubes and fill completely with sterile medium to create anaerobic
conditions. For comparative purposes, incubate one or two uninoculated controls with each set of
inoculated tubes. To sample volumes greater than 10 mL, pass sample through a 0.45-µm
membrane filter and transfer filter to screw-cap test tube with medium. If sulfate-reducing
bacteria are present, tubes will show blackening within 4 to 21 d of incubation at 20 to 30°C.
2) An agar medium suitable for growth and enumeration of sulfate-reducing bacteria also is
available.19 The medium consists of trypticase soy agar, 4.0%, fortified with additional agar,
0.5%, to which is added 60% sodium lactate (0.4% v/v), hydrated magnesium sulfate, 0.2%, and
ferrous ammonium sulfate, 0.2%. Adjust pH to 7.2 to 7.4 and sterilize. Medium should be clear
and free from precipitate. Inoculate all plates within 1 or at most 4 h after agar hardens to
prevent saturation with oxygen. To prevent moisture condensation on petri dish covers, replace
covers with sterile absorbent tops until 10 to 15 min after agar hardens. Place uninverted plates
in desiccator or Brewer jars and replace atmosphere with tank hydrogen or nitrogen by
successive evacuation and gas replacement. Alternatively, use a disposable anaerobic generating
system.*#(51) Incubate at room temperature (21 to 24°C) or at 28 to 30°C, the optimum
temperature for these organisms. Growth and blackening around the colonies is typical of
sulfate-reducing bacteria and may occur at any time between 2 and 21 d, although the usual time
is 2 to 7 d.
3) Media suitable for enumeration or isolation of various species of sulfate-reducing bacteria
are available.20-22
b. Photosynthetic purple and green sulfur bacteria: Because these organisms are so
specialized and rarely cause problems in water and wastewater treatment processes, methods for
their isolation and enumeration are not included here. In certain instances they can be beneficial
because of their ability to oxidize hydrogen sulfide and thus reduce odor. An excellent review is
available.23 Also, media formulations and methods for cultivating specific members of this
group of bacteria are available.23
c. Thiobacillus spp.: The growth and physiology of different species of the single-celled
sulfur-oxidizing bacteria of the genus Thiobacillus have been evaluated carefully.24,25 Media26
suitable for enumeration of Thiobacillus thioparus and Thiobacillus thiooxidans by an MPN
technique are listed in Section 9240D.1. Inoculate medium and incubate for 4 to 5 d at 25 to
30°C. Growth of thiobacilli produces elemental sulfur, which sinks to the bottom with a
coincident decrease in pH and turbidity of the medium. Chemical tests for formation of sulfate
are necessary to confirm presence of Thiobacillus.
d. Filamentous sulfur-oxidizing bacteria:
1) Beggiatoa—Beggiatoa exist in most aquatic habitats where sulfide and oxygen are both
present,27 including fresh and marine water, sediments, and wastewater systems. Because
Beggiatoa is a multicellular bacterium there is no accurate method of determining the number of
viable cells in a sample. To determine the population of Beggiatoa make a direct microscopic
count of the number of filaments.
Marine beggiatoas may be quite large, up to 100 µm or more in diameter.28 They have not
been grown in the laboratory. However, small marine species (up to 5 µm diam) have been
isolated.29 The media for isolating freshwater and marine beggiatoas differ slightly.
a) Enrichment—Inoculate a tube of extracted hay, ¶ 1k1) above, with enough water and a
little mud from the sample site to fill the tube to a depth of at least 8 cm. Incubate for at least 1
week and examine for the presence of ‘‘tufts’’ or ‘‘puff-balls’’ consisting of tangled filaments of
Beggiatoa. Examine by phase-contrast microscopy for the presence of individual filaments.30 If
no ‘‘puff-balls’’ are found, continue incubation for another week and repeat the examination.
Continue examining the enrichment for up to 4 weeks before discarding it.
b) Isolation—With sterilized fine-tipped forceps and using a dissecting microscope, transfer
tufts of Beggiatoa from the enrichments to a small petri dish containing sterile basal medium,30 ¶
1k2) above. Shake the tufts with the forceps to remove adherent bacteria and transfer tufts to a
new petri dish with sterile basal medium. Continue until tufts have been washed at least five
times.
Transfer tufts to a ‘‘drying plate’’ containing basal medium and 1.6% agar for about 1 min to
remove excess fluid, then transfer the tufts to the center of separate plates of either MY or MP
medium,30,31 ¶s 1k3) and 4) above. Incubate plates at room temperature or below and examine
with a dissecting microscope every 5 to 10 h for the presence of gliding filaments of Beggiatoa.
Select filaments that have glided well away from the other filaments and appear to be
uncontaminated and transfer them to separate plates of the same medium with a sterile, flattened
wire, inoculating needle, or toothpick. Take a little agar with the filament to avoid drying the
filament during transfer. Examine the first transfer plates every 5 to 10 h as before and transfer
pure filaments to fresh media.
2) Thiothrix—Obligately myxotrophic Thiothrix have been isolated and characterized from
sulfur springs and other bodies of flowing water that contain sulfide.31,32 Heterotrophic strains
have been isolated and characterized from activated sludge wastewater treatment plants.33-35
While the techniques for the isolation of the myxotrophic and heterotrophic strains are similar,
the isolation media differ.
a) Isolation of myxotrophic Thiothrix—Collect tufts of the bacterium from rocks, water
pipes, or other substrates. Using a dissecting microscope, pick up filaments with fine-tipped
forceps, shake to remove contaminating bacteria, and transfer to sterile basal medium, ¶ 1k2)
above. Repeat at least five times to try to obtain filaments with little or no contamination.
With a sterile Pasteur pipet transfer separate drops to the edge of either MP or MY agar [¶s
1k3) and 4), above] petri dishes and tip the dishes so that the drops run from one side of the dish
to the other. Draw off excess moisture with the pipet and incubate plates at room temperature or
below for about 48 h. Examine plates under a dissecting microscope for the appearance of
typical filamentous colonies. With sterile toothpicks, pick colonies that are widely separated and
transfer individually to fresh plates of the same medium. Streak transferred material with a wire
loop. After about 48 h incubation, examine plates and restreak colonies that appear to be pure.
b) Isolation of heterotrophic Thiothrix—There are two different procedures to use, depending
on the concentration and size of the Thiothrix filaments in the sample.34
In one procedure, wash individual filaments or rosettes of large strains of Thiothrix several
times in MSV broth, ¶ 1l 1) above, by transfer with a Pasteur pipet while observing with a
dissecting microscope. After several washings transfer filaments to a small amount of MSV (1 to
3 mL) and plate on one or more of the following solid media, ¶s 1l 3)–9) above.
If the filaments are small or scarce concentrate the sample by centrifuging. Then dilute 1:5 in
MSV, sonicate at 30 W for 10 s, and wash three times by centrifugation at 1900 × g for 2 to 5
min. The supernatant contains free filaments that are used to inoculate one or more of the solid
media, ¶s 1l 3)–9) above.
4. References
1. GHIORSE, W.C. 1984. Biology of iron- and manganese-depositing bacteria. Annu. Rev.
Microbiol. 38:515.
2. HALLBRECK, E.L. & K. PEDERSON. 1986. The biology of Gallionella. In D.R. Cullimore,
ed. Proc. International Symposium on Biofouled Aquifers: Prevention and Restoration.
3. STARR, M.P., H. STOLP, H.G. TRUPER, A. BALOWS & H.C. SCHLEGE. 1981. The
Prokaryotes. A Handbook on Habitats, Isolation and Identification of Bacteria, Volume
1. Springer-Verlag, New York, N.Y.
4. STALEY, J.T., M.P. BRYANT, N. PFENNIG & J.G. HOLT, eds. 1989. Bergey’s Manual of
5. ARMBRUSTER, E.H. 1969. Improved technique for isolation and identification of
Sphaerotilus. Appl. Microbiol. 17:320.
6. FARQUHAR, G.J. & W.C. BOYLE. 1971. Identification of filamentous microorganisms in
activated sludge. J. Water Pollut. Control Fed. 43:604.
7. VANVEEN, W.L. 1973. Bacteriology of activated sludge, in particular the filamentous
bacteria. Antonie van Leeuwenhoek (Holland) 39: 189.
8. DONDERO, N.C., R.A. PHILIPS & H. HEUKELEKIAN. 1961. Isolation and preservation of
cultures of Sphaerotilus. Appl. Microbiol. 9:219.
9. MULDER, E.G. & W.L. VANVEEN. 1963. Investigations on the Sphaerotilus-Leptothrix
group. Antonie van Leeuwenhoek (Holland) 29:121.
10. UNZ, R.F. & D.G. LUNDGREN. 1961. A comparative nutritional study of three
chemoautotrophic bacteria: Ferrobacillus ferrooxidans, Thiobacillus ferrooxidans, and
Thiobacillus thiooxidans. Soil Sci. 92: 302.
11. MCGORAN, C.J.M., D.W. DUNCAN & C.C. WALDEN. 1969. Growth of Thiobacillus
ferrooxidans on various substrates. Can. J. Microbiol. 15:135.
12. SILVERMAN, M.P. & D.C. LUNDGREN. 1959. Studies on the chemoautotrophic iron
bacterium Ferrobacillus ferrooxidans. J. Bacteriol. 77:642.
13. KUCERA, S. & R.S. WOLFE. 1957. A selective enrichment method for Gallionella
ferruginea. J. Bacteriol. 74:344.
14. WOLFE, R.S. 1958. Cultivation, morphology, and classification of the iron bacteria. J.
15. NUNLEY, J.W. & N.R. KRIEG. 1968. Isolation of Gallionella ferruginea by use of
formalin. Can. J. Microbiol. 14:385.
16. WALSH, F. & R. MITCHELL. 1972. A pH dependent succession of iron bacteria. Environ.
Sci. Technol. 6:809.
17. CLARK, F.M., R.M. SCOTT & E. BONE. 1967. Heterotrophic, iron-precipitating bacteria. J.
18. LEWIS, R.F. 1965. Control of sulfate-reducing bacteria. J. Amer. Water Works Assoc.
57:1011.
19. IVERSON, W.P. 1966. Growth of Desulfovibrio on the surface of agar media. Appl.
Microbiol. 14:529.
20. LECHEVALIER, H.A. & D. PRAMER. 1970. The Microbes, 1st ed. J.B. Lippincott Co.,
Philadelphia, Pa.
21. MARA, D.D. & D.J.A. WILLIAMS. 1970. The evaluation of media used to enumerate
sulphate reducing bacteria. J. Appl. Bacteriol. 33:543.
22. KREIG, N.R. & J.G. HOLT. 1986. Bergey’s Manual of Systematic Bacteriology, Volume
1. Williams & Wilkins. Baltimore, Md.
23. PFENNIG, N. 1967. Photosynthetic bacteria. Annu. Rev. Microbiol. 21: 285.
24. HUTCHINSON, M., K.I. JOHNSTONE & D. WHITE. 1965. The taxonomy of certain
thiobacilli. J. Gen. Microbiol. 41:357.
25. HUTCHINSON, M., K.I. JOHNSTONE & D. WHITE. 1966. Taxonomy of the acidophilic
thiobacilli. J. Gen. Microbiol. 44:373.
26. STARKEY, R.L. 1937. Formation of sulfide by some sulfur bacteria. J. Bacteriol. 33:545.
27. LACKEY, J.B., W.W. LACKEY & G.B. MORGAN. 1965. Taxonomy and ecology of the
sulfur bacteria. Eng. Prog., Univ. Fla. Bull. Ser. 119, 19:3.
28. NELSON, D.C., C.O. WIRSEN & H.W. JANNASCH. 1989. Characterization of large,
autotrophic Beggiatoa spp. abundant at hydrothermal vents of the Guaymas Basin.
29. NELSON, D.C., J.B. WATERBURY & H.W. JANNASCH. 1982. Nitrogen fixation and nitrate
utilization by marine and freshwater Beggiatoa. Arch. Microbiol. 133:172.
30. STROHL, W.R. & J.M. LARKIN. 1978. Enumeration, isolation, and characterization of
31.
32.
33.
34.
35.
Beggiatoa from freshwater sediments. Appl. Environ. Microbiol. 36:755.
LARKIN, J.M. 1980. Isolation of Thiothrix in pure culture and observation of a
filamentous epiphyte on Thiothrix. Curr. Microbiol. 4: 155.
LARKIN, J.M. & D.L. SHINABARGER. 1983. Characterization of Thiothrix nivea. Int. J.
System. Bacteriol. 33:841.
EIKELBOOM, D.H. 1975. Filamentous organisms observed in activated sludge. Water
Res. 9:365.
WILLIAMS, T.M. & R.F. UNZ. 1985. Isolation and characterization of filamentous bacteria
present in bulking activated sludge. Appl. Microbiol. Technol. 22:273.
WILLIAMS, T.M. & R.F. UNZ. 1985. Filamentous sulfur bacteria of activated sludge:
characterization of Thiothrix, Beggiatoa, and Eikelboom type 021N strains. Appl.
9250
DETECTION OF ACTINOMYCETES*#(52)
9250 A.
Introduction
Earthy-musty odors affect the quality and public acceptance of municipal water supplies in
many parts of the world. They are among the naturally occurring odors that plant operators find
most difficult to remove by conventional treatment. As early as 1929, it was assumed that these
odors could be attributed to volatile metabolites formed during normal actinomycete
development.1 Two such compounds, geosmin and 2-methylisoborneol, have been isolated2–8
and identified as the agents responsible for earthy-musty odor problems in surface water.8–10
Both, however, are produced also by some filamentous blue-green algae.11–15 Geosmin and
2-methylisoborneol have threshold odor concentrations well below the microgram-per-liter level.
Thus, traces of these products are sufficient to impart a disagreeable odor to water or a muddy
flavor to fish. In areas periodically plagued by this problem, it is prudent to enumerate
actinomycetes. Identification of their relative abundance in a drinking water source can provide
yet another means to assess water quality. The methods described are well-established
techniques that have been used with success in the isolation and enumeration of actinomycetes
related to public water supplies.16,17 Actinomycetes also have been recognized as a cause of
disruptions in wastewater treatment. Massive growths are capable of producing thick foam in the
activated sludge process.18,19
Of the general properties of actinomycetes, the most striking is their fungal-type
morphology. Although actinomycetes were looked upon initially as fungi, later research revealed
that they were filamentous, branching bacteria.20 The actinomycetes are represented most
commonly by saprophytic forms that have an extensive impact on the environment by
decomposing and transforming a wide variety of complex organic residues. Widely distributed in
nature, actinomycetes constitute a considerable proportion of the population of soil and lake and
river muds. Most actinomycetes from which geosmin and 2-methylisoborneol have been
identified are members of the genus Streptomyces, which is considered the most likely to be
significant in water supply problems.
2. Samples
a. Collection: Collect samples as directed in Section 9060A.
b. Storage: Analyze samples as promptly after collection as possible. Store water samples
below 10°C if they cannot be processed promptly.
3. References
1. ADAMS, B.A. 1929. Cladothrix dichotoma and allied organisms as a cause of an
‘‘indeterminate’’ taste in chlorinated water. Water & Water Eng. 31:327.
2. GERBER, N.N. & H.A. LECHEVALIER. 1965. Geosmin, an earthy-smelling substance
isolated from actinomycetes. Appl. Microbiol. 13:935.
3. GERBER. N.N. 1968. Geosmin, from microorganisms, is
trans-1,10-dimethyl-trans-9-decalol. Tetrahedron Lett. 25:2971.
4. MARSHALL, J.A. & A.R. HOCHSTETLER. 1968. The synthesis of (±)-geosmin and the
other 1,10-dimethyl-9-decalol isomers. J. Org. Chem. 33:2593.
5. ROSEN, A.A., R.S. SAFFERMAN, C.I. MASHNI & A.H. ROMANO. 1968. Identity of odorous
substances produced by Streptomyces griseoluteus. Appl. Microbiol. 16:178.
6. MEDSKER, L.L., D. JENKINS & J.F. THOMAS. 1969. Odorous compounds in natural
waters: 2-exo-hydroxy-2-methylbornane, the major odorous compound produced by
several actinomycetes. Environ. Sci. Technol. 3:476.
7. GERBER, N.N. 1969. A volatile metabolite of actinomycetes, 2-methylisoborneol. J.
Antibiot. 22:508.
8. ROSEN, A.A., C.I. MASHNI & R.S. SAFFERMAN. 1970. Recent developments in the
chemistry of odour in water: The cause of earthy/ musty odour. Water Treat. Exam.
19:106.
9. PIET, G.J., B.C.J. ZOETEMAN & A.J.A. KRAAYEVELD. 1972. Earthy-smelling substances in
surface waters of the Netherlands. Water Treat. Exam. 21:281.
10. YURKOWSKI, M. & J.A.L. TABACHEK. 1974. Identification, analysis, and removal of
geosmin from muddy-flavored trout. J. Fish. Res. Board Can. 31:1851.
11. SAFFERMAN, R.S., A.A. ROSEN, C.I. MASHNI & M.E. MORRIS. 1967. Earthy-smelling
substances from a blue-green alga. Environ. Sci. Technol. 1:429.
12. MEDSKER, L.L., D. JENKINS & J.F. THOMAS. 1968. Odorous compounds in natural
waters. An earthy-smelling compound associated with blue-green algae and
actinomycetes. Environ. Sci. Technol. 2: 461.
13. KIKUCHI, T., T. MIMURA, K. HARIMAYA, H. YANO, M. ARIMOTO, Y. MASADA & T. INOUE.
1973. Odorous metabolites of blue-green alga Schizothrix muelleri Nageli collected in
the southern basin of Lake Biwa. Identification of geosmin. Chem. Pharm. Bull.
21:2342.
14. TABACHEK, J.L. & M. YURKOWSKI. 1976. Isolation and identification of blue-green
algae producing muddy odor metabolites, geosmin and 2-methylisoborneol, in saline
lakes in Manitoba. J. Fish. Res. Board Can. 33:25.
15. IZAGUIRRE, G., C.J. HWANG, S.W. KRASNER & M.J. MCGUIRE. 1982. Geosmin and
2-methylisoborneol from cyanobacteria in three water supply systems. Appl. Environ.
Microbiol. 43:708.
16. SAFFERMAN, R.S. & M.E. MORRIS. 1962. A method for the isolation and enumeration of
actinomycetes related to water supplies. Robert A. Taft Sanitary Engineering Center
Tech. Rep. W62-10, U.S. Public Health Serv., Cincinnati, Ohio.
17. KUSTER, E. & S.T. WILLIAMS. 1964. Selection of media for isolation of Streptomyces.
Nature 202:928.
18. LECHEVALIER, H.A. 1975. Actinomycetes of sewage-treatment plants. Environ.
Protection Technol. Ser., EPA-600/2-75-031, U.S. Environmental Protection Agency,
Cincinnati, Ohio.
19. LECHEVALIER, M.P. & H.A. LECHEVALIER. 1974. Nocardia amarae, sp. nov., an
actinomycete common in foaming activated sludge. Int. J. Syst. Bacteriol. 24:278.
20. LECHEVALIER, H.A. & M.P. LECHEVALIER. 1967. Biology of actinomycetes. Annu. Rev.
Microbiol. 21:71.
9250 B.
Actinomycete Plate Count
A plating method using a double-layer agar technique has been adapted for determining
actinomycete density. Because only the thin top layer of the medium is inoculated with sample,
surface colonies predominate and identification and counting of colonies is facilitated.
2. Preparation and Dilution
Prepare and dilute samples as directed in Section 9215 or Section 9610. Dilutions up to
1:1000 (10–3) usually are suitable for raw water, while treated waters may be examined directly.
For soil samples, use dilutions from 1:1000 (10–3) to 1:1 000 000 (10–6).
3. Medium
Starch-casein agar:
Soluble starch
Casein
Potassium nitrate, KNO3
10.0 g
0.3 g
2.0 g
2.0 g
2.0 g
Magnesium sulfate, hydrate, MgSO4⋅7H2O
g
Calcium carbonate, CaCO3
Ferrous sulfate, hydrate, FeSO4⋅7H2O
Agar
Reagent-grade water
0.05
g
0.02
g
0.01
15.0 g
1 L
No pH adjustment is required. Medium is used to prepare double-layer plates. Store medium
for bottom layer in bulk or in tubes in 15-mL amounts. Store medium for surface layer in tubes
in 17.0-mL amounts.
4. Procedure
a. Plating: Prepare three plates for each dilution to be examined. Aseptically transfer 15 mL
of sterile starch-casein agar to a petri dish and let agar solidify, thus forming the bottom layer.
To a test tube containing 17.0 mL liquefied starch-casein agar at 45 to 48°C, add 2 mL of
appropriately diluted sample and 1 mL of the antifungal antibiotic, cycloheximide,*#(53)
prepared in reagent-grade water (1 mg/mL) and sterilized by autoclaving for 15 min at 121°C.
Pipet 5 mL of inoculated agar over the hardened bottom layer with gentle swirling to obtain even
distribution of the surface layer.
b. Incubation: Invert and incubate at 28°C until no new colonies appear. Usually this
requires 6 to 7 d.
c. Counting: Plates suitable for counting contain 30 to 300 colonies. Identify actinomycetes
by gross colony appearance. If necessary, verify by microscopic examination at a magnification
of 50 to 100×, as shown in Figure 9250:1. Actinomycete colonies, because of filamentous
growth, typically have a fuzzy colonial border. Table 9250:I lists the distinguishing
characteristics commonly used to differentiate actinomycete from other bacterial colonies.
Cycloheximide generally suppresses fungal growth; however, fungal colonies, if present, can be
recognized by their wooly appearance. Microscopically, fungi reveal a considerably larger cell
diameter than actinomycetes.
5. Calculation
Report actinomycetes per milliliter of water or gram (dry weight) of soil. If three plates are
used per sample, the average number of colonies on all plates (total number of colonies/3), times
2, times the reciprocal of the dilution (10/1, 100/1, 1000/1, etc.) equals the actinomycete colony
count per milliliter of original sample. For solid or semisolid samples, correct for water content
and report actinomycete colonies per gram, dry weight, of sample.
9260
DETECTION OF PATHOGENIC BACTERIA*#(54)
9260 A.
Introduction
One purpose of drinking water and wastewater treatment is to reduce the numbers of viable
organisms to acceptable levels, and to remove or inactivate all pathogens capable of causing
human disease. Despite the remarkable success of water treatment and sanitation programs in
improving public health, sporadic cases and point-source outbreaks of waterborne diseases
continue to occur. Water and wastewater may contain a wide variety of bacteria that are
opportunistic or overt pathogens of animals and humans. Waterborne pathogens enter human
hosts through intact or compromised skin, inhalation, ingestion, aspiration, and direct contact
with the mucous membranes of the eye, ear, nose, mouth, and genitals. This section provides an
introduction to the etiologic agents responsible for diseases transmitted by drinking and
recreational waters in the U.S.
Over 80 genera of bacteria that are nonpathogenic for humans have their natural habitat in
water. In addition, some opportunistically pathogenic bacteria (Pseudomonas, Serratia,
Acinetobacter, Chromobacterium, Achromobacter, Aeromonas, etc.) occur naturally in water.
Other opportunists (Bacillus, Enterobacter, Klebsiella, Actinomyces, Streptomyces, etc.) are
sometimes washed into water from their natural habitat in soil or on vegetative matter.
Opportunistic pathogens also may be seeded from regrowth and biofilms in water treatment
plants and distribution systems.
Water contamination and disease transmission may result from conditions generated at
overloaded and/or malfunctioning sanitary waste disposal and potable water treatment systems.
In addition, common outdoor recreational activities such as swimming (including pools and hot
tubs), boating, camping, and hiking, all place humans at risk of waterborne diseases from
ingestion or direct contact with contaminated water.1 Outbreaks of gastroenteritis,
pharyngoconjunctivitis, folliculitis, otitis, and pneumonia are associated with these recreational
activities. Overcrowded parks and recreational areas contribute to the contamination of surface
and groundwater.
National statistics on outbreaks of waterborne diseases have been compiled in the U.S. since
1920.2,3 Since 1971, the Centers for Disease Control and Prevention, the U.S. Environmental
Protection Agency, and the Council of State and Territorial Epidemiologists have maintained a
collaborative surveillance program on waterborne disease outbreaks of drinking water and
recreational water origin.4 A summary of waterborne diseases in the U.S. has been published.5
Summary data from outbreaks reported through the national waterborne disease surveillance
system for drinking water and recreation from 1985 to 1994 are shown in Table 9260:I.
Laboratory diagnosis of infectious disease depends on isolation of the etiologic agent or
demonstration of antibody response in the patient. Environmental microbiological examinations
are conducted for compliance monitoring of the environment, to trouble-shoot problems in
treatment plants and distribution systems, and in support of epidemiological investigations of
disease outbreaks. Ideally, the public health microbiologist can contribute expertise in both
clinical and environmental microbiology, thereby facilitating epidemiological investigations.
When testing for pathogens in environmental samples, it usually is advisable to include
analyses for indicator organisms. Besides coliform indicators (total coliform, fecal coliform, and
E. coli), fecal streptococci, enterococci, Clostridium perfringens, and Aeromonas have been
proposed as indicators of water quality. No single indicator provides assurance that water is
pathogen-free. The choice of monitoring indicator(s) presupposes an understanding of the
parameters to be measured and the relationship of the indicator(s) to the pathogen(s). Some
bacterial pathogens, such as Pseudomonas, Aeromonas, Plesiomonas, Yersinia, Vibrio,
Legionella, and Mycobacterium, may not correlate with coliform indicators. Traditional bacterial
indicators also may not correlate with viruses or parasites in pristine waters or groundwaters, and
they may be of limited utility in estuarine and marine waters. Nevertheless, tests for total and
fecal bacteria and E. coli are useful, because it is rare to isolate bacterial enteric pathogens in the
absence of fecal contamination.
Other more general indicators also may be of value for assessing the potential for pathogen
contamination and interpreting culture results. Heterotrophic plate count provides information
about the total numbers of aerobic organotrophic bacteria and an indication of the total organic
composition of the aquatic environment. Physicochemical factors, such as turbidity, pH, salinity,
temperature, assimilable organic carbon, dissolved oxygen, biochemical oxygen demand, and
ammonia may provide useful information about contamination or the potential of water to
support bacterial growth. For treated waters, chlorine residual should be measured at the sample
collection point.
This section contains methods for Salmonella, Shigella, pathogenic E. coli, Campylobacter,
Vibrio cholerae, Leptospira, Legionella, Yersinia entercolitica, Aeromonas, and Mycobacterium.
Methods for isolation and enumeration of P. aeruginosa are found in Section 9213E and Section
9213F. Methods for other pathogens are found elsewhere.6
The methods outlined below may be used to analyze samples associated with disease
outbreaks, or in other studies on the occurrence of pathogens in water and wastewater. Methods
for recovery of bacterial pathogens from water and wastewater have not changed significantly in
the past 30 years. The methods presented below are not standardized, and the procedures may
need modification to fit a particular set of circumstances. No single procedure is available for
reliable detection of any pathogen or group of pathogens. Because the presence of pathogens is
intermittent and the survival times in the environment are variable, routine examination of water
and wastewater for pathogenic bacteria is not recommended. Even in outbreak situations, the
recovery of pathogens from water and wastewater may be limited by lack of facilities, untrained
personnel, inadequate methods, and high costs.
2. References
1. PITLIK, S., S.A. BERGER & D. HUMINER. 1987. Nonenteric infections acquired through
contact with water. Rev. Infect. Dis. 9:54.
2. CRAUN, G.F., ed. 1986. Waterborne Diseases in the United States. CRC Press, Inc.,
Boca Raton, Fla.
3. LIPPY, E.C. & S.C. WALTRIP. 1984. Waterborne disease outbreaks— 1946–1980: A
thirty-five year perspective. J. Amer. Water Works Assoc. 76:60.
4. KRAMER, M.H., B.L. HERWALDT, G.F. CRAUN, R.L. CALDERON & D.D. JURANEK. 1996.
Waterborne diseases: 1993 and 1994. J. Amer. Water Works Assoc. 88:66.
5. KRAMER, M.H., B.L. HERWALDT, G.F. CRAUN, R.L. CALDERON & D.D. JURANEK. 1996.
Surveillance for waterborne-disease outbreaks— United States, 1993–1994. Morbid.
Mortal. Week. Rep. 45(SS-1):1.
6. MURRAY, P.R., E.J. BARON, M.A. PFALLER, F.C. TENOVER & R.H. YOLDEN, eds. 1995.
Manual of Clinical Microbiology, 6th ed. American Soc. Microbiology Press,
Washington, D.C.
9260 B.
General Qualitative Isolation and Identification Procedures for
Salmonella
Rather than a specific protocol for Salmonella detection in water, a brief summary of
methods suitable for recovery of these organisms is given. Methods currently available have
been used in numerous field investigations to demonstrate Salmonella in both fresh and marine
water environments. The occurrence of Salmonella in water is highly variable; there are
limitations and variations in both the sensitivity and selectivity of accepted Salmonella isolation
procedures for the detection of the more than 2300 Salmonella serotypes currently recognized.
Thus, a negative result by any of these methods does not imply the absence of salmonellae, nor
does it imply the absence of other pathogens.
1. Concentration Techniques
Salmonella are ubiquitous in the environment and can be detected at low concentrations in
most surface waters. These organisms are usually present in small numbers compared to
coliforms; therefore, it is necessary to examine a relatively large sample to isolate the
organisms.1
a. Swab technique: Prepare swabs from cheesecloth 23 cm wide, folded five times at 36-cm
lengths, and cut lengthwise to within 10 cm from the head into strips approximately 4.5 cm wide.
Securely wrap the uncut or folded end of each swab with 16-gauge wire for use in suspending
the swab in water. Place the swabs in kraft-type bags and sterilize at 121°C for 15 min. Place
swab just below the surface of the sampling location for 1 to 3 d.2,3 (Longer swab exposure will
not increase entrapment of pathogens.) Gauze pads of similar thickness may be substituted.
During sampling, particulate matter and microorganisms are concentrated from the water passing
through or over the swab. After exposure, retrieve the swab, place it in a sterile plastic bag, ice,
and send to the laboratory. Maximum storage-transit time allowable is 6 h. Do not transport
swabs in enrichment media; ambient transport temperature may cause sufficient proliferation of
competitive organisms to mask salmonellae. In the laboratory, place pad or portions of it in
enrichment media. When flasks of enrichment medium containing iced swabs are to be incubated
at 40 to 41°C, place flasks in a 44.5°C water bath for 5 min before incubation in an air incubator.
b. Diatomaceous earth technique: Place an absorbent pad (not a membrane filter) on a
membrane filter funnel receptacle, assemble funnel, and add 2.5 g sterile diatomaceous
earth*#(55) to pack the funnel neck loosely. Apply vacuum and filter 2 L of sample. After
filtration, disassemble funnel, divide resulting ‘‘plug’’ of diatomaceous earth and absorbent pad
in half aseptically with a knife-edged, sterile spatula, and add to suitable enrichment media.
Alternatively, place entire plug in enrichment medium.
c. Large-volume sampler: Use a filter composed of borosilicate glass microfibers bonded
with epoxy resin to examine several liters or more of sample, provided that sample turbidity does
not limit filtration.4 The filter apparatus consists of a 2.5- × 6.4-cm cartridge filter and a filter
holder.†#(56) Sterilize by autoclaving at 121°C for 15 min. Place sterile filter apparatus
(connected in series with tubing to a 20-L water bottle reservoir and vacuum pump) in the 20-L
sample container appropriately calibrated to measure volume of sample filtered. Apply vacuum
and filter an appropriate volume. When filtration is complete, remove filter and place in a
selective enrichment medium.
d. Membrane filter technique: To examine low-turbidity water, filter several liters through a
sterile 142-mm-diam membrane of 0.45-µm pore size.5 For turbid waters, precoat the filter:
make 1 L of sterile diatomaceous earth suspension (5 g/L reagent-grade water) and filter about
500 mL. Without interrupting filtration, quickly add sample (1 L or more) to remaining
suspension and filter. After filtration, place membrane in a sterile blender jar containing 100 mL
sterile 0.1% (w/v) peptone water and homogenize at high speed for 1 min. Add entire
homogenate to 100 mL double-strength selective enrichment medium. Alternatively, use
multiple 47-mm-diam membrane filters to filter the sample. Immerse each membrane aseptically
in 50 mL single-strength selective enrichment medium and incubate.
Qualitative detection of Salmonella in suspect potable water also may be achieved
successfully by further analysis of selected M-Endo MF cultures (from 100 mL sample volume)
that contain significant background growth and total coliforms.6 After completing routine
coliform count, place entire filter with mixed growth into 10 mL tetrathionate broth (containing
1:50 000 brilliant green dye) for Salmonella enrichment before differential colony isolation on
brilliant green agar. This unique approach requires no special large sample collections and can
be an extension of the routine total coliform analysis.
2. Enrichment
Selectively enrich the concentrated sample in a growth medium that suppresses growth of
coliform bacteria. Sample enrichment is essential, because the pathogens usually are present in
low numbers and solid selective media for colony isolation are somewhat toxic, even to
pathogens. No single enrichment medium can be recommended that allows optimum growth of
all Salmonella serotypes. Use two or more selective enrichment media in parallel for optimum
detection. Elevated incubation temperatures including 40°, 41.5°, and 43°C and the addition of
brilliant green dye to media help suppress background growth and may improve detection of
Salmonella, but these modifications also suppress growth of some serotypes, including
Salmonella typhi.
a. Selenite cystine broth inhibits gram-positive and nonpathogenic enterobacteria while
allowing for recovery of most species of Salmonella, including Salmonella typhi. Optimum
incubation time for maximum recovery of Salmonella is 48 h at 35 to 37°C. Repeat streaking
from tubes with turbidity several times during first day, and daily up to 5 d to increase potential
recovery of all serotypes that may be present. Transfer 1 mL selenite broth culture to a fresh tube
of same medium for continued incubation to enrich further Salmonella growth and enhance
recovery of streak plates.
b. Selenite-F broth allows for optimum recovery of most Salmonella species, including
Salmonella typhi, after 24 h at 35 to 37°C. This increased recovery of Salmonella is
accompanied by a slight decrease in selectivity when compared to selenite cystine. Most
significantly, E. coli growth is not inhibited. Repeat streaking from tubes with turbidity several
times during first day, and daily up to 5 d to increase potential recovery of all serotypes that may
be present. Transfer 1 mL selenite broth culture to a fresh tube of same medium for continued
incubation to enrich further Salmonella growth and enhance recovery of streak plates.
c. Tetrathionate broth, incubated at 35°C, inhibits coliforms and Gram-positive bacteria,
permitting selective enrichment of most Salmonella species, including S. typhi. It has been
reported that tetrathionate broth is more selective for Salmonella than selenite-based media when
incubated for 48 h at 43°C. While this formulation is highly selective, it is unable to inhibit
Proteus mirabilis, which shows optimum growth. Growth of Proteus and Citrobacter can be
inhibited with addition of brilliant green (see Section 9260B.3a). Incubation at 43°C and
addition of brilliant green also will inhibit some species of Salmonella, including S. typhi.
3. Selective Growth
Further separation of pathogens from the remaining nonpathogenic bacterial population is
facilitated by proper choice of incubation temperature for primary enrichment followed by
secondary differentiation on selective solid media.7 These factors, incubation temperature,
enrichment medium, and isolation medium, are interrelated. No one combination is optimum for
recovery of all Salmonella serotypes. Method comparisons are encouraged to determine the best
combination for a given circumstance.
Solid media commonly used for enteric pathogen detection may be classed into three groups:
(a) differential media with little or no inhibition toward nonpathogenic bacteria, such as EMB
(containing sucrose); (b) selective media containing bile salts or sodium desoxycholate as
inhibitors,8 such as MacConkey’s agar, desoxycholate agar, or xylose lysine desoxycholate
(XLD) agar; and (c) selective media containing brilliant green dye, such as brilliant green agar or
bismuth sulfite agar. Any medium selected must provide optimum suppression of coliforms
while permitting good recovery of the pathogenic group. Great skill at screening for these
pathogens is necessary because of the competing growth of various nonpathogens. Streaking
duplicate plates, one heavily and one lightly, often aids in recognition of enteric pathogens in the
presence of large numbers of interfering organisms.
a. Brilliant green agar: Typical well-isolated Salmonella colonies grown on this medium are
pinkish white with a red background. S. typhi and a few other species of Salmonella grow poorly
because of the brilliant green dye content. Lactose-fermenters not subject to growth suppression
will form greenish colonies or may produce other colorations. Occasionally, slow
lactose-fermenters (Proteus, Citrobacter, and Pseudomonas) will produce colonies resembling
those of a pathogen. Suppress spreading effect of pseudomonads by increasing agar
concentration to 2%. In some instances, Proteus has been observed to ‘‘swarm’’; reduce this
tendency by using agar plates dried to remove surface moisture. If suspect Salmonella colonies
are not observed after 24 h incubation, reincubate for an additional 24 h to permit slow-growing
or partially inhibited organisms to develop visible colonies. If typical colonies are not observed
or if the streak plate is crowded, isolate in pure culture a few colonies for biochemical
characterization. Non-lactose-fermenting colonies in close proximity to lactose-fermenting
colonies may be masked.
b. Bismuth sulfite agar (Wilson and Blair medium9): Luxuriant growth of many Salmonella
species (including S. typhi) can be expected on this medium. Examine bismuth sulfite plates after
24 h incubation for suspect colonies; reincubate for 24 h to detect slow-growing strains. Typical
colonies usually develop a black color, with or without a metallic sheen, and frequently this
blackening extends beyond the colony to give a ‘‘halo’’ effect. A few species of Salmonella
develop a green coloration; therefore, isolate some of these colony types when typical colonies
are absent. As with brilliant green agar, typical colony coloration may be masked by numerous
bordering colonies after 48 h incubation. A black color also is developed by other
H2S-producing colonies, for example, Proteus and certain coliforms.
c. Xylose lysine desoxycholate agar: Compared to brilliant green dye, sodium desoxycholate
is only slightly toxic to fastidious Salmonella. Salmonella and Arizona organisms produce
black-centered red colonies. Coliform bacteria, Proteus, and many Enterobacter produce yellow
colonies. Optimum incubation time is 24 h. If plates are incubated longer, an alkaline reversion
and subsequent blackening occur with H2S-positive nonpathogens (Citrobacter, P. vulgaris, and
P. mirabilis).
d. Xylose lysine brilliant green agar: This medium is especially good for Salmonella from
marine samples. The brilliant green inhibits many Proteus, Enterobacter, and Citrobacter
species.
4. Biochemical Reactions
Many enteric organisms of little or no pathogenicity share certain major biochemical
characteristics with Salmonella. The identification of pathogens by colony characteristics on
selective solid media has limitations inherent in the biological variations of certain organisms
and cannot be relied on for even tentative identification. Suspected colonies grown on selective
solid media must be purified and further characterized by biochemical reactions; final
verification is based on serological identification. Usually a large number of cultures will be
obtained from the screening procedure.
Commercially available differential media kits (see Section 9225) may be used as an
alternative to Phases1, 2, and 3 described below, before serological confirmation. These kits give
95 to 98% agreement with conventional tests, although more significant tests will be necessary
to achieve further differentiation among strains of Enterobacteriaceae.
When such kits are not used, follow a sequential pattern of biochemical testing that will
result in a greater saving of media and time for laboratory personnel.10
Phase 1—Preliminary screening, phenylalanine deaminase activity: Discard phenylalanine
deaminase-positive cultures immediately as indicative of the Proteus group. In this test, spot
isolates on phenylalanine agar and incubate for 24h at either 35 or 37°C. Phenylalanine
deaminase activity is indicated by a green zone that develops around the colony after flooding of
the plate with a 0.5M FeCl3 solution. Subject phenylalanine deaminase-negative cultures to the
biochemical tests of Phase2.
Phase 2—Biochemical tests: The tests used are:
Medium
Purpose of Test
TSI
Fermentation pattern: H2S production
LIA
Lysine decarboxylase activity, H2S
production
Urease production
Urea broth
Conformance to the typical biochemical patterns of the Salmonella determines whether to
process cultures further (Phase3). Aberrant cultures may be encountered that do not conform to
all the classical reactions attributed to each pathogenic group. In all cases, therefore, review
reactions as a whole and do not discard cultures on the basis of a small number of apparent
anomalies.
Phase 3—Fermentation reactions: Test fermentation reactions in dextrose, mannitol,
maltose, dulcitol, xylose, rhamnose, and inositol broths to characterize further the biochemical
capabilities of the isolates. This additional sorting reduces the possible number of positive
cultures to be processed for serological confirmation. If the testing laboratory is equipped for
serological confirmation (see 9260B.5.), this series of biochemical tests may be eliminated.
5. Genus Identification by Serological Techniques
Upon completion of the recommended biochemical tests, inoculate the suspected Salmonella
pure culture onto a brain-heart infusion agar slant and incubate for 18 to 24 h at 35 to 37°C. With
wax pencil (china marker), divide an alcohol-cleaned glass slide into four sections. Prepare a
dense suspension of test organism by suspending growth from an 18- to 24-h agar slant in 0.5
mL 0.85% NaCl solution. Place a drop of Salmonella ‘‘O’’ polyvalent antiserum in the first
section and antiserum plus 0.85% NaCl in the second section. Using a clean inoculating loop,
transfer a loopful of bacterial suspension to the third section containing 0.85% NaCl solution and
to the fourth section containing 0.85% NaCl solution plus antiserum. Gently rock slide back and
forth. If agglutination is not apparent in the fourth section at the end of 1 min, the test is
negative. All other sections should remain clear.
When biochemical reactions are characteristic of S. typhi and the culture reacts with ‘‘O’’
polyvalent antiserum, check other colonies from the same plate for Vi antigen reaction. If there
is no agglutination with Salmonella Vi antiserum, the culture is not S. typhi. Identification of
Salmonella serotypes requires determination of H antigens and phase of the organism as
described by Edwards and Ewing.10 Isolates yielding biochemical reactions consistent for
Salmonella and positive with polyvalent ‘‘O’’ antiserum may be identified as ‘‘Salmonella sp.,
serotype or bioserotype undetermined.’’ If species identification is necessary, send isolates
confirmed as Salmonella by biochemical tests and polyvalent ‘‘O’’ antisera to reference
laboratories for further analysis.
6. References
1. CHERRY, W.B., J.B. HANKS, B.M. THOMASON, A.M. MURLIN, J.W. BIDDLE & J.M. GROOM.
1972. Salmonellae as an index of pollution of surface waters. Appl. Microbiol. 24:334.
2. MOORE, B. 1948. The detection of paratyphoid carriers in towns by means of sewage
examination. Mon. Bull. Mist. Health Pub. Health Lab. Serv. 7:241.
3. MOORE, B., E.L. PERRY & S.T. CHARD. 1952. A survey by the sewage swab method of
latent enteric infection in an urban area. J. Hygiene 50:137.
4. LEVIN, M.A., J.R. FISCHER & V.J. CABELLI. 1974. Quantitative large-volume sampling
technique. Appl. Microbiol. 28:515.
5. PRESNELL, M.W. & W.H. ANDREWS. 1976. Use of the membrane filter and a filter aid for
concentrating and enumerating indicator bacteria and Salmonella from estuarine
6.
7.
8.
9.
10.
waters. Water Res. 10:549.
CANLAS, L. 1975. Personal communication. Guam Environmental Protection Agency,
Agana, Guam.
CHEN, H., A.D.E. FRASER & H. YAMAZAKI. 1993. Evaluation of the toxicity of
Salmonella selective media for shortening the enrichment period. Int. J. Food
Microbiol. 18:151.
LEIFSON, E. 1935. New culture media based on sodium desoxycholate for the isolation
of intestinal pathogens and for enumeration of colon bacilli in milk and water. J.
Pathol. Bacteriol. 40:581.
WILSON, W.J. & E.M. MCV. BLAIR. 1926. Combination of bismuth and sodium sulfite
affording enrichment and selective medium for typhoid and paratyphoid groups of
bacteria. J. Pathol. Bacteriol. 29:310.
EDWARDS, P.R. & W.H. EWING. 1986. Identification of Enterobacteriaeceae, 4th ed.
Elsevier Science Publ. Co., Inc., New York, N.Y.
7. Bibliography
MULLER, G. 1947. Der Nachweis von Keimer der Typhus-Paratyphusgruppe in Wasser. H.H.
Nolke Verlag, Hamburg, Germany.
GREENBERG, A.E., R.W. WICKENDEN & T.W. LEE. 1957. Tracing typhoid carriers by means of
sewage. Sewage Ind. Wastes 29:1237.
MCCOY, J.H. 1964. Salmonella in crude sewage, sewage effluent, and sewage polluted natural
waters. In Int. Conf. Water Pollut. Res., 1st, London, 1962. Vol. 1:205. MacMillan, New
York, N.Y.
BREZENSKI, F.T., R. RUSSOMANNO & P. DEFALCO, JR. 1965. The occurrence of Salmonella and
Shigella in post-chlorinated and nonchlorinated sewage effluents and receiving waters.
Health Lab. Sci. 2:40.
SPINO, D.E. 1966. Elevated temperature technique for the isolation of Salmonella from streams.
GALTON, M.M., G.K. MORRIS & W.T. MARTIN. 1968. Salmonella in foods and feeds. Review of
isolation methods and recommended procedures. Public Health Serv. Bur. Disease
Prevention & Environmental Control, National Center for Disease Control, Atlanta, Ga.
BREZENSKI, F.T. & R. RUSSOMANNO. 1969. The detection and use of Salmonella in studying
polluted tidal estuaries. J. Water Pollut. Control Fed. 41:725.
MORINIGO, M.A., M.A. MUNOZ, E. MARTINEZ-MANZANARES, J.M. SANCHEZ & J.J. BORREGO. 1993.
Laboratory study of several enrichment broths for the detection of Salmonella spp.
particularly in relation to water samples. J. Appl. Bacteriol. 74:330.
U.S. FOOD AND DRUG ADMINISTRATION. 1995. Bacteriological and Analytical Manual, 8th ed.
Assoc. Official Analytical Chemists International, Gaithersburg, Md.
9260 C.
Immunofluorescence Identification Procedure for Salmonella
The direct fluorescent antibody (FA) technique is a rapid and effective means of detecting
salmonellae in fresh- and seawater samples. It may be used as a screening technique to provide
rapid results for large numbers of samples, such as those from recreational or
shellfish-harvesting waters. Positive FA tests are presumptive evidence for the presence of
Salmonella. Because of potential cross-reactivity of antibodies, positive FA results should be
confirmed by other methods. Sample volumes used depend on the degree of contamination.
Where gross pollution is present, use smaller samples. When background information is absent,
analyze a 2-L sample, using the diatomaceous earth concentration technique.
1. Apparatus for Fluorescence Microscopy
Standard fluorescent antibody microscopy equipment may be obtained separately or in a
package containing the essential instrumentation (a-f):
a. Light microscope with microscope stand.
b. Light source, providing energy in the short-wavelength region of the spectrum. A
high-pressure mercury 200-W arc enclosed in a quartz envelope, a 75- to 150-W xenon
high-pressure lamp, or a low-voltage 100-W quartz halogen lamp may satisfy this requirement.
A significant portion of the energy should be emitted in the ultraviolet and blue region of the
spectrum.
c. Power pack to provide constant voltage and wattage output for the selected lamp.
d. Basic filters including heat-absorbing filter (KG-1 or KG-2, or equivalent): red-absorbing
filter (BG-38 or equivalent); exciter filter (BG-12 or equivalent, BG-12 being also a blue filter);
and barrier filter (OG-1 or blue-absorbing filter). New interference excitation filters (KP500 or
equivalent) having very high transmission in the blue portion of the spectrum (490 nm) are
available. Barrier or suppression filters used with these have a sharp cutoff at 500 to 510 nm.
e. Optics: The fluorescence microscope must have high-quality optics. A 100 × objective
with an iris diaphragm to reduce the numerical aperture (N.A.) for dark-field work is essential.
Because the N.A. is similar for all 100 × objectives (1.25 to 1.30), base selection on desire for a
flat-field (plano) lens.
f. Cardioid dark-field condenser for illuminating specimen: A 95 × oil immersion objective
with build-in iris diaphragm is desirable. True dark-field illumination can be achieved only if the
objective N.A. is smaller than the condenser N.A., i.e., of the illuminating cone of light.
(Difference in N.A. between objective and condenser should be at least 0.05.) Reduce N.A. of an
oil immersion objective by using the built-in diaphragm or by putting a funnel stop onto the
objective.
g. FA pre-cleaned micro slides, 7.6- × 2.5-cm, 0.8- to 1.0-mm thickness.
h. Cover glass for FA slides, No. 1 1/2, 0.16- to 0.19-mm thickness.
i. Staining assembly consisting of dish, cover, and slide rack with handle. Five dishes are
required; for Kirkpatrick’s fixative, 95% ethanol, first PBS rinse, second PBS rinse, and reagent
water.
j. Moist chamber used to incubate slides containing smears with added conjugate. A simple
chamber consists of water-saturated toweling with a culture dish bottom (150 by 20 mm) placed
over the wet toweling.
2. Reagents
a. Nondrying immersion oil, Type A (low fluorescence, PCB-free).*#(57)
b. FA Kirkpatrick fixative, consisting of 60 mL absolute ethanol, 30 mL chloroform, and 10
mL formaldehyde.†#(58)
c. Phosphate-buffered saline (PBS): Add 10 g buffer‡#(59) to 1000 mL freshly prepared
distilled water. Stir until the powder dissolves completely. Adjust with 0.2N NaOH to pH 8.0.
d. FA mounting fluid: Use standardized reagent-grade glycerine adjusted to pH 9.0 with
0.2N NaOH and intended for mounting slides to be viewed with the FA microscope.
e. Reagent (laboratory pure) water: Use double-distilled water from an all-glass still or
other high-quality analytical-grade laboratory water.
f. FA Salmonella panvalent conjugate is a fluorescein-conjugated anti-Salmonella
globulin.§#(60) To rehydrate, add 5 mL reagent water to a vial or conjugate. Determine working
dilution (see ¶ 5e). Store unused rehydrated conjugate in a freezer, preferably at −60°C. Avoid
repeated freezing and thawing.
g. Zn-CdS: Ag phosphor particle.i#(61)
3. Concentration Technique
Place an absorbent pad on a membrane filter funnel and add sufficient sterile diatomaceous
earth##(62) to pack funnel neck loosely. Filter 2 L of sample. Rinse funnel with 50 to 100 mL
sterile phosphate-buffered dilution water or 0.1% peptone water. Disassemble funnel and remove
resulting ‘‘plug’’ of diatomaceous earth and the absorbent pad. Repeat with a second 2-L
sample.
4. Enrichment
Immerse one plug and absorbent pad in a flask containing 300 mL selenite cystine broth.
Immerse second plug and absorbent pad in a flask containing 300 mL tetrathionate broth
supplemented with 3 mL 1:1000 aqueous solution of brilliant green dye and 3 mg l-cystine.
Incubate at either 35 or 37°C for 24 h.
5. Fluorescent Antibody Reaction and Analysis
a. Prepare spot plates of brilliant green agar (BGA) and xylose lysine brilliant green (XLBG)
agar by placing 1 drop (about 0.01 mL, delivered with a wire or sterile plastic loop) of the
enrichment medium (selenite cystine or tetrathionate broth) at each of four separate points on the
agar surface.1 Space drops on agar plate so that FA microscope slide will cover two inoculation
points. This is essential because glass slide impression smears of the inoculated points will be
made after incubation of plates.
b. Incubate BGA and XLBG plates at 37 ± 0.5°C for 2.5 to 3 h. After incubation, micro
CFUs will develop. Make impression smears by taking a clean FA microscope glass slide and
placing it over two inoculated points on the medium. Press down lightly, being careful not to
move glass slide horizontally. Do not apply too much pressure, because it will cause movement
of the slide and collection of additional agar. Repeat this process for the other two inoculation
points and for inoculation points on second agar medium. Prepare a total of four FA slides in this
manner.
c. Air-dry smears and fix for 2 min in Kirkpatrick’s fixative. Rinse slides briefly in 95%
ethanol and let air dry. Do not blot.
d. Cover fixed smears with 1 drop of Salmonella panvalent conjugate. Before use, dilute
commercial conjugate and determine appropriate working dilution. Most batches are effective at
a 1:4 dilution but this will vary with the type of fluorescence equipment used, light source,
alignment, magnification, cultures, etc. Determine working dilution (titer) of each lot of
conjugate.
e. To determine conjugate titer use a known 18- to 24-h Salmonella culture grown in veal
infusion broth and make smears on FA glass slide. Dilute conjugate and treat as outlined in c and
d above. For example, if the following results are obtained:
Dilution of Conjugate
Fluorescence
1:2
1:4
1:6
1:8
1:10
4+
4+
4+
2+
1+
use the second highest dilution giving 4+ fluorescence. In the above example use a 1:4 dilution
of conjugate. Diluting conjugate insures minimum cross-reactivity. Prepare fresh diluted
conjugate daily.
f. After covering each smear with 1 drop of appropriate dilution of conjugate, place slides in
a moist chamber to prevent evaporation of staining reagent. After 30 min wash away excess
reagent by dipping slides into phosphate-buffered saline (pH 8.0). Place slides in second bath of
buffered saline for 10 min. Remove, rinse in distilled water, and drain dry. Do not blot.
g. Place a small drop of mounting fluid (pH 9.0) on the smear and cover with a No. 1 1/2
cover slip. Seal edges of cover slip with clear fingernail polish. Examine sealed slides within a
few hours while fluorescence is of optimum intensity. Examine under a fluorescence microscope
unit fitted with appropriate filters.
h. Include a positive control slide with each set of samples. This checks conjugate reactivity
and FA equipment generally.
6. Recording and Interpreting Results
The intensity of organisms fluorescing in any given field is important in assessing positive
Salmonella smears. If the majority of cells present fluoresce (4+ or 3+) the smear is positive.
Carefully scrutinize smears showing only a few scattered fluorescing cells. Critical examination
of cellular morphology may distinguish between these cells and salmonellae. The degree of
fluorescence is the criterion on which positivity is based. Consider weakly fluorescing cells (2+
and 1+) negative. Confirm all positive FA results by cultural techniques (see Section 9260B).
Reaction
Positive
Positive
Negative
Negative
Negative
Description
Brilliant yellow-green fluorescence, cells
sharply outlined
Bright yellow-green fluorescence, cells sharply
outlined with dark center
Dull yellow-green fluorescence, cells not
sharply outlined
Faint green fluorescence discernible in dense
areas, cells not outlined
No fluorescence
Fluorescence
Intensity
4+
3+
2+
1+
0
7. Quantitative Immunofluorescence Microspectrofluorometric Microscopy
To make such analyses use a system consisting of analyzing and illumination sections. The
analyzing section includes an eyepiece monochromator assembly and a
photomultiplier-photometer. The eyepiece uses a beam splitter that reflects to the
monochromator and the observer’s eye, allowing for simultaneous visual observation and
quantitative analysis of the yellow-green fluorescence intensity. The photometer package
provides meter readout in milliamperes so that visual observation of fluorescence can be
correlated with objective reading. Microspectrofluorometry can be done with a conventional
fluorescence microscope.
8. Reference
1. KATZ, I.J. & F.T. BREZENSKI. 1973. Detection of Salmonella by fluorescent antibody.
U.S. Environmental Protection Agency, Edison, N.J.
9. Bibliography
SCHULTE, S.J., J.S. WITZEMAN & W.M. HALL. 1968. Immunofluorescent screening for Salmonella
in foods: comparison with culture methods. J. Amer. Org. Agr. Chem. 51:1334.
THOMASON, B.M. & J.G. WALLS. 1971. Preparation and testing of polyvalent conjugates for F.A.
detection of Salmonellae. Appl. Microbiol. 22:876.
THOMASON, B.M. 1971. Rapid detection of Salmonella microcolonies by fluorescent antibody.
9260 D.
Quantitative Salmonella Procedures
This procedure describes one approach for estimating Salmonella density in water samples.
Other methods have been described in the literature and a comparative study is recommended to
select the best quantitative method for any given application. The following procedure must be
modified for use with solid or semisolid samples.
Because of the high ratio of coliform bacteria to pathogens, large samples (1 L or more) are
required. Any concentration method in Section 9260B.1 may be used but preferably concentrate
the sample by the membrane filter technique (Section 9260B.1d). After blending the membrane
with 100 mL sterile 0.1% (w/v) peptone water, use a quantitative MPN procedure by
proportioning homogenate into a five-tube, three-dilution multiple-tube procedure using either
selenite cystine, selenite-F, or tetrathionate broth as the selective enrichment medium (See
Section 9260B.3). Incubate for 24 h as specified or required for the enrichment medium used and
streak from each tube to plates of brilliant green and xylose lysine desoxycholate agars. Incubate
for 24 h at 35°C. Select from each plate at least one, and preferably two to three, colonies
suspected of being Salmonella, inoculate a slant each of triple sugar iron (TSI) and lysine iron
(LIA) agars, and incubate for 24 h at 35°C. Test cultures giving a positive reaction for
Salmonella by serological techniques (see Section 9260B.5). From the combination of
Salmonella negative and positive tubes, calculate the MPN/1.0 L of original sample (see Section
9221E).
9260 E.
Shigella
Shigellosis is an acute gastrointestinal disease of humans, caused by four species or
serogroups of the genus Shigella, S. dysentariae (Group A), S. flexneri (Group B), S. boydii
(Group C), and S. sonnei (Group D). Shigellae invade the intestinal mucosa, producing dysentery
characterized by abdominal pain, fever, and diarrhea. The infectious dose for Shigella spp. is
low, and most cases result from person-to-person transmission. When outbreaks occur, they are
usually associated with fecal contamination of foods and, less frequently, water. The shigellosis
case rate has gradually risen in the U.S. over the past 30 years from 6 cases/100 000 population
in 1965 to 12 cases/100 000 population in 1995.1 In the U.S., S. sonnei (66.5%) is the most
common cause of shigellosis, followed by S. flexneri (16.4%), S. boydii (1.1%), and S.
dysentariae (0.5%). The serogroup is not reported for 15.5% of cases.
Shigellosis is most common among children. Outbreaks from direct transmission have been
reported in schools, day-care centers, and institutions providing custodial care. Waterborne
outbreaks are associated with fecal contamination together with inadequate chlorination of
private or noncommunity water supplies, as the result of cross-connections between wastewater
and potable water lines, and from exposure to fecally contaminated recreational waters.
Shigellae are sensitive to chlorination at normal levels, and they do not compete favorably
with other organisms in the environment. Their survival time is measured in hours and days, and
is a function of the extent of pollution, as well as physical conditions such as temperature and
pH. Shigellae survive up to 4 d in river water. However, the time required to establish a
laboratory diagnosis by culture of patient specimens (1 to 2 d) makes it improbable that shigellae
can be recovered from an environmental source unless there is a continuous source of
contamination such as wastewater seepage. Shigellae can survive in a viable but nonculturable
state after 21 d.2 The public health significance of nonculturable shigellae in the environment is
unknown.
Methods for the reliable quantitative recovery of shigellae from the environment are not yet
available. Culture of shigellae is usually either not attempted or unsuccessful. Methods that have
resulted in isolation of Shigella include membrane filtration3,4 and centrifugation5,6 with or
without subsequent broth enrichment. Recently, the polymerase chain reaction (PCR) has shown
promise for detection of shigellae in environmental samples.7-9
1. Sampling and Storage
Collect a water sample in a sterile 1-L container. Collect soil, sediment, sludge, or other
samples in plastic bags*#(63) or glass or plastic bottles. Hold samples at 2 to 8°C until they are
processed. Process samples as soon as possible after collection.
2. Enrichment
Choose a selective enrichment medium to minimize accumulation of volatile acid
by-products derived from growth of potentially antagonistic bacteria. Selenite F broth has been
used successfully to recover shigellae from water and sand.5,6 While GN broth facilitates better
recovery of shigellae from stools than Selenite F broth, the only reported attempt to use GN
broth as an enrichment for membrane filters for isolation of shigellae failed to recover the
organism.10
Alternatively, use reduced-strength nutrient medium adjusted to pH 8.0 (0.15 g tryptic soy
broth, added directly to the sample). During outbreak investigations, the enrichment medium
may be made selective by incorporation of antibiotics to which the clinical isolates have shown
resistance, such as tetracycline and streptomycin at concentrations of 150 µg/mL.11
3. Membrane Filter Procedure
This procedure is suitable for low-turbidity potable and surface waters with low
concentrations of coliform bacteria. Filter 100-mL to 1-L samples through 0.45-µm pore size
membranes and place filters face up on the surface of XLD or MacConkey agar plates; incubate
plates at 35°C overnight. Where growth is confluent, sweep growth from plate and inoculate GN
or Selenite F broth enrichments; incubate for 6 h and streak onto MacConkey and XLD plates for
colony isolation. Pick colorless colonies (lactose nonfermenters) from membrane or plates to TSI
and LIA slants; incubate overnight at 35°C. For biochemical reactions and serological grouping,
see ¶ 5 below.
4. Centrifugation Procedure
This procedure is suitable for surface waters, wastewater, and sediments. Centrifuge 200- to
250-mL water samples at 1520 × g for 15 min and pour off all but last 2 mL of supernatant.
Resuspend pellet and add 8 mL Selenite F or GN broth. Incubate suspension for 24 h at 35°C.
Mix suspension and inoculate one loopful to each of several MacConkey and XLD plates. Streak
plates for isolation and incubate overnight at 35°C. Examine plates for colorless colonies, and
pick suspect colonies to TSI and LIA slants; incubate at 35°C overnight. For biochemical
reactions and serological grouping, see ¶ 5 below.
For solid samples (sediments, soil, sludge, etc.) suspend 10 g sample in 100 mL Selenite F or
GN broth and mix thoroughly. Incubate suspension overnight at 35°C. Resuspend sediment and
streak one loopful onto each of several MacConkey and XLD agar plates; incubate overnight at
35°C. Pick colorless colonies to TSI and LIA slants, and proceed as above. For biochemical
reactions and serological grouping, see ¶ 5 below.
5. Biochemical Identification and Serological Grouping
Examine the TSI and LIA slants for the reactions shown in Table 9260:II. Cultures that are
presumptively identified as Shigella spp. are serogrouped by a slide agglutination test using
polyvalent and group specific antisera. Refer cultures to a public health reference laboratory if
molecular typing is desirable for outbreak-related strains.
6. References
1. CENTERS FOR DISEASE CONTROL AND PREVENTION. 1996. Summary of notifiable
diseases, United States 1995. Morbid. Mortal. Week. Rep. 44:1.
2. COLWELL, R.R., P.R. BRAYTON, D.J. GRIMES, D.B. ROSZAK, S.A. HUQ & L.M. PALMER.
1985. Viable but non-culturable Vibrio cholerae and related pathogens in the
environment: implications for release of genetically engineered microorganisms.
Bio/Technology 3:817.
3. DANIELSSON, D. & G. LAURELL. 1968. A membrane filter method for the demonstration
of bacteria by the fluorescent antibody technique. Acta. Path. Microbiol. Scand.
72:251.
4. LINDELL, S.S. & P. QUINN. 1973. Shigella sonnei isolated from well water. Appl.
Microbiol. 26:424.
5. CODY, R.M. & R.G. TISCHER. 1965. Isolation and frequency of occurrence of Salmonella
and Shigella in stabilization ponds. J. Water Pollut. Control Fed. 37:1399.
6. DABROWSKI, J. 1982. Isolation of the Shigella genus bacteria from the beach sand and
water of the bay of Gdansk. Biul. Inst. Med. Morskiej. 33:49.
7. BEJ, A.K., J.L. DICESARE, L. HAFF & R.M. ATLAS. 1991. Detection of Escherichia coli and
Shigella spp. in water by using the polymerase chain reaction and gene probes for uid.
8. ISLAM, M.S., M.K. HASAN, M.A. MIAH, G.C. SUR, A. FELSENSTEIN, M. VENKATESAN, R.B.
SACK & M.J. ALGERT. 1993. Use of the polymerase chain reaction and
fluorescent-antibody methods for detecting viable but nonculturable Shigella
dysenteriae Type 1 in laboratory microcosms. Appl. Environ. Microbiol. 59:536.
9. SETHABUTR, O., P. ECHEVERRIA, C.W. HOGE, L. BODHIDATTA & C. PITARANGSI. 1994.
Detection of Shigella and enteroinvasive Escherichia coli by PCR in the stools of
patients with dysentery in Thailand. J. Diarrh. Dis. Res. 12:265.
10. MAKINTUBEE, S., J. MALLONEE & G. ISTRE. 1987. Shigellosis outbreak associated with
swimming. Amer. J. Pub. Health 77:166.
11. ROSENBERG, M.L., K.K. HAZLET, J. SCHAEFER, J.G. WELLS & R.C. PRUNEDA. 1976.
Shigellosis from swimming. J. Amer. Water Works Assoc. 236:1849.
9260 F.
Pathogenic Escherichia coli
Escherichia coli is a normal inhabitant of the human digestive tract; however, some E. coli
cause diarrheal diseases in humans.1 These pathogenic E. coli are classed into five groups:
enterotoxigenic (ETEC), enterohemorrhagic (EHEC), enteroinvasive (EIEC), enteropathogenic
(EPEC), and the newly recognized group called enteroadherent-aggregative E. coli (EA-AggEC)
for its aggregative or ‘‘stacked-brick’’-like adherence to cultured mammalian cells.2 Pathogenic
E. coli can be grouped on the basis of serology but, because they are classed on the basis of
distinct pathogenic factors, definitive identification requires the determination of the
characteristic virulence properties associated with each group. These include: plasmid-mediated
cell invasion, plasmid-mediated colonization and enteroadherence factors, production of several
potent cytotoxins, hemolysins, as well as heat-labile and stable enterotoxins.3
Although pathogenic E. coli have most often been implicated in foodborne illness, several
major waterborne outbreaks have been reported.4 Outbreaks have involved both water
supplies5-7 and recreational waters.8,9 Some E. coli pathogens have a low infectious dose.
1. Examination Procedures
The pathogenic E. coli groups are phenotypically diverse; hence, no standard
microbiological methods have been developed for these pathogens. Unlike typical E. coli, some
pathogenic groups like EIEC do not ferment lactose3; hence, coliform methods based on lactose
fermentation are not suitable for detection of EIEC. Also, many fecal coliform confirmation or
enrichment procedures use elevated incubation temperature, which is inhibitory to the growth of
EHEC.10 Elevated temperatures and sodium lauryl sulfate used in lauryl tryptose broth (LTB)
for MPN analysis also have been found to cause the loss of plasmid, which encodes many of the
virulence-associated factors.11
Pathogenic E. coli that ferment lactose and are not affected by elevated temperatures still can
be presumptively distinguished from non-E. coli by the MPN fecal coliform procedure (9221E)
or the fecal coliform membrane filter method (9222D) followed by serotyping and virulence
analysis. These methods, as well as methods from other sources,12 also have been modified to
detect specific pathogenic groups. Regardless of the method, however, when testing for
pathogenic E. coli, first identify isolates as E. coli either by conventional biochemical testing or
by using commercially available biochemical identification kits (see Section 9260B.4) before
serotyping and assaying for the virulence factors associated with the respective pathogenic
groups.
a. EHEC O157:H7: The following procedure is a modification of the standard total coliform
fermentation technique (9221B) for detecting E. coli O157:H7 in water.13 Inoculate a 100-mL
sample into 50 mL 3× lauryl tryptose broth (LTB) and incubate at 35°C for 24 h. Serially dilute
the sample, spread plate (0.1 mL) onto sorbitol MacConkey agar (SMAC)*#(64) and incubate at
35°C for 18 to 24 h. EHEC O157:H7 form colorless colonies because they do not ferment, or are
slow fermenters of, sorbitol. Pick ten sorbitol-negative colonies, transfer individually into
LTB-MUG (4-methylumbelliferone glucuronide; 0.1 g/L)14 and incubate at 35°C for 18 to 24 h.
EHEC O157:H7 ferment lactose, but do not have β-glucuronidase activity to hydrolyze MUG, so
cultures will appear gas-positive and nonfluorescent. Assay these for positive glutamate
decarboxylase activity,13 then identify biochemically as E. coli.
Larger volumes of sample also may be examined by the following procedure modified from
a procedure for detecting O157: H7 in food.15 This procedure has not been tested for use in
water analysis; however, it has been used extensively to detect O157: H7 bacteria in apple juice.
Centrifuge 200 mL sample at 10 000 × g for 10 min. Resuspend pellet in 225 mL EHEC
enrichment broth (EEB) and incubate at 35°C for 6 h. Spread plate 0.1 mL from EEB and a 1:10
dilution of EEB onto tellurite cefixime SMAC (TC SMAC). Both EEB and TC SMAC contain
antibiotics to reduce growth of normal flora bacteria; therefore, they are best suited for highly
contaminated samples. Incubate EEB sample and TC SMAC at 35°C for 18 to 24 h. Observe TC
SMAC plates for isolated, colorless colonies. If none are evident, serially dilute the overnight
EEB sample and replate onto TC SMAC. Test colorless colonies for positive indole reaction and
identify biochemically as E. coli before serotyping and virulence analysis for the Shiga toxin
genes.
b. EPEC, ETEC, EIEC: With the exception of EIEC, use either the MPN fecal coliform
procedure (9221E) or the fecal coliform membrane filter method (9222D) for presumptive
isolation of these pathogenic E. coli groups from water. Alternatively plate presumptive positive
samples onto selective media, such as LES Endo and MacConkey (MAC) agars (see Section
9221B.3, Completed Phase). In food analysis, L-EMB agar also has been used. For EIEC, which
ferment lactose slowly or not at all, the MPN method is not useful; however, the membrane filter
method (9222D) can be used. In food testing for EIEC, Hektoen agar (HE), Salmonella-Shigella
(SS) agar and MAC are used for selective plating, but HE and MAC appear less inhibitory and
are best suited for the isolation of EIEC10. In the analysis of each pathogenic E. coli group,
preferably pick 10 typical (lactose-positive) and 10 atypical (lactose-negative) colonies for
biochemical identification. Identify all isolates as E. coli before serological typing and analysis
for the group-specific virulence factors.
2. Serotyping
For definitive identification, serotype for the O:H antigens any isolates presumptively
identified as pathogenic E. coli by microbiological methods. Polyvalent antisera are available
commercially, but only for the common serotypes. Several anti-O157 and anti-H7 latex
agglutination kits are available for typing O157:H7 isolates. Serotype information also is
essential for epidemiological investigations.
3. Virulence Analysis
The pathogenic potential of an E. coli isolate can be determined only by testing for its
distinctive virulence properties. A simple antibody-bound latex agglutination kit and several
enzyme linked immunosorbent assay kits are available for testing Shiga cytotoxins of
EHEC†#(65). An agglutination kit also is available for testing labile and stable enterotoxins of
ETEC,‡#(66) but analysis of other virulence factors may require bioassays using animal models,
tissue cultures, or other antibody and nucleic-acid-based molecular methods. A partial listing of
commercially available assays and media for pathogenic E. coli is available.12 Most of the
assays are specific for EHEC O157:H7 and introduced only recently for food analysis; hence,
few have been evaluated by collaborative studies.
4. References
1. ORSKOV, F. & ORSKOV, I. 1992. Escherichia coli serotyping and disease in man and
animals. Can. J. Microbiol. 38:699.
2. VIAL, P.A., R. ROBINS-BROWNE, H. LIOR, V. PRADO, J.B. KAPER, J.P. NATARO, D.
MENEVAL, A.-E.-D. ELSAYED & M.M. LEVINE. 1988. Characterization of
enteroadherent-aggregative Escherichia coli, a putative agent of diarrheal disease. J.
Infect. Dis. 158:70.
3. LEVINE, M.M. 1987. Escherichia coli that cause diarrhea: enterotoxigenic,
enteropathogenic, enteroinvasive, enterohemorrhagic and enteroadherent. J. Infect. Dis.
155:377.
4. FENG, P. 1995. Escherichia coli serotype O157:H7: novel vehicles of infection and
emergence of phenotypic variants. Emerging Infec. Dis. 2:47.
5. SCHROEDER, S.A., J.R. CALDWELL, T.M. VERNON, P.C. WHITE, S.I. GRANGER & J.V.
BENNETT. 1968. A waterborne outbreak of gastroenteritis in adults associated with
Escherichia coli. Lancet 1:737.
6. ROSENBERG, M.L., J.P. KOPLAN, I.K. WACHSMUTH, J.G. WELLS, E.J. GANGAROSA, R.L.
GUERRANT & D.A. SACK. 1977. Epidemic diarrhea at Crater Lake from enterotoxigenic
Escherichia coli. Ann. Intern. Med. 86:714.
7. SWERDLOW, D.L., B.A. WOODRUFF, R.C. BRADY, P.M. GRIFFIN, S. TIPPEN, H.D. DONNELL,
E. GELDREICH, B.J. PAYNE, A. MEYER, J.G. WELLS, K.D. GREENE, M. BRIGHT, N.H. BEAN
& P.A. BLAKE. 1992. A waterborne outbreak in Missouri of Escherichia coli O157:H7
associated with bloody diarrhea and death. Ann. Intern. Med. 117:812.
8. KEENE, W.E., J.M. MCANULTY, F.C. HOESLY, L.P. WILLIAMS, K. HEDBERG, G.L. OXMAN,
T.J. BARRETT, M.A. PFALLER & D.W. FLEMING. 1994. A swimming-associated outbreak
of hemorrhagic colitis caused by Escherichia coli O157:H7 and Shigella sonnei. N.
England J. Med. 331:579.
9. BREWSTER, D.H., M.I. BROWNE, D. ROBERTSON, G.L. HOUGHTON, J. BIMSON & J.C.M.
SHARP. 1994. An outbreak of Escherichia coli O157 associated with a children’s
paddling pool. Epidemiol. Infect. 112:441.
10. DOYLE, M.P. & V.V. PADHYE. 1989. Escherichia coli. In M.P. Doyle, ed. Foodborne
Bacterial Pathogens. Marcel Dekker, Inc., N.Y.
11. HILL, W.E. & C.L. CARLISLE. 1981. Loss of plasmids during enrichment for Escherichia
coli. Appl. Environ. Microbiol. 41:1046.
12. U.S. FOOD AND DRUG ADMINISTRATION. 1995. Bacteriological Analytical Manual, 8th
ed. Assoc. Official Analytical Chemists International, Gaithersburg, Md.
13. RICE, E.W., C.H. JOHNSON & D.J. REASONER. 1996. Detection of Escherichia coli
O157:H7 in water from coliform enrichment cultures. Lett. Appl. Microbiol. 23:179.
14. FENG, P. & P.A. HARTMAN. 1982. Fluorogenic assay for immediate confirmation of
Escherichia coli. Appl. Environ. Microbiol. 43:1320.
15. HITCHINS, A.D., P. FENG, W.D. WATKINS, S.R. RIPPEY & L.A. CHANDLER. 1995.
Escherichia coli and the coliform bacteria. In Bacteriological Analytical Manual, 8th
ed. Assoc. Official Analytical Chemists International, Gaithersburg, Md.
9260 G.
Campylobacter jejuni
Campylobacters are commonly found in the normal gastrointestinal and genitourinary flora
of wild animals, birds, and domestic animals including sheep, cattle, swine, goats, and chickens.1
Campylobacter infections often are acquired by the fecal oral route, often as zoonoses through
exposure to infected animals. Large outbreaks have resulted from contaminated milk, uncooked
meat or fowl, and contaminated water systems.2 Campylobacter has been reported to be the most
common cause of bacterial enteritis worldwide.3
Waterborne transmission of Campylobacter has resulted from drinking untreated surface
water, contamination of groundwater with surface water, faulty disinfection, and contamination
by wild bird feces.4 In remote mountain areas, the infection has been associated with drinking
surface water from cold mountain streams.5 Occurrence of campylobacters in surface water is
variable and appears to be seasonally dependent, with lowest levels occurring in summer.
Survival in surface water is affected by both temperature and sunlight.6 Between 1978 and 1986,
57 outbreaks of campylobacteriosis were reported, including 11 waterborne outbreaks, 7 of
which occurred in community water supplies.
1. Water Collection and Filtration Method
Collect large-volume water samples in sterile 10-L plastic containers. Process samples
immediately after collection or store at 4°C and process as soon as possible. Filter one to several
liters of the water through a 0.45- or 0.22-µm-pore-size, 47-mm-diam, cellulose nitrate
membrane filter. Remove filter and place face down on selective medium (see isolation section).
Incubate microaerophilically at 42°C for 24 h. Remove filter from the plate and place it face
down on another selective plate. Incubate both plates at 42°C for up to 5 d.7
For turbid water pre-filtration is necessary. Use a stainless steel filtration device with a 1.5-L
reservoir.*#(67) Assemble with the following filter sequence: Place a 142-mm, 3.0-µm filter on
the screen inside reservoir with a 124-mm prefilter on top. In the bottom tubing adapter place a
47-mm, 1.2-µm filter. Then place Swinnex filter holders in parallel with a 47-mm, 0.65-µm filter
in the upstream filter holder and a 47-mm, 0.45-µm filter in the downstream holder. Add 1 L
sample to the reservoir, seal, and apply pressure of about 350 kPa. After filtration, remove the
0.45-µm pore-size filter and culture on selective plate medium as described above.
2. Isolation
Campylobacter isolation requires use of selective media containing antimicrobial agents,
microaerophilic atmosphere (5% O2, 10% CO2, and 85% N2), and 42°C incubation temperature,
to suppress the growth of most common bacteria.8 The thermophilic campylobacters (C. jejuni,
C. coli, C. lari, and C. upsaliensis) grow well at 42°C. However, other campylobacters (C. jejuni
subsp. doylei and C. fetus) do not grow well at 42°C; incubate plates at both 37°C and 42°C for
optimal isolation of these bacteria.9 Microaerophilic conditions can be provided by using
commercially available systems and equipment.†#(68)
Several selective media for plating campylobacters are commercially available. Skirrow’s
medium contains blood agar base with lysed horse blood, trimethoprim, vancomycin, and
polymixin B. Campy-BAP contains Brucella agar base with sheep blood, trimethoprim,
vancomycin, polymixin B, amphotericin B, and cephalothin (to which some campylobacters are
sensitive). Butzler’s medium contains thioglycollate agar with sheep blood, bacitracin,
novobiocin, cycloheximide, and cefazolin. Preston’s medium contains Campylobacter agar base
with horse blood, cycloheximide, rifampicin, trimethoprim, and polymyxin B. Other media, such
as Campylobacter blood-free selective medium and Campylobacter charcoal differential agar,
can be used to isolate campylobacters.10 Use of enrichment broth will improve recovery of
campylobacters.
Several enrichment media, such as Campylobacter broth, Campy-thio broth, Gifu
anaerobe-modified semisolid medium, and Preston medium, are used to enhance recovery of
campylobacters.9 Add 10 mL water sample to 10 mL Campylobacter enrichment broth tubes in
duplicate, and incubate cultures at 37°C and 42°C for 8 h or overnight. Pre-enrichment of water
sample in a selective enrichment broth for 4 h at 37°C may be important for recovery of stressed
cells of C. jejuni that show less tolerance to elevated growth temperatures. For pre-enrichment of
water sample, add 10 mL water to 10 mL enrichment medium and incubate culture for 4 h at
37°C, then transfer the cultures to another incubator at 42°C for overnight incubation.11,12
C. jejuni may be induced to a nonculturable state in water, and it is not clear whether
pre-enrichment or enrichment will facilitate isolation of these bacteria.13 Use of a decreased
substrate concentration enhances metabolic activity in nonculturable campylobacters from
water.14
3. Identification
a. Culture examination: Examine Campylobacter plates at 24 and 48 h for characteristic
colonies, which can range from flat, spreading colonies that cover the entire surface of the plate,
to very small, convex, translucent colonies. Colony colors range from gray to yellowish or
pinkish.
b. Microscopy identification: Campylobacter spp. do not stain well by the conventional
Gram stain. If safranin is used as a counterstain, apply it for 2 to 3 min; carbol fuchsin is a better
alternative. Even 24-h cultures of campylobacters appear pleomorphic in stained smears, and
cells range from small Gram-negative rods and coccoid forms to longer rods that may show an
‘‘S’’ or seagull shape, and long spirals, particularly from older cultures.15
c. Motility test: Campylobacter normally are motile by a single polar flagellum at one or
both ends. Suspend cells in Mueller-Hinton or nutrient broth, and observe motility using phase
microscopy or brightfield microscopy with reduced illumination. Do not use saline or distilled
water because they may inhibit motility.8 Young cells are 0.2 to 0.8 µm wide by 1.5 to 5 µm
long, curved or spiral, and motile with darting or corkscrew-like motion.16
d. Biochemical tests: Despite numerous studies, campylobacters remain relatively difficult
to rapidly identify, classify, and type biochemically.17 Campylobacters do not ferment or oxidize
carbohydrates, and they are inert in most biochemical media used to characterize bacterial
isolates.18 Although no standard methods for the characterization of campylobacters have been
published, oxidase, catalase, nitrite and nitrate reduction, H2S production, hippurate hydrolysis,
resistance to various agents, temperature tolerances, and growth requirements are among the
common phenotypic tests used to characterize campylobacters.3
4. Serological Identification Tests
Commercially available kits‡#(69) for serotyping campylobacters are available. These kits
use latex particles coated with polyvalent immunoglobulins for several Campylobacter species.
They are designed for rapid presumptive identification of the thermophilic, enteropathogenic
Campylobacter species (C. jejunei, C. coli, and C. lari); use in accordance with manufacturer’s
instructions.19
Other techniques that are not widely available in all laboratories include lectin agglutination,
cellular fatty acid profiles, nucleic acid probes, polymerase chain reaction, and other genomic
methods that can be used in reference and research laboratories for detection and identification
of campylobacters.3
5. References
1. RYAN, K.J. 1990. Vibrio and Campylobacter. In J.C. Sherris, ed. Medical Microbiology:
An Introduction to Infectious Diseases. Elsevier, New York, N.Y.
2. BARON, E.J., R.S. CHANG, D.H. HOWARD, J.N. MILLER & J.A. TURNER, eds. 1994. Medical
Microbiology: A Short Course. Wiley-Liss, New York, N.Y.
3. ON, S.L.W. 1996. Identification methods for campylobacters, helicobacters, and related
organisms. Clin. Microbiol. Rev. 9:405.
4. TAUXE, R.V. 1992. Epidemiology of Campylobacter jejuni infections in the United
States and other industrialized nations. In I. Nachamkin, M.J. Blaser & L.S. Tompkins,
eds. Campylobacter jejuni: Current Status and Future Trends. American Soc.
5. TAYLOR, D.N., K.T. MCDERMOTT, J.R. LITTLE, J.G. WELLS & M.J. BLASER. 1983.
Campylobacter enteritis from untreated water in the Rocky Mountains. Ann. Intern.
Med. 99:38.
6. VOGT, R.L., H.E. SOURS, T. BARRETT, R.A. FELDMAN, R.J. DICKINSON & L. WITHERELL.
1982. Campylobacter enteritis associated with contaminated water. Ann. Intern. Med.
96:292.
7. PEARSON, A.D., M. GREENWOOD, T.D. HEALING, D. ROLLINS, M. SHAHAMAT, J.
DONALSON & R.R. COLWELL. 1993. Colonization of broiler chickens by waterborne
Campylobacter jejuni. Appl. Environ. Microbiol. 59:987.
8. ISENBERG, H.D., ed. 1992. Clinical Microbiology Procedures Handbook. Vol. 1.
American Soc. Microbiology, Washington, D.C.
9. GOOSSENS, H. & J.P. BUTZLER. 1992. Isolation and identification of Campylobacter spp.
In I. Nachamkin, M.J. Blaser & L.S. Tompkins, eds. Campylobacter jejuni: Current
Status and Future Trends. American Soc. Microbiology, Washington, D.C.
10. PARKS, L.C., ed. 1993. Handbook of Microbiological Media. CRC Press, Boca Raton,
Fla.
11. HUMPHREY, T.J. 1989. An appraisal of the efficacy of preenrichment for the isolation of
Campylobacter jejuni from water and food. J. Appl. Bacteriol. 66:119.
12. HUMPHREY, T.J. 1986. Techniques for the optimum recovery of cold injured
Campylobacter jejuni from milk or water. J. Appl. Bacteriol. 61:125.
13. ROLLINS, D.M. & R.R. COLWELL. 1986. Viable but nonculturable stage of
Campylobacter jejuni and its role in survival in the natural aquatic environment. Appl.
14. ROLLINS, D.M. 1987. Characterization of Growth, Decline, and the Viable but
Nonculturable State of Campylobacter jejuni. Ph.D dissertation, Univ. Maryland,
College Park.
15. KAPLAN, R.L. & A.S. WEISSFELD. 1994. Campylobacter, Helicobacter and related
organisms. In B.J. Howard et al., eds. Clinical and Pathogenic Microbiology, 2nd ed.
Mosby, St. Louis, Mo.
16. BEUCHAT, L.R. 1986. Methods for detecting and enumerating Campylobacter jejuni and
Campylobacter coli in poultry. Poultry Sci. 65:2192.
17. DUBREUIL, J.D., M. KOSTRZYNSKA, S.M. LOGAN, L.A. HARRIS, J.W. AUSTIN & T.J. TRUST.
1990. Purification, characterization, and localization of a protein antigen shared by
thermophilic campylobacters. J. Clin. Microbiol. 28:1321.
18. CARDARELLI-LEITE, P., K. BLOM, C.M. PATTON, M.A. NICHOLSON, A.G. STEIGERWALT,
S.B. HUNTER, D.J. BRENNER, T.J. BARRETT & B. SWAMINATHAN. 1996. Rapid
identification of Campylobacter species by restriction fragment length polymorphism
analysis of a PCR-amplified fragment of the gene coding for 16S-rRNA. J. Clin.
Microbiol. 34:62.
19. HODINKA, R.L. & P.H. GILLIGAN. 1988. Evaluation of the Campyslide agglutination test
for confirmatory identification of selected Campylobacter species. J. Clin. Microbiol.
26:47.
9260 H.
Vibrio cholerae
Vibrio cholerae is the causative agent of cholera, a waterborne illness with symptoms
ranging from mild to severe and potentially fatal diarrheal disease.1,2 This is a well-defined
species on the basis of biochemical tests and DNA studies, but the serotypes within the species
can be quite diverse in their ability to produce infection. The O1 serogroup is associated with
epidemic and pandemic cholera, especially in developing countries. The current (seventh)
pandemic has affected over 100 countries, including the United States, with over one million
reported cases and 10 000 deaths.3 The newly identified O139 Bengal serogroup4 also is capable
of producing epidemic cholera. In contrast, the great majority of non-O1/non-O139 strains,
which are more common in the environment, do not produce cholera toxin, and are not
associated with epidemic cholera. However, these strains occasionally are associated with
potentially fatal extra-intestinal infections. V. cholerae occurs as part of the normal microflora in
estuarine areas, with non-O1/non-O139 strains being much more common than are O1 strains.
1. Concentration Techniques
Levels of V. cholerae in natural waters and sewage usually are quite low. Thus, methods of
concentration or enrichment usually are employed. One method for isolating V. cholerae O1
from contaminated waters is placement of Moore swabs in flowing wastewater for periods up to
1 week, followed by placement into enrichment media at a 1:1 (weight/volume) ratio.5
2. Enrichment Procedures
Samples are enriched in alkaline peptone broth (1% peptone, 1% NaCl, pH 8.4), using
appropriate concentration of broth relative to sample volume. Incubate enrichment cultures for 6
to 8 h at 35°C, then streak a loopful of the enrichment broth onto thiosulfate-citrate-bile
salts-sucrose (TCBS) agar and incubate these plates at 35°C for 18 to 24 h.6 Other enrichment
and plating media have been reviewed.7,8
3. Selective Growth
Suspected V. cholerae colonies appear yellow, a result of sucrose fermentation. A variety of
other sucrose-fermenting vibrios also appear on TCBS, however, including V. fluvialis, V.
furnissii, V. alginolyticus, V. metschnikovii, V. cincinnatiensis, and V. carchariae.2
4. Presumptive Tests to Differentiate V. cholerae
The following key tests are used to identify V. cholerae:
Test
Reaction
Gram-negative rod
Cytochrome oxidase
Glucose fermented (no gas)
Growthin nutrient broth:
No NaCl added
8% NaCl added
+
+
+
Arginine dihydrolase
−
+
+
Ornithine decarboxylase
ONPG hydrolysis
+
−
After isolation on TCBS, streak presumptive V. cholerae isolates to a nonselective medium,
such as trypticase soy agar, containing a minimum of 0.5% NaCl.
5. Classification of Isolates as V. cholerae
The tests listed below may be used for a more extensive phenotypic characterization of V.
cholerae.7 To determine the serogroup, use agglutination assays.
Test
ONPG
Nitrate reduction
Indole
0/129 sensitivity:
10 mg
150 mg
Swarming
Luminescence
Thornley’s arginine dihydrolase
Lysine decarboxylase
Ornithine decarboxylase
Growth at 42°C
Growth at % NaCl:
0%
3%
6%
8%
10%
Voges-Proskauer reaction
Gas from glucose fermentation
Fermentation to acid:
L-Arabinose
m-Inositol
D-Mannose
Sucrose
Enzyme production:
Alginase
Amylase
Chitinase
Gelatinase
Lipase
Utilization as sole source of carbon:
Reaction
+
+
+
+
+
−
v*
−
+
+
+
+
+
v
−
−
v
−
−
−
v
+
−
+
+
+
+
Test
Reaction
γ-Aminobutyrate
Cellobiose
−
L-Citruline
−
Ethanol
D-Gluconate
−
+
D-Glucuronate
−
L-Leucine
−
Putrescine
−
+
−
Sucrose
D-Xylose
−
* v = variable, differs for strains within the species.
6. Serological Identification
Slide agglutination with polyvalent antisera can be used to identify the serogroups of V.
cholerae. Polyvalent antiserum for V. cholerae O1 is available commercially.*#(70) The O1
serogroup can be further divided into two primary serotypes, Ogawa and Inaba.
7. Biotypes of Serogroup O1 V. cholerae
V. cholerae can be divided into two biotypes or biovars, classical and El Tor, which differ in
several characteristics. The El Tor biotype currently is the most important biotype.
Biovar
Test7
Classical
El Tor
Hemolysis of sheep erythrocytes
−
v*
Voges-Proskauer reaction
−
+
Chicken erythrocyte agglutination
−
+
Antibiotic sensitivity:
Polymyxin B (50 IU)
+
−
Lysis
No lysis
No lysis
Lysis
Bacteriophage susceptibility:
Mukerjee classical phage IV
Mukerjee El Tor phage 5
* v = different reaction within the serovar.
8. Other Procedures
Environmental samples also may be examined by fluorescent-antibody techniques, but the
number of V. cholerae cells in aquatic samples is generally quite low.7 Nucleic acid probes are
not routinely used for the identification of V. cholerae, although DNA probes are extremely
useful in determining which strains of this species contain the cholera toxin gene.2 This
distinction is especially important in examining environmental isolates of V. cholerae because
the great majority of these strains lack the cholera toxin gene.
9. References
1. KAPER, J.B., J.G. MORRIS, JR. & M.M. LEVINE. 1995. Cholera. Clin. Microbiol. Rev. 8:48.
2. OLIVER, J.D. & J.B. KAPER. 1997. Vibrio species. In M.P. Doyle, L.R. Beuchat & T.J.
Montville, eds. Fundamentals of Food Microbiology. American Soc. Microbiology,
Washington, D.C.
3. CENTERS FOR DISEASE CONTROL. 1995. Update: Vibrio cholerae O1— Western
hemisphere, 1991–1994, and V. cholerae O139—Asia, 1994. Morbid. Mortal. Week.
Rep. 44:215.
4. ALBERT, M.J. 1994. Vibrio cholerae O139 Bengal. J. Clin. Microbiol. 32:2345.
5. BARRETT, T.J., A. BLAKE, G.K. MORRIS, N.D. PUHR, H.B. BRADFORD & J.G. WELLS. 1980.
Use of Moore swabs for isolating Vibrio cholerae from sewage. J. Clin. Microbiol.
11:385.
6. SPECK, M.L., ed. 1984. Compendium of Methods for the Microbiological Examination
of Foods, 2nd ed. American Public Health Assoc., Washington, D.C.
7. WEST, P.A. & R.R. COLWELL. 1984. Identification and classification of
Vibrionaceae—an overview. In R.R. Colwell, ed. Vibrios in the Environment. John
Wiley & Sons, New York, N.Y.
8. KAYSNER, C.A. & W.E. HILL. 1994. Toxigenic Vibrio cholerae O1 in food and water. In
I.K. Wachsmuth, P.A. Blake & O. Olsvik, eds. Vibrio cholerae and Cholera: Molecular
to Global Perspectives. ASM Press, Washington, D.C.
9260 I.
Leptospira
Leptospira spp. are motile, aerobic spirochetes that require fatty acids for growth.1 Serum or
polysorbate enrichments must be incorporated into artificial media, and some pathogenic strains
may require CO2 upon initial isolation. Leptospires are divided into two groups, based on their
pathogenicity and growth characteristics. The saprophytic leptospires are assigned to the Biflexa
Complex, and the pathogenic leptospires make up the Interrogens Complex. Pathogenic strains
have an optimal growth temperature of 28 to 30°C, and they grow over a pH range from 5.2 to
7.7. Saprophytic strains prefer a growth temperature between 5 and 10°C below pathogenic
strains. Leptospires prefer alkaline conditions, and they persist longest in warm, moist
environments protected from sunlight. Under favorable temperature and pH conditions,
leptospires survive for 3 to 5 d in damp soil and up to 10 d in natural waters. They survive for 12
to 14 h in undiluted wastewater, up to 3 d in aerated wastewater, and up to 4 weeks in sterile
tapwater at pH 7. Nonpathogenic leptospires are ubiquitous, and they have been isolated from
municipal water supplies.2 Generally, pathogenic leprospires require an animal host and do not
survive and propagate in the environment.
Leptospirosis is a worldwide zoonotic disease of wild animals.3 Reservoirs of leptospires in
wildlife include deer, foxes, raccoons, skunks, opossums, muskrats, and rodents. Domestic
animals harboring leptospires include horses, cattle, goats, pigs, and sheep. Dogs may become
infected but cats are spared. Humans are incidental hosts.
Humans acquire leptospirosis (Weil’s disease) directly from animals, and from occupational
or recreational exposure to urine-contaminated water4-6 or environmental surfaces. Swimming
and other water sports,7 travel to tropical areas with occupational or recreational exposure to
surface waters,8 and natural disasters that affect sewer systems and runoff9,10 increase risk of the
disease. Outbreaks of leptospirosis associated with drinking water are extremely unusual, and are
invariably caused by contamination of domestic water reservoirs with urine of infected
rodents.11
Leptospirosis ranges from mild nonspecific febrile illness to severe or fatal renal, hepatic, or
meningeal disease.12,13 Leptospires enter through imperfections in the skin, through mucous
membranes, or by ingestion of contaminated water. Urine of infected animals and humans may
contain 106 to 108 organisms/ mL. Leptospires may be shed into the environment up to 3 months
after clinical recovery from disease.
Diagnosis of disease in animals and humans usually is based upon serology, darkfield
examination of urine sediments, examination of histopathological stains, or culture of the
organism from urine or tissues. Recently polymerase chain reaction (PCR) methods have been
introduced for diagnosis and typing of leptospires.
While leptospirosis remains relatively common in tropical regions of the world, only 40 to
120 cases/year have been reported in the U.S. over the past 30 years. Leptospirosis was dropped
from the list of notifiable diseases in 1994.
Leptospires are recovered from environmental sources with great difficulty.14-17 Because
both saprophytic and pathogenic strains of leptospires may be recovered from environmental
samples, their presence has no public health significance apart from an epidemiological context.
1. Sample Collection
Collect water samples of 100 mL to 1 L in sterile containers for transport to the laboratory at
ambient temperature within 72 h of collection. Multiple samples from each sample site usually
are required for successful isolation because finding leptospires in 10 to 20% of samples of
surface waters receiving farm runoff is considered a high yield. Leptospires find their ecological
niche at the interface between sediment and shallow water. Gently agitate the water to bring
some of the sediment to the surface of shallow bodies of water to improve the probability of
recovering organisms.18 For soil samples, collect 10 to 20 g of soil in sterile bottles or plastic
bags. Use a small, tightly sealed container to protect sample from drying. A small amount of
sterile deionized water may be added to soil samples to prevent drying.
2. Sample Processing
Centrifuge a portion of a water sample at 5000 × g for 10 min and examine sediment by
darkfield microscopy for leptospires. Their presence indicates that conditions are favorable for
leptospire survival, but does not differentiate saprophytic from pathogenic forms. In the
laboratory, thoroughly mix soil samples with three volumes of sterile deionized water and let
coarse particulate material settle by gravity. Process remaining suspension as a water sample.
Leptospira can pass through 0.22-µm membrane filters (¶ a below); this ability has been
exploited to separate them from other bacteria in environmental samples and in mixed cultures.
Similarly, guinea pig inoculation (¶ b below) has been used as a biological filter for isolation of
leptospires from contaminated samples.
a. Filtration method: Filter surface water samples through filter paper*#(71) to remove
coarse debris before membrane filtration. Occasionally, samples may have to be passed through
a series of prefilters of decreasing pore sizes (8-µm, 4-µm, 1-µm, 0.65-µm, and 0.45-µm) to
prevent clogging of the final 0.22-µm filter.
b. Animal inoculation method: Filter water through a 0.45-µm membrane filter and inoculate
1 to 3 mL intraperitoneally into weanling guinea pigs. After 3 to 6 d, inject a small amount of
sterile saline and withdraw fluid for darkfield examination. If leptospires are seen, perform a
cardiac puncture to obtain blood for inoculation of culture media. If no leptospira are seen by
darkfield examination, record rectal temperatures daily until a fever spike indicates infection,
then repeat the darkfield examination of peritoneal fluid for leptospires. Exsanguinate guinea
pigs at 4 weeks and save serum for serological tests. Culture blood, kidney, and brain of guinea
pigs with serological evidence of infection. Details of the method are described elsewhere.19
3. Culture
Cultures of environmental samples usually will be contaminated with other bacteria unless
the samples are filtered through a 0.22-µm membrane filter before inoculation. Filtration also
may be used to isolate leptospires from mixed cultures, by direct filtration or another method.20
Unless sample filtration is used in conjunction with selective media or animal inoculation, a
culture contamination rate of 60 to 80% is not uncommon. The amount of sample cultured will
depend on the amount of particulate material in the sample. Generally, culture sample volumes
from a few drops to 3.5 mL.
a. Culture media: Pathogenic leptospires have been cultured in liquid, semisolid, and solid
media, but not all pathogenic strains will grow on solid media. Optimal pH of culture media is
7.2 to 7.4 and optimal incubation temperature is 30°C. Leptospires are sensitive to detergents, so
keep glassware free of detergent residues. When using serum enrichments in culture media, use
serum free of antibody to leptospires. Bovine serum albumin shows manufacturer and lot
variations; test new batches for their ability to support growth of leptospires.
Modifications of the Ellinghausen-McMullough formulation (EMJH) that incorporate
bovine serum albumin fraction V and polysorbates are used as serum replacements.21-24 EMJH
base is available commercially. Neomycin is used in culture media at concentrations between 5
and 25 µg/mL to inhibit competing microflora, but it may be toxic to some strains.25
5-fluorouracil is used at 100 or 200 µg/mL in culture media, but it too is toxic for some strains,
particularly at concentrations above 100 µg/mL.26
b. Culture methods:
1) Direct culture method—To recover leptospires from surface waters, place a few drops of
water in EMJH liquid medium and incubate overnight at 30°C. Filter inoculated medium through
a 0.22-µm membrane filter into a sterile tube and reincubate at 30°C for up to 6 weeks.
2) Dilution method—When samples may contain reasonable numbers of organisms in the
presence of inhibitors or competing microflora, prepare 10-fold dilutions in duplicate, and
inoculate 0.1 mL undiluted sample and each dilution into EMHJ medium. One tube of each pair
may be made selective by addition of a single 30-µg neomycin antimicrobial susceptibility disk
to the media before incubation. Incubate cultures at 20 to 30°C for up to 4 months.
3) Animal inoculation method—Add 1 to 2 drops of heart blood from infected guinea pigs to
each of three to five tubes of EMJH medium. Incubate cultures at 20°C for up to 4 months.
c. Culture examination: Leptospires usually are detected in cultures of environmental
samples within 7 to 14 d; however, incubate and examine cultures weekly for 6 weeks before
discarding them as negative. Observe tubes for a lightly turbid ring of growth just below the
surface of the medium. This band of maximum turbidity at the zone of optimal oxygen tension is
referred to as Dinger’s ring. Remove a drop of the culture weekly for darkfield examination and
prepare subcultures if motile leptospires are observed. Generally, saprophytic leptospires grow at
lower temperatures, and form rings closer to the surface of culture media than pathogenic
serovars. Cultures remain viable in semisolid media for at least 8 weeks at room temperature.
4. Identification
Experience and skill are required to differentiate artifacts from leptospires by darkfield
microscopy. The biochemical tests previously thought to differentiate between pathogenic and
saprophytic serovars do not reliably predict pathogenicity of leptospires, and they are not
recommended. Leptospira are identified to serogroup by the microscopic agglutination test using
reference antisera. Identification to serovar requires use of adsorbed antisera that are available
only in reference laboratories. Over 200 serotypes of Leptospira are known.
5. References
1. FAINE, S. 1992. The genus Leptospira. In A. Balows, H.G. Truper, M. Dworkin, W.
2.
3.
4.
5.
6.
7.
8.
9.
10.
Harder & K.H. Schleifer, eds. The Prokaryotes, Vol. IV. Springer-Verlag, New York,
N.Y.
HENRY, R.A. & R.C. JOHNSON. 1978. Distribution of the genus Leptospira in soil and
water. Appl. Envrion. Microbiol. 35:492.
MICHNA, S.W. 1970. Leptospirosis. Vet. Record 86:484.
ANDERSON, D.C., D.S. FOLLAND, M.D. FOX, C.M. PATTON & A.F. KAUFMANN. 1978.
Leptospirosis: a common-source outbreak due to leptospires of the grippotyphosa
serogroup. Amer. J. Epidemiol. 107:538.
COGGINS, W.J. 1962. Leptospirosis due to Leptospira pomona: an outbreak of nine
cases. J. Amer. Med. Assoc. 181:1077.
VENKATARAMAN, K.S. & S. NEDUNCHELLIYAN. 1992. Epidemiology of an outbreak of
leptospirosis in man and dog. Comp. Immun. Microbiol. Infect. Dis. 15:243.
SHAW, R.D. 1992. Kayaking as a risk factor for leptospirosis. Missouri Med. 89:354.
VAN CREVEL, R., P. SPEELMAN, C. GRAVEKAMP & W.J. TERPSTRA. 1994. Leptospirosis
in travelers. Clin. Infect. Dis. 19:132.
FUORTES, L. & M. NETTLEMAN. 1994. Leptospirosis: a consequence of the Iowa flood.
Iowa Med. 84:449.
KAT, A.R., S. MANEA & D.M. SASAKI. 1991. Leptospirosis on Kauai: investigation of a
common source waterborne outbreak. Amer. J. Pub. Health 81:1310.
11. CACCIAPUOTI, B., L. CICERONI, C. MAFFEI, F. DI STANISLAO, P. STRUSI, L. CALEGARI, R.
LUPIDI, G. SCALISE, G. CAGNONI & G. RENGA. 1987. A waterborne outbreak of
leptospirosis. Amer. J .Epidemiol. 126:535.
12. HEATH, C.W., A.D. ALEXANDER & M.M. GALTON. 1965. Leptospirosis in the United
States (concluded). Analysis of 483 cases in man, 1949–1961. N. England J. Med.
272:915.
13. HEATH, C.W., A.D. ALEXANDER & M.M. GALTON. 1965. Leptospirosis in the United
States. Analysis of 483 cases in men, 1949–1961. N. England J. Med. 273:857.
14. ALEXANDER, A.D., H.G. STOENNER, G.E. WOOD & R.J. BYRNE. 1962. A new pathogenic
Leptospira, not readily cultivated. J. Bacteriol. 83:754.
15. BAKER, M.F. & H.J. BAKER. 1970. Pathogenic Leptospira in Malaysian surface waters I.
A method of survey for Leptospira in natural waters and soils. Amer. J. Trop. Med.
Hyg. 19:485.
16. DIESCH, S.L. & W.F. MCCULLOCH. 1966. Isolation of pathogenic leptospires from water
used for recreation. Pub. Health Rep. 81:299.
17. GILLESPIE, W.H., S.G. KENZY, L.M. RINGEN & F.K. BRACKEN. 1957. Studies on bovine
leptospirosis. III. Isolation of Leptospira pomona from surface water. Amer. J. Vet.
Res. 18:76.
18. BRAUN, J.L., S.L. DIESCH & W.F. MCCULLOCH. 1968. A method for isolating leptospires
from natural surface waters. Can. J. Microbiol. 14:1011.
19. FAINE, S. 1982. Guidelines for the control of leptospirosis. WHO offset publ. No. 67.
World Health Organization, Geneva, Switzerland.
20. SMIBERT, R.M. 1965. A technique for the isolation of leptospirae from contaminating
microorganisms. Can. J. Microbiol. 11:743.
21. ELLINGHAUSEN, H.C., JR. & W.G. MCCULLOUGH. 1965. Nutrition of Leptospira pomona
and growth of 13 other serotypes: a serum-free medium employing oleic albumin
complex. Amer. J. Vet. Res. 26:39.
22. ELLINGHAUSEN, H.C., JR. & W.G. MCCULLOUGH. 1965. Nutrition of Leptospira pomona
and growth of 13 other serotypes: fraction of oleic albumin complex and a medium of
bovine albumin and polysorbate 80. Amer. J. Vet. Res. 26:45.
23. TURNER, L.H. 1970. Leptospirosis III. Trans. Roy. Soc. Trop. Med. Hyg. 64:623.
24. ADLER, B., S. FAINE, W.L. CHRISTOPHER & R.J. CHAPPEL. 1986. Development of an
improved selective medium for isolation of leptospires from clinical material. Vet.
Microbiol. 12:377.
25. MYERS, D.M. & V.M. VARELA-DIAZ. 1973. Selective isolation of leptospiras from
contaminated material by incorporation of neomycin to culture media. Appl. Microbiol.
25:781.
26. JOHNSON, R.C. & P. ROGERS. 1964. 5-fluorouracil as a selective agent for growth of
leptospirae. J. Bacteriol. 87:422.
9260 J.
Legionella
The Legionellaceae have been implicated in outbreaks of disease occurring since 1947.1
Two forms of disease are recognized: a pneumonic form called Legionnaires’ Disease and a
nonpneumonic form called Pontiac fever. The first species was isolated following the historic
outbreak associated with the Legionnaires’ Convention in Philadelphia, Pa., in 1976.
Epidemiological findings and animal studies have shown that the organism is transmitted via the
airborne route2 and is ubiquitous in moist environments. The reservoirs for most outbreaks have
been either contaminated air conditioning cooling tower water or contaminated potable water
distribution systems.3,4 Legionella species also have been isolated in non-disease-related
circumstances from a wide variety of aquatic environments such as lakes, streams, reservoirs,
and sewage.5,6 The organisms are able to survive for prolonged periods in laboratory distilled
and tap water.7
The Legionellaceae are composed of a single genus, Legionella, and more than 35 different
species.8 The organisms are Gram-negative, aerobic, non-spore-forming bacteria. They are 0.5 to
0.7 µm wide and 2 to 20 µm long. They possess polar, subpolar, and/or lateral flagella. With the
exception of L. oakridgensis, all require cysteine and iron salts for growth.
Although Legionella originally were isolated in guinea pigs and embryonated hen’s eggs, it
has been shown that plating directly on artificial media is more sensitive than animal inoculation
for L. pneumophila.9 The most widely used medium is an ACES
(N-2-acetamideo-2-aminoethanesulfonic acid) buffered (pH 6.9) charcoal yeast extract (BCYE)
agar supplemented with cysteine, ferric pyrophosphate, and optimally, alpha-ketoglutarate
(BCYE-alpha).10
No one medium will be optimal for the recovery of Legionella from every environmental
site; thus different selective media with various antibiotic combinations in a BCYE base may be
necessary.10-12 Also, pretreating samples with hydrochloric acid-potassium chloride, pH 2.2, is
useful for eliminating non-Legionella organisms.13 The two most commonly used selective
media are GPVA medium (BCYE-alpha supplemented with glycine anisomycin, vancomycin,
and polymyxin B) and CCVC medium (BCYE-alpha supplemented with polymyxin B,
cephalothin, vancomycin, and cycloheximide). The GPVA medium is less inhibitory to some
Legionella species. Use CCVC medium in combination with a less selective medium.
Recovery of legionellae from environmental water samples sometimes is difficult.
Legionellae may take up to a week to grow on plate media, and even with acid pretreatment and
the addition of antibiotics to the medium, faster-growing organisms may overgrow legionellae.
In addition, other organisms, including Pseudomonas spp., secrete into surrounding media
bacterial products that can inhibit Legionella growth.14
Rapid methods for detecting Legionella utilizing direct fluorescent antibody staining (DFA)
or polymerase chain reaction technology (PCR) also are available and may be more sensitive
than culture-based assays.15-17 DFA can be quantitative, but may be subject to interference due
to cross-reactivity with other organisms.18 The PCR method is semiquantitative. Both DFA and
PCR may not distinguish between viable and nonviable bacteria.
Collect water samples from the littoral zone or from cooling towers, condenser coils, storage
tanks, showers, water taps, etc. In most instances, a 1-L water sample is sufficient. Larger
volumes of water (1 to 10 L)6 may be needed in water having low bacterial counts. In addition to
collecting water samples, it may be useful to swab various fixtures (e.g., shower heads) and plate
directly on selective media. Transport samples to the laboratory in insulated containers.
Refrigerate samples that cannot be processed immediately. Treat chlorinated water with sodium
thiosulfate (see Section 9060A.2).
2. Immunofluorescence Procedure
Centrifuge 100 mL at 3500 × g for 30 min at room temperature and reconstitute the
sedimented material in 6 to 10 mL filter-sterilized (0.2-mm filter) water from sample. Prepare
smears for DFA by filling two 1.5-cm circles on a microscope slide with the concentrate. Air-dry
sample smears, gently heat-fix, treat with 10% formalin for 10 min, rinse with
phosphate-buffered saline (pH 7.6), and react with specific fluorescent antibodies.6,19 The DFA
procedure lacks specificity20 and cannot determine viability. Some environmental bacteria (i.e.,
Pseudomonas spp. and Xanthomonas-Flavobacterium group) cross-react with the Legionella
DFA reagents.
To determine whether organisms are viable, use secondary staining with a tetrazolium dye.18
Confirm Legionella using direct isolation procedures.
3. Media and Reagents
a. Buffered charcoal yeast extract alpha base:19
Norit SG charcoal
Yeast extract
ACES buffer
Ferric pyrophosphate, soluble
L-cysteine, HCl⋅H2O
2.0
10.0
10.0
0.25
0.4
g
g
g
g
g
Agar
Potassium alpha-ketoglutarate
Reagent-grade water
17.0
1.0
1.0
g
g
L
Dissolve yeast extract, agar, charcoal, glycine, and alphaketoglutarate in approximately 850
mL water; boil. Dissolve 10g ACES buffer in 100mL warm water, adjust pH to 6.9 with 1N
KOH and add. Autoclave 15min at 121°C. Cool to 50°C. Dissolve 0.4g cysteine and 0.25g ferric
pyrophosphate in 10mL of water each and filter sterilize separately (0.22µm). After base has
cooled, add cysteine, ferric pyrophosphate, and dyes in that order. Adjust pH to 6.9 with sterile
1N KOH and dispense.
b. GPVA medium:11,12*#(72)
Glycine
Polymyxin B
0.3 %
100
units/mL
Vancomycin
5
µg/mL
Anisomycin
80
µg/mL
To cooled BCYE-alpha base with glycine, add filter-sterilized antibiotics and mix. Adjust
pH to 6.9 with sterile 1N KOH and dispense.
c. CCVC medium11†#(73)
Cephalothin
4
µg/ml
Colistin
Vancomycin
Cycloheximide
16
µg/mL
0.5 µg/mL
80
µg/mL
To cooled BCYE-alpha base add filter-sterilized antibiotics and mix. Adjust to pH 6.9 with
sterile 1N KOH and dispense.
d. Acid treatment reagent,11 pH 2.0 (0.2M KCl/HCl):
Solution A—0.2M KCl (14.9 g/L in distilled water).
Solution B—0.2M HCl (16.7 mL/L 10N HCl in distilled water).
Mix 18 parts of Solution A with 1 part of Solution B. Check pH against a pH 2.0 standard
buffer. Dispense into screw-cap tubes in 1.0-mL volumes and sterilize by autoclaving.
e. Alkaline neutralizer reagent11 (0.1N KOH):
Stock solution—0.1N KOH (6.46 g/L in deionized water). Dilute 10.7 mL of stock solution
with deionized water to 100 mL. Dispense into screw-cap tubes in convenient volumes and
sterilize by autoclaving. The pH of d and e combined in equal volumes should be 6.9.
4. Sample Preparation
a. Low-bacterial-count water: Concentrate water that has a low total bacterial count either by
filtration11 or continuous-flow centrifugation.21 Filter samples through sterile 47-mm filter
funnel assemblies containing a 0.2-µm porosity polycarbonate filter.‡#(74) After filtration,
immediately remove the filter aseptically and place it in a 50-mL centrifuge tube or similar-size
vessel containing 10 mL sterile tap water or phosphate buffer. If more than one filter is required
to concentrate a sample, combine them.
b. High-bacterial-count water: Process water that has a high total bacterial count directly.
Place 10 mL sample in a 50-mL centrifuge tube or similar-size vessel containing 10 mL of sterile
tap water or phosphate buffer.
c. Sample dispersion: Disperse organisms from filter or aggregates by mixing with a vortex
mixer (3 × 30 s).
d. Plating: Plate acid-treated and non-acid-treated samples on two types of BCYE: plain and
selective with antibiotics.
1) No acid treatment—Inoculate three plates each of BCYE-alpha and selective
BCYE-alpha (GPVA or CCVC) with 0.1 mL of suspension. Spread with a sterile smooth glass
rod. Save remainder of specimen for acid treatment and store at 4°C.
2) Acid treatment—Place 1.0 mL of suspension in a sterile 13 × 100-mm screw-capped tube
containing 1.0 mL acid treatment reagent and mix. Final pH of mixture should be approximately
2.2. Let stand for 15 min at room temperature, neutralize by adding 1.0 mL alkaline neutralizer
reagent, and mix. Inoculate 0.1 mL onto three plates each of BCYE-alpha and selective
BCYE-alpha (GPVA or CCVC) and spread with a sterile smooth glass rod.
3) Incubation—Incubate all plates at 35°C in a humidified atmosphere (>50%) for up to 10
d. A candle jar or humidified CO2 incubator (2 to 5% CO2) is acceptable.
e. Total bacterial count examination: Determine the adequacy of processing for each
high-bacterial-count water. Some samples may require dilution, concentration, or animal
inoculation. If the total count of the acid-treated sample exceeds 300 colonies on BCYE selective
medium, make a further 10-fold dilution of the sample stored at 4°C. Repeat acid-treatment and
plating.
If the total count of the non-acid-treated sample is less than 30 colonies on BCYE agar,
concentrate and treat the collected water as previously described for low-bacterial-count water.
5. Examination of Cultures of Legionellae
With the aid of a dissecting microscope, examine all cultures daily after 48 h incubation for
the presence of opaque bacterial colonies that have a ‘‘ground-glass’’ appearance. Place plates
with Legionella-like colonies in a biological safety cabinet equipped with a burner, a
bacteriological needle, and a loop. Aseptically pick each suspect colony onto BCYE-alpha agar
and a BCYE agar plate prepared without L-cysteine. Streak the inoculated portion of each plate
with a sterile loop to provide areas of heavy growth and incubate for 24 h.
Reincubate plates without growth an additional 24 h. Plates demonstrating growth on only
BCYE-alpha agar are presumptive for Legionella. Confirm Legionella by slide agglutination or
direct immunofluorescence. If these confirmatory techniques are not available, send subcultures
of the presumptive legionellae to a reference laboratory for further identification. Because there
are many serotypes in some species, especially L. pneumophila, investigation of environmental
sites as possible reservoirs of epidemic-causing strains may be useful.22 Effective investigatory
techniques include monoclonal antibody subtyping, electrophoretic isoenzyme analysis,
restriction endonuclease tests, and plasmid analysis.
6. Polymerase Chain Reaction Procedure
A test kit utilizing the polymerase chain reaction (PCR) is available commercially§#(75) and
has been used successfully in an epidemiological investigation of an outbreak of Pontiac fever.23
Perform tests according to manufacturer’s instructions. The kit provides sample processing
reagents, PCR primers, detection strips, and positive and negative controls. Specific probes allow
for the detection of twenty-five Legionella species as well as specific detection of Legionella
pneumophila. The test is semiquantitative, based on a colorimetric comparison to control strips
equivalent to 103 cells/mL.
7. References
1. MCDADE, J.E., C.C. SHEPARD, D.W. FRASIER, T.R. TSAI, M.A. REDUS, W.T. DOWDLE & THE
LABORATORY INVESTIGATION TEAM. 1977. Legionnaires’s Disease: isolation of a
bacterium and demonstration of its role in other respiratory disease. N. England J.
Med. 297:1197.
2. BERENDT, R.F., et al. 1980. Dose-response of guinea pigs experimentally infected with
aerosols of Legionella pneumophila. J. Infect. Dis. 141:186.
3. FLIERMANS, C.B., W.B. CHERRY, L.H. ORRISON, S.J. SMITH, D.L. TISON & D.H. POPE. 1981.
Ecological distribution of Legionella pneumophila. Appl. Environ. Microbiol. 41:9.
4. TOBIN, J.O.H., R.A. SWAN & C.L.R. BARTLETT. 1981. Isolation of Legionella
pneumophila from water systems: methods and preliminary results. Brit. Med. J.
282:515.
5. CHERRY, W.B., G.W. GORMAN, L.H. ORRISON, C.W. MOSS, A.G. STEIGERWALT, H.W.
WILKINSON, S.E. JOHNSON, R.M. MCKINNEY & D.J. BRENNER. 1982. Legionella
jordanis: a new species of Legionella isolated from water and sewage. J. Clin.
Microbiol. 15:290.
6. FLIERMANS, C.B., W.B. CHERRY, L.H. ORRISON & L. THACKER. 1979. Isolation of
Legionella pneumophila from nonepidemic related aquatic habitats. Appl. Environ.
Microbiol. 37:1239.
7. SKALIY, P. & H.V. MCEACHERN. 1979. Survival of the Legionnaires’s Disease bacterium
in water. Ann. Intern. Med. 90:662.
8. BRENNER, D.J., A.G. STEIGERWALT, G.W. GORMAN, H.W. WILKINSON, W.F. BIBB, M.
HACKEL, R.L. TYNDALL, J. CAMPBELL, J.C. FEELEY, W.L. THACKER, P. SKALIY, W.T.
MARTIN, B.J. BRAKE, B.S. FIELDS, H.W. MCEACHERN & L.K. CORCORAN. 1985. Ten new
species of Legionella. Int. J. System. Bacteriol. 35:50.
9. FEELEY, J.C., R.J. GIBSON, G.W. GORMAN, N.C. LANGFORD, J.K. RASHEED, D.C. MACEL &
W.B. BAINE. 1979. Charcoal-yeast extract agar: primary isolation medium for
Legionella pneumophila. J. Clin. Microbiol. 10:437.
10. EDELSTEIN, P.H. 1982. Comparative studies of selective media for isolation of
Legionella pneumophila from potable water. J. Clin. Microbiol. 16:697.
11. GORMAN, G.W., J.M. BARBAREE & J.C. FEELEY. 1983. Procedures for the Recovery of
Legionella from Water. Developmental Manual, Centers for Disease Control, Atlanta,
Ga.
12. WADOWSKY, R.M. & R.B. YEE. 1981. Glycine-containing selective medium for isolation
of Legionellaceae from environmental specimens. Appl. Environ. Microbiol. 42:768.
13. BOPP, C.A., J.W. SUMNER, G.K. MORRIS & J.G. WELLS. 1981. Isolation of Legionella spp.
from environmental water samples by low-pH treatment and use of selective medium.
J. Clin. Microbiol. 13:714.
14. PASZKO-KOLVA, C., P.A. HACKER, M.A. STEWART & R.L. WOLFE. 1993. Inhibitory effect
of heterotrophic bacteria on the cultivation of Legionella dumoffi. In J.M. Barbaree,
R.F. Breiman & A.P. Dufour, eds. Legionella: Current Status and Emerging
Perspectives. ASM Press, Washington, D.C.
15. PALMER, C.J., Y. TSAI, C. PASZKO-LOLVA, C. MAYER & L.R. SANGERMANO. 1993.
Detection of Legionella species in sewage and ocean water by polymerase chain
reaction, direct fluorescent-antibody, and plate culture methods. Appl. Environ.
Microbiol. 59:3618.
16. PALMER, C.J., G.F. BONILLA, B. ROLL, C. PASZKO-KOLVA, L.R. SANGERMANO & R.S.
FUJIOKA. 1995. Detection of Legionella species in reclaimed water and air with the
Enviroamp Legionella PCR kit and direct fluorescent antibody staining. Appl. Environ.
Microbiol. 61:407.
17. WILLIAMS, H.N., C. PASZKO-KOLVA, M. SHAHAMAT, C.J. PALMER, C. PETTIS & J. KELLEY.
1996. Molecular techniques reveal high prevalence of Legionella in dental units. J.
Amer. Dental Assoc. 127:1188.
18. FLIERMANS, C.B., R.J. SORACCO & D.H. POPE. 1981. Measure of Legionella pneumophila
activity in situ. Curr. Microbiol. 6:89.
19. JONES, G.L. & G.A. HEBERT. 1979. Legionnaires—the disease, the bacterium and
methodology. U.S. Dep. Health, Education, & Welfare, Centers for Disease Control,
Atlanta, Ga.
20. EDELSTEIN, P.H., R.M. MCKINNEY, R.D. MEYER, M.A.C. EDELSTEIN, C.J. KRAUSE & S.M.
FINEGOLD. 1980. Immunologic diagnosis of Legionnaires’ Disease: cross reactions
with anaerobic and microaerophilic organisms and infections caused by them. J. Infect.
Dis. 141:652.
21. VOSS, L., K.S. BUTTON, M.S. RHEINS & O.H. TUOVINEN. 1984. Sampling methodology for
enumeration of Legionella spp. in water distribution systems. In C. Thornsberry, A.
Balows, J.C. Feeley & W. Jakubowski, eds. Legionella, Proc. 2nd International
Symposium. American Soc. Microbiology, Washington, D.C.
22. BARBAREE, J.M., G.W. GORMAN, W.T. MARTIN, B.S. FIELDS & W.E. MORRILL. 1987.
Protocol for sampling environmental sites for Legionellae. Appl. Environ. Microbiol.
53:1454.
23. MILLER, L.A., J.I. BEEBE, J.C. BUTLER, W. MARTIN, R. BENSON, R.E. HOFFMAN & B.S.
FIELDS. 1993. Use of polymerase chain reaction in epidemiological investigations of
Pontiac fever. J. Infect. Dis. 168:769.
8. Bibliography
CENTERS FOR DISEASE CONTROL, NATIONAL INSTITUTE OF ALLERGY AND INFECTIOUS
DISEASES & WORLD HEALTH ORGANIZATION. 1979. International Symposium on
Legionnaire’s Disease. Ann. Intern. Med. 90:489.
BLACKMAN, J.A., F.W. CHANDLER, W.B. CHERRY, A.C. ENGLAND, J.C. FEELEY, M.D. HICKLIN, R.M.
MCKINNEY & H.W. WILKINSON. 1981. Legionellosis. Amer. J. Pathol. 103:427.
DUFOUR, A. & W. JAKUBOWSKI. 1982. Drinking water and Legionnaire’s Disease. J. Amer.
THORNSBERRY, C., A. BALOWS, J.C. FEELEY & W. JAKUBOWSKI, eds. 1984. Legionella, Proc. 2nd
International Symposium. American Soc. Microbiology, Washington, D.C.
9260 K.
Yersinia enterocolitica
Yersinia enterocolitica is a gram-negative bacterium that can cause acute gastroenteritis and
can be found in water in cold or temperate areas of the United States. Many wild, domestic, and
farm animals are reservoirs of this organism, including wild animals associated with water
habitats (beavers, minks, muskrats, nutrias, otters, and racoons).1,2 The organism can grow at
temperatures as low as 4°C with a generation time of 3.5 to 4.5 h if at least trace amounts of
organic nitrogen are present.3 Most environmental strains of Y. enterocolitica and the closely
related species, Y. kristensenii, Y. frederiksenii, and Y. intermedia, generally are considered
nonpathogenic, but disease outbreaks have been associated with environmental sources. Some
strains lacking classic virulence markers also may be associated with disease.4 Y. enterocolitica
has become recognized worldwide as an important human pathogen and in several countries it is
nearly as common as Salmonella and Campylobacter as a leading cause of acute or chronic
enteritis.5 Y. enterocolitica usually is associated with sporadic cases of gastroenteritis in the
U.S.; however, epidemiologic investigations suggest that the predominant pathogenic serotype
isolated in the U.S. has been changing.4 Y. enterocolitica serogroup O:3 has replaced O:8 as the
most common species recovered from patients, reflecting the same pattern seen in other parts of
the world.4,5 Two reported incidents of waterborne gastroenteritis possibly caused by Yersinia
occurred during the period 1971 to 1978.3,6,7
Yersinia has been isolated from untreated surface and ground waters in the Pacific
Northwest, New York, and other regions of North America, with highest isolations during the
colder months.8-10 Concentrations have ranged from 3 to 7900 CFU/100 mL. Laboratory tests
used to isolate and enumerate yersiniae do not discriminate between pathogenic and
nonpathogenic strains. Yersinia isolations do not correlate with levels of total and fecal coliforms
or total plate count bacteria.9 There is little information on Yersinia survival in natural waters
and water treatment processes.
In studies of Yersinia in chlorinated-dechlorinated secondary effluent and receiving (river)
water, the organism was isolated in 27% of the effluent samples, 9% of the upstream samples,
and 36% of the downstream samples.11 Mean total and fecal coliform reductions in effluent
chlorination were 99.93 and 99.95%, respectively. In a survey of untreated and treated
(chlorination or filtration plus chlorination) drinking water supplies, Yersinia was found in 14.0
and 5.7% of the samples, respectively.9 Of water samples with less than 2.2 coliforms/100 mL,
15.9% were Yersinia-positive. Yersinia isolation did not correlate with presence of total or fecal
coliforms in this study. Another study confirmed that E. coli also is not a good indicator for
Yersinia in water and that Y. enterocolitica O:3 strains harboring a virulence plasmid have
enhanced resistance to chlorine compared to non-virulent strains.12
Because of the existence of animal reservoirs, widespread occurrence and persistence of
Yersinia in natural and treated water in at least some geographic areas, the evidence for possible
waterborne outbreaks, and the lack of definitive information on its reduction by treatment
processes, this pathogen is of potential importance in drinking water.
1. Concentration and Cultivation
A membrane filter method for enumerating and isolating Yersinia enterocolitica is
available.13 The method may be used for examining large volumes of low-turbidity water and for
presumptively identifying the organism without transferring colonies to multiple confirmatory
media.
Filter sample through a membrane filter (see Section 9260B.1d). Place membrane filter on a
cellulose pad saturated with m-YE recovery broth. Incubate for 48 h at 25°C. Aseptically transfer
the membrane to a lysine-arginine agar substrate and incubate anaerobically at 35°C. After 1 h,
puncture a hole in the membrane next to each yellow to yellow-orange colony with a needle,
transfer the membrane to a urease-saturated absorbent pad, and incubate at 25°C for 5 to 10 min.
Immediately count all distinctly green or deep bluish-purple colonies. The green or bluish
colonies are sorbitol-positive, lysine- and arginine-negative, and urease-positive. They may be
presumptively identified as Y. enterocolitica or a closely related Yersinia species. Additional
biochemical testing is necessary to determine species. Reasonably simple tests have been
described to screen isolates for pathogenicity.14 Comprehensive biochemical and serological
characterization or the use of molecular methods is necessary to confirm virulence, but these
methods are not generally available.
2. References
1. WETZLER, T.F. & J. ALLARD. 1977. Yersinia enterocolitica from trapped animals in
Washington State. Paper presented at International Conf. Disease in Nature
Communicable to Man. Yellow Bay, Mont.
2. WETZLER, T.F., J.T. REA, G. YUEN & W. TURNBERG. 1978. Yersinia enterocolitica in
waters and wastewaters. Paper presented at 106th Annual Meeting, American Public
Health Assoc., Los Angeles, Calif.
3. HIGHSMITH, A.K., J.C. FEELEY, P. SKALIY, J.G. WELLS & B.T. WOOD. 1977. The isolation
and enumeration of Yersinia enterocolitica from well water and growth in distilled
4. BISSETT, M.J., C. POWERS, S.L. ABBOTT & J.M. JANDA. 1990. Epidemiologic
investigations of Yersinia enterocolitica and related species: sources, frequency, and
serogroup distribution. J. Clin. Microbiol. 28:910.
5. FENWICK, S.G. & M.D. MCCARTY. 1995. Yersinia enterocolitica is a common cause of
gastroenteritis in Auckland. N. Zealand Med. J. 108:269.
6. EDEN, K.V., M.L. ROSENBERG, M. STOOPLER, B.T. WOOD, A.K. HIGHSMITH, P. SKALIY,
J.G. WELLS & J.C. FEELEY. 1977. Waterborne gastrointestinal illness at a
ski-resort—isolation of Yersinia enterocolitica from drinking water. Pub. Health Rep.
92:245.
7. KEET, E. 1974. Yersinia enterocolitica septicemia. N.Y. State J. Med. 74:2226.
8. HARVEY, S., J.R. GREENWOOD, M.J. PICKETT & R.A. MAH. 1976. Recovery of Yersinia
enterocolitica from streams and lakes of California. Appl. Environ. Microbiol. 32:352.
9. WETZLER, T.F., J.R. REA, G.J. MA & M. GLASS. 1979. Non-association of Yersinia with
traditional coliform indicators. In Proc. Annu. Meeting American Water Works Assoc.,
10. SHAYEGANI, M., I. DEFORGE, D.M. MCGLYNN & T. ROOT. 1981. Characteristics of
Yersinia enterocolitica and related species isolated from human, animal, and
environmental sources. J. Clin. Microbiol. 14:304.
11. TURNBERG, W.L. 1980. Impact of Renton Treatment Plant effluent upon the
Green-Duwamish River. Masters Thesis, Univ. Washington, Seattle.
12. LUND, D. 1996. Evaluation of E. coli as an indicator for the presence of Campylobacter
jejuni and Yersinia enterocolitica in chlorinated and untreated oligotrophic lake water.
Water Res. 30:1528.
13. BARTLEY, T.D., T.J. QUAN, M.T. COLLINS & S.M. MORRISON. 1982. Membrane filter
technique for the isolation of Yersinia enterocolitica. Appl. Environ. Microbiol.
43:829.
14. FARMER, J.J., G.P. CARTER, V.L. MILLER, S. FALKOW & I.W. WACHSMUTH. 1992.
Pyrazinamidase, CR-MOX agar, salicin fermentation-esculin hydrolysis, and d-xylose
fermentation for identifying pathogenic serotypes of Yersinia enterocolitica. J. Clin.
Microbiol. 30:2589.
3. Bibliography
HIGHSMITH, A.K., J.C. FEELEY & G.K. MORRIS. 1977. Yersinia enterocolitica: a review of the
bacterium and recommended laboratory methodology. Health Lab. Sci. 14:253.
BOTTONE, E.J. 1977. Yersinia enterocolitica: a panoramic view of a charismatic microorganism.
CRC Crit. Rev. Microbiol. 5:211.
YANKO, W.A. 1993. Occurrence of Pathogens in Distribution and Marketing Municipal Sludges.
National Technical Information Serv. Rep. PB88-154273-AS, Springfield, Va.
9260 L.
Aeromonas (PROPOSED)
1. Introduction
Aeromonas spp. are natural inhabitants of aquatic environments worldwide. These
Gram-negative, facultatively anaerobic, glucose-fermenting organisms have been isolated from
groundwater, treated drinking water, surface waters, wastewater, sludge, and sediment. Their
populations are seasonal in all natural waters, with the highest numbers present in warmer
months. Aeromonads cause serious diseases of aquatic animals and represent an economic threat
to the aquaculture industry. The motile aeromonads have emerged as a serious microbial threat
to human populations, especially the immunocompromised.1
As a result of recent taxonomic studies, Aeromonas bacteria have been removed from the
family Vibrionaceae and established as the sole genus of the new family Aeromonadaceae. The
genus Aeromonas comprises 14 recognized and 2 proposed DNA hybridization groups with 13
named phenospecies and 4 unnamed genospecies. The extreme difficulty of phenotypically
differentiating aeromonads and the unavailability of DNA hybridization techniques in most
laboratories have lead clinical microbiologists to report aeromonads as A. hydrophila, A. sobria,
or A. caviae, according to a published classification scheme.2 Environmental microbiologists
usually combine all motile, mesophilic aeromonads into the Aeromonas hydrophila complex, or
simply report isolates as A. hydrophila. These practices obscure understanding of the medical
and public health significance of aeromonads isolated from clinical specimens, environmental
samples, and public water supplies; identification of Aeromonas isolates according to established
taxonomic principles is preferable.3
While no waterborne outbreaks of gastroenteritis attributed to aeromonads have implicated
public drinking water supplies in the U.S., this does not mean that none have occurred. The
epidemiologic association between ingestion of untreated well water and subsequent Aeromonas
gastrointestinal illness has been widely documented. Numerous cases and outbreak
investigations of water- and food-transmitted illnesses caused by aeromonads have been
reported.4 Outbreaks of gastroenteritis caused by aeromonads have occurred in custodial care
institutions, nursing homes, and day-care centers. Aeromonas contamination of drinking water
has been documented as a cause of travelers’ diarrhea.5
For many years, Aeromonas have been considered nuisance organisms by environmental
microbiologists because they were reported to interfere with coliform multiple tube fermentation
(MTF) methods. While aeromonads comprise 12% of bacteria isolated from drinking water by
presence-absence methods, no data have demonstrated inhibition of coliform organisms by
aeromonads in drinking water. Slight turbidity of LTB tubes, with or without a small bubble of
gas in the inverted tube, is suggestive of aeromonads. When the MTF method is used for
drinking water samples, cultures producing turbidity at 35°C that remain clear at 44.5°C are
suggestive of aeromonads. The presence of aeromonads can be verified by subculturing a loopful
of turbid broth to a MacConkey plate and screening colorless colonies for gelatinase and oxidase
production. No data are available to support invalidation of coliform MTF tests based on
turbidity of tubes in the absence of gas production.
The ecology of mesophilic aeromonads in aquatic environments, including water treatment
plants and distribution systems, has been reviewed.6 The Netherlands and the Province of
Quebec have established drinking water standards for Aeromonas at 20 CFU/100 mL for water
leaving the treatment plant, and 200 CFU/100 mL for distribution system water. Canada has
established an Aeromonas MCL of 0 (zero) for bottled water. A resuscitation method for
recovery of aeromonads in bottled water has been published.7
The ability to isolate, enumerate, and identify aeromonads from water and wastewater
sources is important because of their role in causing human and animal disease, their ability to
colonize treatment plants and distribution systems, and their presence and distribution as
alternative indicators of the trophic state of waters. The diversity of aeromonads in drinking
water plants and distribution systems was shown by several investigators.8-10
Many media and methods have been proposed for the isolation and enumeration of
aeromonads.11,12 The methods presented below represent a compromise, because no single
enrichment method, isolation medium, or enumeration method is capable of recovering all
aeromonads present in a water sample. The methods were chosen on the basis of reproducibility
of results, objectivity of interpretation, availability of materials, and specificity of the method for
detection of aeromonads in the presence of other heterotrophic bacteria. Consult the literature for
additional methods for use in special circumstances.13
Collect water samples in sterile screw-capped glass or plastic bottles or plastic bags.*#(76)
Sample volumes of 200 mL to 1 L are sufficient for most analyses. For chlorinated waters, add
sodium thiosulfate (see Section 9060A.2). The potentially toxic effect of heavy metals is
neutralized by adding EDTA (see Section 9060A.2).
Transport samples to the laboratory at 2 to 8°C within 8 h. Samples for presence-absence
analyses may be transported at ambient temperatures within 24 h. Grab samples are most
common. Moore swabs (see 9260B.1a) have been used for wastewater sampling, and Spira
bottles have been used for tapwater sampling.13 Both of these methods are used in conjunction
with enrichment in 1% alkaline peptone water (APW), pH 8.6.13 Place sediment and sludge
samples in bottles or bags and submit in same way as water samples.
3. Enrichment Methods
Do not use enrichment methods for ecological studies because the predominant strain(s) will
overgrow other organisms. Reserve enrichments for presence-absence tests for aeromonads in
drinking water, foods, stools, or for monitoring aeromonad populations in wastewater or marine
environments, where organisms may be present in low numbers or require resuscitation due to
injury from exposure to inimical agents or hostile physical environments. For isolation of
aeromonads from clear water samples, filter through 0.45 µm membrane filters, place filters in a
bottle with 10 mL APW, incubate overnight at 35°C, and inoculate to plating media for isolation.
Optimally, to sample clear water intended for drinking, filter a volume of water through a
mini-capsule filter†#(77), decant residual water from inlet, plug ends with sterile rubber
stoppers, and fill filter with APW, pH 8.6, through syringe port. Incubate filter at 35°C for 6 h or
overnight and streak loopfuls of broth onto selective and differential plating media.14
4. Enumeration Methods
a. Spread plates: Enumerate samples expected to contain predominantly aeromonads in high
numbers (sludge, sediments, wastewater effluents, polluted surface waters, etc.) directly by
spreading 0.1-mL portions of decimal dilutions on ampicillin dextrin agar (ADA)15-17 plates.
Incubate plates at 35°C overnight and count bright yellow colonies 1 to 1.5 mm in diameter.
Presumptively identify colonies using the screening methods below.
b. Membrane filtration (MF): Enumerate aeromonads in drinking water samples or other
low-turbidity waters by using MF procedures with ADA medium and incubating aerobically
overnight at 35°C. Filter sample volumes equivalent to 1 mL, 10 mL, and 100 mL. To achieve a
countable plate (1 to 30 colonies), prepare decimal dilutions when aeromonads are present in
high numbers. Count bright yellow colonies, 1 to 1.5 mm in diameter, and pick to screening
media.
c. Multiple-tube fermentation tests (MTF): Multiple-tube fermentation tests using APW, pH
8.6, or trypticase soy broth (TSB) containing ampicillin at 30 µg/mL (TSB30) have been applied
to foods; however, they have not been used for enumeration of aeromonads in water samples.
Some aeromonads are sensitive to ampicillin and will not grow in TSB30 medium. ADA without
agar has been used to enumerate aeromonads in drinking water.8 Use MTF methods only for
clean samples such as groundwater or treated drinking water samples, because the effect of
competing microflora present in surface waters on recovery of aeromonads in broth media has
not been studied adequately. Similarly, the correlation between MTF population estimates and
other enumeration methods has not been examined adequately for matrices other than foods.
5. Screening Tests
Pick 3 to 10 colonies resembling aeromonads on differential and selective plating media or
membrane filters and stab-inoculate into deeps of Kaper’s multi-test medium19 or one tube each
of triple sugar iron (TSI) agar and lysine iron agar (LIA). Incubate cultures at 30°C for 24 h.
Perform a spot oxidase test on growth taken from the LIA slant. Do not test for oxidase on
growth from TSI slants, MacConkey agar, or other selective or differential media, because acid
production interferes with the oxidase reaction. Reactions of enteric bacteria on TSI and LIA
media are shown in Table 9260:III. When Kaper’s medium is used instead of TSI/LIA slants,
colonies may be picked and inoculated onto sheep blood agar plates; incubate at 35°C overnight
to provide growth for the oxidase test and to record hemolysin production. Cultures are
identified presumptively using Kaper’s medium according to the characteristics shown in Table
9260:IV. If species identification is desirable, submit presumptively identified Aeromonas
cultures to a reference laboratory. Cultures with potential public health or regulatory significance
may be subtyped using various molecular methods to determine clonality for outbreak
investigations and trouble-shooting of treatment plant or distribution system problems.1
6. References
1. JANDA, J.M. & S.L. ABBOTT. 1996. Human Pathogens. In B. Austin, M. Altwegg, P.
Gosling & S.W. Joseph, eds. The Genus Aermonas, p. 151. John Wiley & Sons,
Chichester, U.K.
2. POPOFF, M. & M. VERON. 1976. A taxonomic study of the Aeromonas
hydrophila-Aeromonas punctata group. J. Gen. Microbiol. 94:11.
3. CARNAHAN, A.M. & M. ALTWEGG. 1996. Taxonomy. In B. Austin, M. Altwegg, P.
Gosling & S.W. Joseph, eds. The Genus Aeromonas, p. 1. John Wiley & Sons,
Chichester, U.K.
4. JOSEPH, S.W. 1996. Aeromonas gastrointestinal disease: a case study in causation?. In
B. Austin, M. Altwegg, P. Gosling & S.W. Joseph, eds. The Genus Aeromonas, p. 311.
John Wiley & Sons, Chichester, U.K.
5. HANNINEN, M.L., S. SALMI, L. MATTILA, R. TAIPALINEN & A. SIITONEN. 1995.
Association of Aeromonas spp. with travellers’ diarrhoea in Finland. J. Med.
Microbiol. 42:26.
6. HOLMES, P., L.M. NICCOLLS & D.P. SARTORY. 1996. The ecology of mesophilic
Aeromonas in the aquatic environment. In B. Austin, M. Altwegg, P. Gosling & S.W.
Joseph, eds. The Genus Aeromonas, p. 127. John Wiley & Sons, Chichester, U.K.
7. WARBURTON, D.W., J.K. MCCORMICK & B. BOWEN. 1993. Survival and recovery of
Aeromonas hydrophila in water: development of methodology for testing bottled water
in Canada. Can. J. Microbiol. 40:145.
8. HANNINEN, M.-L. & A. SIITONEN. 1995. Distribution of Aeromonas phenospecies and
genospecies among strains isolated from water, foods or from human clinical samples.
Epidemiol. Infect. 115:39.
9. HUYS, G., I. KERSTERS, M. VANCANNEYT, R. COOPMAN, P. JANSSEN & K. KERSTERS.
1995. Diversity of Aeromonas sp. in Flemish drinking water production plants as
determined by gas-liquid chromatographic analysis of cellular fatty acid methyl esters
(FAMEs). J. Appl. Bacteriol. 78:445.
10. MOYER, N.P., G.M. LUCCINI, L.A. HOLCOMB, N.H. HALL & M. ALTWEGG. 1992.
Application of ribotyping for differentiating aeromonads isolated from clinical and
environmental sources. Appl. Environ. Microbiol. 58:1940.
11. GAVRIEL, A. & A.J. LAMB. 1995. Assessment of media used for selective isolation of
Aeromonas spp. Lett. Appl. Microbiol. 21:313.
12. JEPPESEN, C. 1995. Media for Aeromonas spp., Plesiomonas shigelloides and
Pseudomonas spp. from food and environment. Int. J. Food Microbiol. 26:25.
13. MOYER, N.P. 1996. Isolation and enumeration of aeromonads. In B. Austin, M.
Altwegg, P. Gosling & S.W. Joseph, eds. The Genus Aeromonas, p. 39. John Wiley &
Sons, Chichester, U.K.
14. MOYER, N.P., G. MARTINETTE, J. LUTHY-HOTTENSTEIN & M. ALTWEGG. 1992. Value of
rRNA gene restriction patterns of Aeromonas spp. for epidemiological investigations.
Curr. Microbiol. 24:15.
15. HANDFIELD, M., P. SIMARD & R. LETARTE. 1996. Differential media for quantitative
16.
17.
18.
19.
recovery of waterborne Aeromonas hydrophila. Appl. Environ. Microbiol. 62:3544.
HAVELAAR, A.H., M. DURING & J.F. VERSTEEGH. 1987. Ampicillin-dextrin agar medium
for the enumeration of Aeromonas species in water by membrane filtration. J. Appl.
Bacteriol. 62:279.
HAVELAAR, A.H. & M. VONK. 1988. The preparation of ampicillin dextrin agar for the
enumeration of Aeromonas in water. Lett. Appl. Microbiol. 7:169.
ALTWEGG, M. 1996. Subtyping methods for Aeromonas species. In B. Austin, M.
Altwegg, P. Gosling & S.W. Joseph, eds. The Genus Aeromonas, p. 109. John Wiley &
Sons, Chichester, U.K.
KAPER, J., R.J. SEIDLER, H. LOCKMAN & R.R. COLWELL. 1979. Medium for the
presumptive identification of Aeromonas hydrophila and Enterobacteriaceae. Appl.
9260 M.
Mycobacterium (PROPOSED)
The genus Mycobacterium comprises over 70 characterized species that are non-motile
spore-forming, aerobic, acid-fast bacilli measuring 0.2 to 0.6 × 1 to 10 µm. Most organisms in
this genus are saprophytes, but some species are capable of causing disease in humans. The
primary pathogens in this group include Mycobacterium tuberculosis and Mycobacterium leprae,
the causative agents of tuberculosis and leprosy, respectively. Recently there has been an
increase in the incidence of disease caused by nontuberculosis mycobacteria, probably related to
the increasing numbers of immunocompromised patients.1-3 In the genus Mycobacterium, the
most important opportunistic pathogens include M. avium-intracellulare, M. kansasii, M.
marinum, and M. simiae, which are capable of causing disease when the immune system is
compromised. Some of the common hosts and environmental reservoirs of Mycobacteria are
shown in Table 9260:V.
Because of the complex nature of the cell wall, which is rich in lipids and therefore has a
hydrophobic surface, this genus is resistant to many common disinfectants. As a result, several
members of this genus are becoming important waterborne pathogens in the
immunocompromised population. Mycobacteria also are acid-fast and extremely slow-growing.
Some species such as M. avium-intracellulare require from 3 to 8 weeks to form colonies on
culture media.
Mycobacterium avium and Mycobacterium intracellulare exhibit overlapping properties,
making speciation extremely difficult. As a result, these two pathogens are grouped together and
called M. avium-intracellulare or refered to as the MAC complex. Organisms from this group
are ubiquitous in the environment and have been isolated from potable water systems, including
those in hospitals4-6 as well as from soil and dairy products. This pathogen causes a chronic
pulmonary disease in immunocompetent hosts that is clinically and pathologically
indistinguishable from tuberculosis; it also causes disseminated disease in immunocompromised
hosts. The primary route of transmission is believed to be through ingestion, but increasing
numbers of cases originate in the respiratory tract, indicating an aerosol route of transmission.
1. Sample Collection and Concentration
Mycobacteria typically constitute a minority of the microflora, especially in finished waters,
and require sample concentration. Collect water samples in sterile 1-L polypropylene containers.
For finished, disinfected waters, add 1 mL 10% sodium thiosulfate solution/L water collected.
Transport samples to laboratory immediately after collection. If samples cannot be analyzed
immediately, store at 4°C and begin analysis within 24 h of sampling.
2. Screening Water Samples by Direct Fluorescent Assay
Before committing the sample to a lengthy culture incubation, survey for acid-fast bacteria
by using a combination solution of Auramine-Rhodamine (A-R) fluorescent dye.7*#(78)
Auramine and Rhodamine nonspecifically bind to mycolic acids and resist decolorization by acid
alcohol.8
Filter a minimum of 500 mL finished water or 100 mL source water (depending on turbidity),
through a sterile 0.45-µm-porosity, 47-mm-diam black filter. Aseptically transfer filter to a
sterile polypropylene 50-mL tube and add 5 mL of buffered dilution water. Resuspend organisms
from filter by vortexing for 2 min. Aspirate suspension and aseptically transfer to a sterile 15-mL
polypropylene centrifuge tube. Centrifuge suspension at 5000 g for 10 min and discard all but
about 0.5 mL of supernatant. Resuspend pellet by vortexing. Transfer 100 mL of the concentrate
to a clean glass slide and air-dry and heat-fix at 60 to 70°C for 2 h or overnight. Primary stain the
smear with A-R (15 min), decolorize with acid-alcohol†#(79) for 2 to 3 min, rinse with
deionized water, apply secondary potassium permanganate counterstain (no longer than 2 to 4
min), rinse, and let air-dry. Examine smear at 100 × and 400 × with a microscope fitted with a
BG-12 or 5113 primary filter with a OG-1 barrier filter. Acid-fast organisms will stain
yellow-orange on a black background. To confirm for acid-fastness, apply a traditional acid-fast
stain (Ziehl-Nielsen with Kenyon modification) directly to the prepared smear following the A-R
stain.
For wastewater or highly turbid source waters, collect a 10-mL subsample and transfer to a
sterile polypropylene 15-mL tube. Centrifuge at 5000 g for 10 min and discard all but about 0.5
mL of supernatant. Follow slide preparation procedure and staining as above.
3. Decontamination and Culture Methods
Mycobacteria grow very slowly on laboratory media. Therefore, eliminate from the sample
naturally occurring organisms that can out-compete and overgrow the mycobacteria. Various
isolation and identification methods have been described for the recovery of mycobacteria,
especially in the hospital environment.4-6 Decontamination of the sample concentrate is required
for the selection for mycobacteria before culture. In addition, the matrix may affect the success
of the recovery of mycobacteria. Several methods (a through c below) are detailed for recovering
mycobacteria from water samples; determine which method performs best with the matrix to be
examined.
a. Filter 500-mL water sample through a sterile 0.45-µm-porosity, 47-mm-diam filter.
Aseptically transfer filter to a sterile polypropylene 50-mL tube. Add 5 mL sterile distilled water
and resuspend organisms off the filter by shaking with two 5-mm glass beads for 1 h on a
mechanical shaker.9 Add a 3% sodium lauryl sulfate, 1% NaOH solution.10 Spread portions of
this suspension onto a selective agar medium as described in ¶ 4 below.
b. Filter 500-mL water sample through sterile 0.45-µm-porosity, 47-mm-diam filter.
Aseptically transfer filter to a sterile polypropylene 50-mL tube. Add 5 mL sterile distilled water
and resuspend organisms off the filter by shaking with glass beads for 5 min on a mechanical
shaker. Add 10 mL 1M NaOH for 20 min followed by centrifugation at 8600 g at 4°C for 15
min. Discard supernatant and add 5 mL 5% oxalic acid for 20 min. Re-centrifuge, discard
supernatant, and add 30 mL sterile distilled water to neutralize. Centrifuge again, and resuspend
in 0.7 mL distilled water.11 Use portions of this material for selective growth (¶ 4 below).
c. Add 20 mL 0.04% (w/v) cetylpridinium chloride (CPC) to 500-mL water sample and leave
at room temperature for approximately 24 h. Filter sample and wash filter with 500 mL sterile
water.12 A study of decontamination methods for the isolation of mycobacteria from drinking
water samples found a CPC concentration of 0.005% (w/v) to yield the highest isolation rate and
lowest contamination rate for the water examined.13
4. Selective Growth
Culture all samples in duplicate. After sample decontamination, either spread portions of the
concentrates or use sterile forceps to place filters on selective media. One common egg-based
medium that successfully isolates mycobacteria from environmental concentrates is
Lowenstein-Jensen agar. An agar-based medium containing cycloheximide (7H10) is a general
growth medium for mycobacteria as well. Place plates in humid chambers or gas-permeable bags
to prevent dehydration, and incubate at 37°C. Additional plates also can be incubated at 30°C in
a humidified chamber to detect mycobacteria that grow optimally at lower temperatures.
Examine plates periodically during a 3- to 8-week incubation period. Count suspect colonies
(acid-fast coccobacilli) and subculture to a tube of 7H9 broth. After 5 d, remove subsamples and
stain with Ziehl-Nielsen stain with Kenyon modification. Subculture coccobacillary acid-fast
organisms further onto 7H10 plates. Conduct phenotypic tests (Table 9260:VI) as a first step
toward identification. If biochemical tests do not allow speciation, use other methods, such as
fatty acid profile by HPLC or GLC, serological typing, and/or molecular tests such as DNA
probes, and RFLP, which have been used for rapid detection of a limited number of species.14
Although phenotypic tests have been the standard for species identification, there are several
inherent problems in this approach. First, because initial identification of mycobacteria can take
3 to 8 weeks, observing biochemical changes entails additional time for the isolates (especially
those of nontuberculosis mycobacteria) to metabolize specific substrates or to exhibit certain
characteristics. Second, phenotypic traits are not stable; thus some species of mycobacteria are
untypable by conventional methods.
One approach that can successfully speciate Mycobacterium is sequencing amplified
rDNA.15 The method produces objective results in 2 d, gives reproducible results (due to the
stability of the 16S rRNA), and can identify new species. These techniques have been used in
clinical diagnostic laboratories, and now are available in some full-service environmental testing
laboratories.
5. References
1. GOOD, R.C. Opportunistic pathogens in the genus Mycobacterium. 1985. Annu. Rev.
Microbiol. 39:347.
2. CARSON, L.A., L.A. BLAND, L.B. CUSICK, M.S. FAVERO, G.A. BOLAN, A.L. REINGOLD &
R.C. GOOD. 1988. Prevalence of nontuberculous mycobacteria in water samples of
hemodialysis centers. Appl. Environ. Microbiol. 54:3122.
3. DUMOULIN, G.C. & K.D. STOTTMEIR. 1986. Waterborne mycobacteria: an increasing
threat to health. ASM News 52: 525.
4. DUMOULIN, G.C., K.D. STOTTMEIR, P.A. PELLETIER, A.Y. TSANG & J. HEDLEY-WHITE.
1988. Concentration of Mycobacterium avium by hospital water systems. J. Amer.
Med. Assoc. 260:1599.
5. POWELL, B.L. & J.E. STEADHAM. 1981. Improved technique for isolation of
Mycobacterium kansasii from water. J. Clin. Microbiol. 13:969.
6. CARSON, L.A., L.B. CUSICK, L.A. BLAND & M.S. FAVERO. 1988. Efficiency of chemical
dosing methods for isolating nontuberculous mycobacteria from water supplies of
dialysis centers. Appl. Environ. Microbiol. 54:1756.
7. NOLTE, F.S. & B. METCHOCK. 1995. Mycobacterium. In P.R. Murray, E.J. Baron, M.A.
Pfaller, F.C. Tenover & R.H. Yolken, eds. Manual of Clinical Microbiology. American
Soc. Microbiology Press, Washington, D.C.
8. CHAPIN, K. 1995. Clinical Miscroscopy. In P.R. Murray, E.J. Baron, M.A. Pfaller, F.C.
Tenover & R.H. Volken, eds. Manual of Clinical Microbiology. American Soc.
Microbiology Press, Washington, D.C.
9. KAMALA, T., C.N. PARAMASIVAN, D. HERBERT, P. VENKATESAN & R. PRABHAKAR.
1994. Evaluation of procedures for isolation of nontuberculous mycobacteria from soil
and water. Appl. Environ. Microbiol. 60:1021.
10. ENGEL, H.W.B., L.G. BERWALD & A.H. HAVELAAR. 1980. The occurrence of
Mycobacterium kansasii in tapwater. Tubercle 61:21.
11. IIVANANINEN, E.K., P.J. MARTIKAINEN, P.K. VAANANEN & M.-L.KATILA. 1993.
Environmental factors affecting the occurrence of mycobacteria in brook waters. Appl.
12. DUMOULIN, G.C. & K.D. STOTTMEIR. 1978. Use of cetylpyridinium chloride in the
decontamination of water culture of mycobacteria. Appl. Environ. Microbiol. 36:771.
13. SCHULZE-ROBBECKE, R., A. WEBER & R. FISCHEDER. 1991. Comparison of
decontamination methods for the isolation of mycobacteria from drinking water
samples. J. Microbiol. Methods 14:177.
14. ANDREW, P.W. & G.J. BOULNOIS. 1990. Early days in the use of DNA probes for
Mycobacterium tuberculosis and Mycobacterium avium complexes. In A.J.L. Macario
& E.C. de Macario, eds. Gene Probes for Bacteria. Academic Press, San Diego, Calif.
15. ROGALL, T., T. FLOHR & E.C. BOTTGER. 1990. Differentiation of Mycobacterium species
by direct sequencing of amplified DNA. J. Gen. Microbiol. 136:1915.
6. Bibliography
TSUKAMURA, M. 1981. A review of the methods of identification and differentiation of
mycobacteria. Rev. Infect. Dis. 3:841.
GOOD, R.C. 1985. Opportunistic pathogens in the genus Mycobacterium. Annu. Rev. Microbiol.
39:347.
ICHIYAMA, S. & K. SHIMOKATA. 1988. The isolation of Mycobacterium avium complex from
soil, water, and dusts. Microbiol. Immunol. 32:733.
BROADLEY, S.J., P.A. JENKINS, J.R. FURR & A.D. RUSSELL. 1991. Anti-mycobacterial activity of
biocides. Lett. Appl. Microbiol. 13:118.
FISCHEDER, R., R. SCHULZE-ROBBECKE & A. WEBER. 1991. Occurrence of mycobacteria in
drinking water samples. Zentralbl. Hyg. Umweltmed. 192:154.
JENKINS, P. A. 1991. Mycobacteria in the environment. J. Appl. Bacteriol. 70:137.
SCHULZE-ROBBECKE, R., B. JANNING & R. FISCHEDER. 1992. Occurrence of mycobacteria in
biofilm samples. Tubercle Lung Dis. 73:141.
COLLINS, J. & M. YATES. 1994. Mycobacteria in water. J. Appl. Bacteriol. 1984. 57:193.
JENSEN, P.A. 1997. Airborne Mycobacterium spp. In C.J. Hurst, G.R. Knudsen, M.J. McInerney,
L.D. Stetzenbach & M.V. Walter, eds. Manual of Environmental Microbiology. American
Soc. Microbiology Press, Washington, D.C.
9510
DETECTION OF ENTERIC VIRUSES*#(80)
9510 A.
Introduction
1. Occurrence
Viruses excreted with feces or urine from any species of animal may pollute water.
Especially numerous, and of particular importance to health, are the viruses that infect the
gastrointestinal tract of man and are excreted with the feces of infected individuals. These
viruses are transmitted most frequently from person to person by the fecal-oral route. However,
they also are present in domestic sewage which, after various degrees of treatment, is discharged
to either surface waters or the land. Consequently, enteric viruses may be present in
sewage-contaminated surface and ground waters that are used as sources of drinking water. The
viruses known to be excreted in relatively large numbers with feces include polioviruses,
coxsackieviruses, echoviruses, and other enteroviruses, adenoviruses, reoviruses, rotaviruses, the
hepatitis A (infectious hepatitis) virus(es), and the Norwalk-type agents that can cause acute
infectious nonbacterial gastroenteritis. With the possible exception of hepatitis A, each group or
subgroup consists of a number of different serological types; thus more than 100 different human
enteric viruses are recognized.1-4
In temperate climates enteroviruses occur at peak levels in sewage during the late summer
and early fall. However, hepatitis A virus (HAV), Norwalk-type viruses, and rotaviruses may be
important exceptions because the incidence of the diseases due to these viruses increases in the
colder months. Quantitative information on seasonal patterns of occurrence in water and
wastewater of these latter viruses is lacking because they cannot be assayed readily with
conventional cell culture techniques. The Norwalk-type viruses have not been cultivated in any
cell cultures, although immunochemical assay methods have been developed to detect them as
antigens.5,6 Human rotaviruses and HAV have been cultivated recently in cell cultures, but the
techniques are difficult and require concomitant use of immunoassays such as
immunofluorescence to detect virus growth or gene probes.7-11
Viruses are not normal flora in the intestinal tract; they are excreted only by infected
individuals, mostly infants and young children. Infection rates vary considerably from area to
area, depending on sanitary and socioeconomic conditions. Viruses usually are excreted in
numbers several orders of magnitude lower than those of coliform bacteria. Because enteric
viruses multiply only within living, susceptible cells, their numbers cannot increase in sewage.
Sewage treatment, dilution, natural inactivation, and water treatment further reduce viral
numbers. Thus, although large outbreaks of waterborne viral disease may occur when massive
sewage contamination of a water supply takes place,12 waterborne transmission of viral infection
and disease in technologically advanced nations depends on whether minimal quantities of
viruses are capable of producing infections. It has been demonstrated that infection can be
produced experimentally by a very few virus units,13 although the risk of infection increases
with increasing ingested doses.14 The risk of infection incurred by an individual in a community
with a water supply containing a very few virus units has not been determined. Risk analysis has
suggested that significant risk of infection could result from low numbers of enteric viruses
present in a drinking water supply.15 The percentage of individuals who develop clinical illness
may be as low as 1% for poliovirus and as great as 97% for hepatitis A.15
Most recognized waterborne virus disease outbreaks in the U.S. have been caused by obvious
sewage contamination of untreated or inadequately treated private and semipublic water
supplies. Virus disease outbreaks in community water supply systems usually are caused by
contamination through the distribution system.16
2. Testing for Viruses
The routine examination of water and wastewater for enteric viruses is not recommended
now. However, in special circumstances such as wastewater reclamation, disease outbreaks, or
special research studies, it may be prudent or essential to conduct virus testing. Such testing
should be done only by competent and specially trained water virologists having adequate
facilities.
Laboratories planning to concentrate viruses from water and wastewater should do so with
the clear understanding that the available methodology has important limitations.17 Even the
most current methods for concentrating viruses from water still are being researched and
continue to be modified and improved. The efficiency of a virus concentration method may vary
widely depending on water quality. Furthermore, none of the available virus detection methods
have been tested adequately with representatives from all of the virus groups of public health
importance. Most virus concentration methods have achieved adequate virus recoveries with
water or wastewater samples that have been contaminated experimentally with known quantities
of a few specific enteric viruses. Although method effectiveness in field trials is difficult to
evaluate, some virus concentration methods have been used successfully to recover naturally
occurring enteric viruses. Some of these methods require large equipment for sample processing
and virus assay and identification procedures usually require cell culture and related virology
laboratory facilities.
Detecting viruses in water through recovery of infectious virus requires three general steps:
(a) collecting a representative sample, (b) concentrating the viruses in the sample, and (c)
identifying and estimating quantities of the concentrated viruses. Particular problems associated
with the detection of viruses of public health interest in the aquatic environment are: (a) the
small size of virus particles (about 20 to 100 nm in diameter), (b) the low virus concentrations in
water and the variability in amounts and types that may be present, (c) the inherent instability of
viruses as biological entities, (d) the various dissolved and suspended materials in water and
wastewater that interfere with virus detection procedures, and (e) the present limitations of virus
estimation and identification methods.
3. Selection of Concentration Method
The densities of enteric viruses in water and wastewater usually are so low that virus
concentration is necessary, except possibly for raw sewage in certain areas or seasons.18
Numerous methods for concentrating waterborne enteric viruses have been proposed, tested
under laboratory conditions with experimentally contaminated samples, and in some cases used
to detect viruses under field conditions.19,20
Virus concentration methods often are capable of processing only limited volumes of water
of a given quality. In selecting a virus concentration method consider the probable virus density,
the volume limitations of the concentration method for that type of water, and the presence of
interfering constituents. A sample volume less than 1 L and possibly as small as a few milliliters
may suffice for recovery of viruses from raw or primary treated sewage. For drinking water and
other relatively nonpolluted waters, the virus levels are likely to be so low that hundreds or
perhaps thousands of liters must be sampled to increase the probability of virus detection.
Three different techniques used to concentrate viruses from water are described herein:
adsorption to and elution from microporous filters (Methods B and C); aluminum hydroxide
adsorption-precipitation (Method D); and polyethylene glycol (PEG) hydroextraction-dialysis
(Method E).19,20 A separate technique (Method F) for recovering viruses from solids in small
volumes of water also is described. Virus concentration by adsorption to and elution from
microporous filters can be used for both small volumes of wastewater and large volumes of
natural and finished waters. The aluminum hydroxide adsorption-precipitation and PEG
hydroextraction-dialysis methods are impractical for processing large fluid volumes. However,
they are suitable for concentrating viruses from wastewater or other waters having relatively
high virus densities and for second-step concentration (reconcentration) of viruses in primary
eluates obtained by processing large sample volumes through microporous filters.
4. Recovery Efficiencies
In examining a particular water include a preliminary evaluation of virus recovery
efficiency. To do this add a known quantity of one or more test virus types to the required
volume of sample, process the sample by the concentration method, and assay the concentrate
for test viruses to determine virus recovery efficiency. Ideally, such seeded samples should be
used whenever field samples are processed. If seeded samples are used concurrently with field
samples, take appropriate steps, including disinfection and sterilization and the use of aseptic
technique, to prevent accidental contamination of samples.
5. References
1. RAO, V.C. & J.M. MELNICK. 1986. Environmental Virology. American Soc.
2. FEACHEM, R.G., D.J. BRADLEY, H. GARELICK & D.D. MARA. 1983. Sanitation and
Disease. Health Aspects of Excreta and Wastewater Management. John Wiley & Sons,
New York, N.Y.
3. WILLIAMS, F.P. & E.W. AKIN. 1986. Waterborne gastroenteritis. J. Amer. Water Works
Assoc. 78:34.
4. FEACHEM, R., H. GARELICK & J. SLADE. 1981. Enteroviruses in the environment. Trop.
Dis. Bull. 78:185.
5. BLACKLOW, N.R. & G. CUKOR. 1980. Viral gastroenteritis agents, Chap. 90 in E.H.
Lennette, A. Balows, W.J. Hausler, Jr. & J.P. Truant, eds. Manual of Clinical
Microbiology, 3rd ed. American Soc. Microbiology, Washington, D.C.
6. KAPIKIAN, A.Z., R.H. YOLKEN, H.B. GREENBERG, R.G. WYATT, A.R. KALICA, R.M.
CHANOCK & H.W. KIM. 1979. Gastroenteritis viruses. In E.H. Lennette & N.J. Schmidt,
eds. Diagnostic Procedures for Viral, Rickettsial and Chlamydial Infections. American
Public Health Assoc., Washington, D.C.
7. SOBSEY, M.D., S.E. OGLESBEE, D.A. WAIT & A.I. CUENEA. 1984. Detection of hepatitis A
in drinking water. Water Sci. Technol. 17: 23.
8. SMITH, E.M. & C.P. GERBA. 1984. Development of a method for detection of human
rotavirus in water. Appl. Environ. Microbiol. 43: 1440.
9. SATO, K., Y. INABA, T. SHINOZAKI, R. FUJII & M. MATUMOTO. 1981. Isolation of human
rotavirus in cell cultures. Arch. Virol. 69:155.
10. HEJKAL, T.W., E.M. SMITH & C.P. GERBA. 1984. Seasonal occurrence of rotavirus in
sewage. Appl. Environ. Microbiol. 47:588.
11. JIANG, X., M.K. ESTES & T.G. METCALF. 1987. Detection of hepatitis A virus by
hybridization with single-stranded RNA probes. Appl. Environ. Microbiol. 53:2487.
12. MELNICK, J.L. 1957. A water-borne urban epidemic of hepatitis. In Hepatitis Frontiers.
Little, Brown & Co., Boston, Mass.
13. WARD, R.L., D.I. BERNSTEIN & E.C. YOUNG. 1986. Human rotavirus studies in
volunteers: Determination of infectious dose and serological response to infection. J.
Infect. Dis. 154:871.
14. AKIN, E. 1981. A review of infective dose data for enteroviruses and other enteric
microorganisms in human subjects. In Microbial Health Considerations of Soil
Disposal of Domestic Wastewaters. EPA-600/9-83-017, U.S. Environmental Protection
Agency, Washington, D.C.
15. GERBA, C.P. & C.N. HAAS. 1988. Assessment of risks associated with enteric viruses in
contaminated drinking water. In J.J. Lichtenberg, J.A. Winter, C.I. Weber & L.
Frankin, eds. Chemical and Biological Characterization of Sludges, Sediments, Dredge
Spoils, and Drilling Muds. ASTM STP 976. American Soc. Testing & Materials,
Philadelphia, Pa.
16. CRAUN, G.F. 1986. Waterborne Disease in the United States. CRC Press, Boca Raton,
Fla.
17. SOBSEY, M.D. 1982. Quality of currently available methodology for monitoring viruses
in the environment. Environ. Internat. 7:39.
18. BURAS, N. 1976. Concentration of enteric viruses in wastewater and effluent: A two
year survey. Water Res. 10:295.
19. SOBSEY, M.D. 1976. Methods for detecting enteric viruses in water and wastewater. In
G. Berg, H.L. Bodily, E.H. Lennette, J.L. Melnick & T.G. Metcalf, eds. Viruses in
Water. American Public Health Assoc., Washington, D.C.
20. GERBA, C.P. & S.M. GOYAL. 1982. Methods in Environmental Virology. Marcel Dekker,
New York.
9510 B.
Virus Concentration from Small Sample Volumes by Adsorption to
and Elution from Microporous Filters
Viruses can be concentrated from aqueous samples by reversibly adsorbing them to
microporous filters and then eluting them from the filters in a small liquid volume.1 The
virus-containing sample is pressure-filtered through microporous filters having large surface
areas to which viruses adsorb, presumably by both electrostatic and hydrophobic interactions.2
Two general types of adsorbent filters are available: electronegative (negative surface charge)
and electropositive (positive surface charge). The former filters are composed of either cellulose
esters or fiberglass with organic resin binders. They adsorb viruses most efficiently in the
presence of multivalent cations such as Al3+ and Mg2+ and/or at low pH, usually pH 3.5. The
latter filters are composed of either fiberglass or cellulose and a positively charged organic,
polymeric resin. They adsorb viruses efficiently over a wide pH range without added polyvalent
salts. If the sample is neutral or acidic, it can be processed with these filters without chemical
conditioning.
Electropositive filters have given virus recoveries comparable to those with electronegative
filters.3-5 They have been used in field studies,6,7 and were evaluated with a variety of virus
types8-13 and waters.
Adsorbed viruses usually are eluted from the surfaces of microporous filters by
pressure-filtering a small volume of eluent fluid through the filters in situ. The eluent is either a
slightly alkaline proteinaceous fluid such as beef extract or a more alkaline buffer such as
glycine-NaOH, pH 10.5 to 11.5. If glycine-NaOH is used as eluent, preferably use pH 10.5
because of the greater likelihood of virus inactivation at the higher pH.14,15
Microporous filter methods suffer from three main limitations. Sample suspended matter
tends to clog the adsorbent filter, thereby limiting the volume that can be processed and possibly
interfering with the elution process.16 Dissolved and colloidal organic matter in some waters can
interfere with virus adsorption to filters, presumably by competing with viruses for adsorption
sites,17-19 and they also can interfere with virus elution. Finally, viruses adsorbed to suspended
matter may be removed in any clarification procedure applied before virus adsorption. These
solids-associated viruses are lost from the sample unless special efforts are made to recover the
solids and process them for viruses.16 A method for recovering solids-associated viruses from
small volumes of water and wastewater is given in Section 9510F. Despite these limitations,
virus concentration by adsorption to and elution from microporous filters is a most promising
technique for detecting viruses.
2. Equipment and Apparatus
a. Adsorbent filter holder, 47-, 90-, or 142-mm diam, equipped with pressure relief valve.
b. Pressure vessel, 12- or 20-L capacity.
c. Positive pressure source up to about 400 kPa with regulator: laboratory air line, air pump,
or cylinder of compressed air or nitrogen gas.
d. Autoclavable vinyl plastic tubing with plastic or metal connectors (quick-disconnect type),
for connecting positive pressure source, pressure vessel, and filter holder in series.
e. pH meter.
f. Beakers, 50- to 500-mL.
g. Laboratory balance.
h. Graduated cylinders, 25- to 100-mL.
i. Pipets, 1-, 5-, and 10-mL.
3. Materials
a. Electronegative virus adsorbent filter: Use either:
1) Cellulose nitrate filter, 0.45-µm porosity.*#(81)
2) Fiberglass-acrylic resin filter, 0.45-µm porosity.†#(82) Filter media available
commercially only as flat sheets can be cut to the desired disk diameter with scissors.
b. Electropositive virus adsorbent filter: Use either:
1) Surface modified cellulose and filter aid disk depth-filter.‡#(83)
2) Surface modified cellulose and filter aid thin-sheet medium, 0.20-µm porosity.§#(84)
c. Prefilter: Use one or more cellulose nitrate or fiberglass-acrylic resin filters or equivalent,
with porosities greater than 0.45 µm to prevent clogging of the virus adsorbent filter by
suspended matter. Place prefilters on top of the 0.45-µm-porosity virus adsorbent filter in the
same filter holder.
4. Reagents
a. Hydrochloric acid, HCl, 0.1, 1.0, and 10N.
b. Sodium hydroxide, NaOH, 0.1, 1.0, and 10N.
c. Aluminum chloride, AlCl3⋅6H2O, 0.15N, or magnesium chloride, MgCl2⋅6H2O, 5N
(necessary only for electronegative filters).
d. Sodium thiosulfate, Na2S2O3⋅5H2O, 0.5% (w/v).
e. Sodium chloride, 0.14N, pH 3.5: Dissolve 8.18 g in 1 L reagent-grade water and adjust to
pH 3.5 with HCl (necessary only for electronegative filters).
f. Virus eluent: Use either:
1) Glycine-NaOH, pH 10.5 or 11.5: Prepare 0.05M glycine solution, autoclave, and adjust to
pH 10.5 or 11.5 with 1 to 10N NaOH. Add phenol red, 0.0005%, as a pH indicator.
2) Beef extract, 3%, pH 9.0: Dissolve 30 g beef extract paste or 24 g beef extract powder in
1000 mL reagent-grade water, adjust to pH 9.0 with 1 to 10N NaOH, and sterilize by
autoclaving.
g. Glycine-HCl, pH 1.5: Prepare 0.05M glycine solution, autoclave, and adjust to pH 1.5 with
1 to 10N HCl. Add phenol red, 0.0005%, as a pH indicator.
h. Nutrient broth, 10X, pH 7.5: Dissolve 8.0 g nutrient broth in 90 mL reagent-grade water,
adjust to pH 7.5, dilute to 100 mL with reagent-grade water, and sterilize by autoclaving.
i. Antibiotics: Use either:
1) Penicillin-streptomycin, 10X: Contains 5000 IU penicillin/mL and 5000 µg
streptomycin/mL. Use commercially available form or prepare by dissolving powdered sodium
or potassium penicillin-G and streptomycin sulfate in reagent-grade water and sterilizing by
filtration. Store frozen.
2) Gentamycin-kanamycin, 100X: Contains 5000 µg/mL each of gentamycin (base) and
kanamycin (base). Prepare by combining aseptically equal volumes of commercially available
sterile gentamycin and kanamycin solutions, 10 000 µg/mL, respectively, or by dissolving
powdered gentamycin sulfate and kanamycin sulfate in reagent-grade water and sterilizing by
filtration. Store refrigerated or frozen.
j. Hanks balanced salt solution, 10X: Use commercially available form or prepare following
a standard protocol.20
k. Sodium hypochlorite, 5.25% available chlorine (household bleach).
5. Procedure
a. Sterilization of apparatus, materials, and reagents: Most reagents, virus adsorbent filters,
filter holders, tubing, and labware can be sterilized by autoclaving or made virus-free by
streaming steam. To sterilize filters load into their holders; if several filters are to be placed in
one holder, place filter with smallest porosity on the bottom with progressively larger filters on
top. Do not use an automatic drying cycle when autoclaving virus adsorbent filters. Sterilize
apparatus and material that cannot be autoclaved or treated with streaming steam by treating with
10-mg/L free chlorine solution, pH 7.0, for 30 min and rinse or flush with 50-mg/ L sterile
Na2S2O3 solution. Do not treat adsorbent filters with chlorine. Use aseptic technique during all
virus concentration operations to prevent extraneous microbial contamination.
b. Sample size and choice of filter size: Sample size and, hence, filter diameter depend partly
on water quality and the probable virus concentration. Single-stage microporous filter
adsorption-elution methods have been used to recover viruses from 100 mL raw sewage on
47-mm-diam filters21 and from 3.8 to 4.6 L secondary and tertiary sewage effluent on 90- or
142-mm-diam filters.18,21,22 Based on the diameter and solids-holding characteristics of the
filters, the scale and volume capacity of the apparatus and materials, and the quality of the
samples, the practical limits for sample size are 20, 8, and 2 L for 142-, 90-, and 47-mm-diam
filters, respectively.
c. Choice of filter type: Virus adsorption to electropositive filters decreases above pH 8 and
pH adjustment below this value may be necessary for optimal virus adsorption.4 Virus recovery
from raw sewage may be less than with electronegative filters.10
d. Sample collection and storage: Collect samples aseptically in sterile containers. If they
contain residual chlorine, immediately add Na2S2O3 solution to give a final concentration of 50
mg/L. Process samples as soon as possible after collection; do not hold samples for more than 2
h at up to 25°C or 48 h at 2 to 10°C. Do not freeze samples unless they cannot be processed
within 48 h; then freeze and store at −70°C or less.
e. Sample processing of electronegative filters: Adjust sample to pH 3.5 and 0.0015N AlCl3
or to between pH 6.0 and 3.5 and 0.1N MgCl2. Make sample adjustments either in a pressure
vessel or in another appropriate container. Mix sample vigorously during addition of 1.0 or 0.1N
HCl and AlCl3 solution (1 part solution to 100 parts sample) or MgCl2 solution (1 part solution
to 50 parts sample). Because AlCl3 is an acid salt, it may decrease sample pH slightly. Do not let
sample pH fall below 3.0.
Place sample in a pressure vessel connected to a source of positive pressure and connect
pressure vessel outlet to inlet of virus adsorbent filter holder. With pressure relief valve on filter
holder opened, apply a slight positive pressure to purge air from filter holder. When sample just
begins to flow from pressure relief valve, quickly close valve and continue filtration at a rate not
exceeding 28 mL/min/cm2 of filter area (about 130, 250, and 4000 mL/min for 47-, 90-, and
142-mm-diam filters, respectively). After filtering entire sample let positive pressure source
purge excess fluid from filter holder.
Wash filters with 0.14N NaCl to remove excess Al3+ or Mg2+ from virus adsorbent filter.
Use about 1.5 mL NaCl solution/cm2 filter area (25, 100, and 240 mL for 47-, 90-, and
142-mm-diam filters, respectively). Place wash solution in a pressure vessel connected to filter
holder inlet, use positive pressure to filter solution through virus adsorbent filter, discard filtrate,
and let positive pressure purge virus adsorbent filter of excess wash solution.
Elute viruses from filters with a recommended eluent. Use about 0.45 mL eluent/cm2 filter
surface area (about 7.5, 28, and 71 mL for 47-, 90-, and 142-mm-diam filters, respectively). With
pressure relief valve on filter holder open, add eluent to filter holder so that it completely covers
filter surface. When eluent begins to discharge from pressure relief valve, quickly close valve. If
pH 11.5 glycine-NaOH is the eluent, place a sterile beaker under filter outlet and apply positive
pressure so that filtrate flows slowly from filter holder outlet. Collect filtrate in sterile beaker
and, when filtrate no longer flows, slowly increase pressure to force retained fluid from filters.
Quickly check eluate (filtrate) pH. If it is less than 11.0, elute with additional pH 11.5
glycine-NaOH until an eluate with a pH ≥ 11.0 is obtained. Immediately after checking pH,
adjust eluate to a pH between 9.5 and 7.5 with pH 1.5 glycine-HCl or 0.1N HCl while mixing
vigorously. Complete elution and eluate pH adjustment to 7.5 to 9.5 in 5 min or less to avoid the
possibility of appreciable virus inactivation.
If pH 10.5 glycine-NaOH is the eluent, proceed as with pH 11.5 glycine-NaOH, but pass the
eluate through the filters a total of five times. For each elution, collect the filtrate, readjust to pH
10.5 with 1.0 or 0.1N NaOH, and then pass through the filter. After the fifth elution, adjust
filtrate to pH 7.4 with glycine-HCl, pH 1.5, or 0.1N HCl.
If 3% beef extract, pH 9.0, is the eluent, place a sterile beaker under filter outlet, apply a
slight positive pressure to eluent-containing filter holder so that filtrate flows slowly from the
outlet, and collect filtrate. Slowly increase pressure to force additional retained fluid from filters.
Measure eluate volume and add 1/10 of the measured volume each of penicillin-streptomycin
or gentamycin-kanamycin, Hanks balanced salt solution, and 10X nutrient broth (add last item to
glycine eluates only). Adjust sample to pH 7.4 with glycine-HCl or 0.1N HCl while mixing
vigorously. Store at either 4 or −70°C, depending on the time until virus assay. Maximum
storage at 4°C is 48 h.
f. Processing of electropositive filters: Processing for electropositive filters is identical to
that for electronegative filters except that addition of Al3+ and Mg2+ and sample pH adjustments
are unnecessary; because Al3+ and Mg2+ are not added, it is not necessary to wash filters with
0.14N NaOH before elution. If sample pH is greater than 8.0, adjust to less than pH 8 by adding
1.0 or 0.1N HCl.
6. References
1. FARRAH, S.R., C.P. GERBA, C. WALLIS & J.L. MELNICK. 1976. Concentration of viruses
from large volumes of tapwater using pleated membrane filters. Appl. Environ.
Microbiol. 31:221.
2. FARRAH, S.R., D.O. SHAH & L.O. INGRAM. 1981. Effects of chaotropic and antichaotropic
agents on the elution of poliovirus adsorbed to membrane filters. Proc. Nat. Acad. Sci.
U.S. 18:1229.
3. SOBSEY, M.D. & B.L. JONES. 1979. Concentration of poliovirus from tap water using
positively charged microporous filters. Appl. Environ. Microbiol. 37:588.
4. SOBSEY, M.D. & J.S. GLASS. 1980. Poliovirus concentration from tap water with
electropositive adsorbent filters. Appl. Environ. Microbiol. 40:201.
5. SOBSEY, M.D., R.S. MOORE & J.S. GLASS. 1981. Evaluating adsorbent filter performance
for enteric virus concentrations in tap water. J. Amer. Water Works Assoc. 73:542.
6. CHANG, L.T., S.R. FARRAH & G. BITTON. 1981. Positively charged filters for virus
recovery from wastewater treatment plant effluents. Appl. Environ. Microbiol. 42:921.
7. HEJKAL, T.W., B. KESWICK, R.L. LABELLE, C.P. GERBA, Y. SANCHEZ, G. DREESMAN, B.
HAFKIN & J.L. MELNICK. 1982. Viruses in a community water supply associated with an
outbreak of gastroenteritis and infectious hepatitis. J. Amer. Water Works Assoc.
74:318.
8. SCHLAAK, M., E. TISCHER & J.M. LOPEZ. 1983. Evaluation of current procedures for the
concentration of viruses in water. Zentralbl. Bakteriol. Microbiol. Hyg. I. Abt. Orig. B
177:127.
9. GUTTMAN-BASS, N. & R. ARMON. 1983. Concentration of Simian rotavirus SA-11 from
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
tap water by membrane filtration and organic flocculation. Appl. Environ. Microbiol.
45:850.
ROSE, J.B., S.N. SINGH, C.P. GERBA & L.M. KELLEY. 1984. Comparison of microporous
filters for concentration of viruses from wastewater. Appl. Environ. Microbiol. 45:989.
RAPHAEL, R.A., S.A. SATTAR & V.S. SPRINGTHROPE. 1985. Rotavirus concentration from
raw water using positively charged filters. J. Virol. Methods 11:131.
NUPEN, E.M. & B.W. BATEMAN. 1985. The recovery of viruses from drinking water by
means of an in-line electropositive filter. Water Sci. Technol. 17:63.
TORANZOS, G.A. & C.P. GERBA. 1989. An improved method for the concentration of
rotaviruses from large volumes of water. J. Virol. Methods 24:131.
SOBSEY, M.D., J.S. GLASS, R.J. CARRICK, R.R. JACOBS & W.A. RUTALA. 1980. Evaluation
of the tentative standard method for enteric virus concentration from large volumes of
tap water. J. Amer. Water Works Assoc. 72:292.
SOBSEY, M.D., J.S. GLASS, R.R. JACOBS & W.A. RUTALA. 1980. Modification of the
tentative standard method for improved virus recovery efficiency. J. Amer. Water
WELLINGS, F.M., A.L. LEWIS & C.W. MOUNTAIN. 1976. Demonstration of
solids-associated virus in wastewater and sludge. Appl. Environ. Microbiol. 31:354.
FARRAH, S.R., S.M. GOYAL, C.P. GERBA, C. WALLIS & P.T.B. SHAFFER. 1976.
Characteristics of humic acid and organic compounds concentrated from tapwater
using the aquella virus concentrator. Water Res. 10:897.
WALLIS, C. & J.L. MELNICK. 1967. Concentration of viruses from sewage by adsorption
on Millipore membranes. Bull. World Health Org. 36:219.
SOBSEY, M.D., C. WALLIS, M. HENDERSON & J.L. MELNICK. 1973. Concentration of
enteroviruses from large volumes of water. Appl. Microbiol. 26:529.
SCHMIDT, N.J. 1979. Tissue culture technics for diagnostic virology. In E.H. Lennette &
N.J. Schmidt, eds. Diagnostic Procedures for Viral and Rickettsial Infections, 5th ed.
RAO, V.C., U. CHANDORKAR, N.U. RAO, P. KUMARAN & S.B. LAKHE. 1972. A simple
method for concentrating and detecting viruses in wastewater. Water Res. 6:1565.
GERBA, C.P., S.R. FARRAH, S.M. GOYAL, C. WALLIS & J.L. MELNICK. 1978. Concentration
of enteroviruses from large volumes of tap water, treated sewage, and seawater. Appl.
9510 C.
Virus Concentration from Large Sample Volumes by Adsorption to
and Elution from Microporous Filters
This section describes a two-stage process for concentrating viruses from large sample
volumes. Viruses in eluate volumes too large to be conveniently and economically assayed
directly in cell cultures, such as those obtained from processing large volumes of water through
cartridge or large disk filters, can be concentrated further (reconcentrated) by several alternative
methods. Viruses in proteinaceous eluates can be reconcentrated by either ‘‘organic
flocculation,’’1,2 aluminum hydroxide adsorption-precipitation (Section 9510D), or polyethylene
glycol hydroextraction-dialysis (Section 9510E). These reconcentration techniques can be used
for both proteinaceous and organic buffer eluates from all types of water. Organic flocculation,
now used widely, involves precipitating viruses by acidifying eluates to pH 3.5, recovering the
precipitate by centrifugation, and then resuspending it in a small volume of alkaline buffer.1
Additionally, viruses in nonproteinaceous eluates such as glycine-NaOH can be
reconcentrated by adsorption to and elution from small microporous filters. The eluate is
adjusted to pH and ionic conditions for optimum virus adsorption, filtered through a secondary
adsorbent, and adsorbed viruses are eluted with a small volume of eluent. This procedure can be
used only for reconcentrating primary eluates obtained from processing drinking water and other
highly finished waters because of potential interfering substances likely to be present in primary
eluates from natural and less finished waters.
Figure 9510:1 shows the alternative microporous filter adsorption-elution and
reconcentration methods.
For general information on microporous filter techniques, see Section 9510B.1.
a. Apparatus for first-stage concentration (Figure 9510:2):
1) First-stage virus adsorbent filter holder.
2) Chemical additive system. Use either:
a) Fluid proportioner with four feed pumps (quadraplex) and a mixing chamber.*#(85)
b) Venturi-type proportioning injector†#(86) with plastic or metal connectors
(quick-disconnect type) and a length of vinyl tubing for the chemical feed line.3 To feed two
separate additives, attach a ‘‘Y’’ or ‘‘T’’ connector and two lengths of vinyl tubing to the
chemical feed port, or alternatively, use two separate proportioning injectors. It may be
necessary to use a bypass system with the injector to prevent loss of chemical feed due to back
pressure from the water line.4 This bypass system consists of ‘‘T’’ pipe fittings on the injector
inlet and outlet ports connected by a length of flexible hose with an in-line shut-off/control valve
(see Figure 9510:2).
Proportioning injectors available commercially will process water at flow rates of 3 to 33
L/min with water-to-chemical feed ratios between 10 to 1 and 1110 to 1. Select equipment and
operating conditions providing a water-to-chemical feed ratio of 100 to 1.
3) Water flow meter.
4) Pressure gauge, 0 to 400 kPa.
5) Vinyl plastic tubing, autoclavable, with plastic or metal connectors (quick-disconnect
type).
6) Pressure relief valve (optional).
7) Carboys, 20- to 50-L, or similar containers.
8) Positive pressure source up to 400 kPa with regulator: laboratory air line, positive
pressure pump, or cylinder of compressed air or nitrogen gas.
9) Pump (if source water is not under pressure).
b. pH meter.
c. Laboratory balance.
d. Beakers, 2- or 4-L.
e. Pressure vessel, 4-L.
f. Graduated cylinders, 1- and 2-L.
g. Pipets, 1-, 5-, and 10-mL.
h. Centrifuge with rotor and buckets for 250- to 500-mL-capacity bottles.‡#(87)
i. Centrifuge bottles, 250- to 500-mL.
3. Materials
a. First-stage electronegative virus adsorbent filters: Use one of the following:
1) 293-mm-diam, 8.0- and 1.2-µm-porosity cellulose nitrate filter series.§#(88)
2) 17.8-cm-long, 8.0-µm-porosity fiberglass-epoxy filter tube.i#(89)
3) 25.4-cm-long, 0.25- or 0.45-µm-porosity fiberglass-acrylic resin pleated filter
cartridge.##(90)
b. Second-stage electronegative virus adsorbent filters: 47-mm-diam, 3.0-, 0.45-, and
0.25-µm-porosity fiberglass-acrylic resin filter series. Use to reconcentrate highly finished water
samples only.##(91)
c. First-stage electropositive adsorbent filters: Use one of the following:
1) 293-mm-diam surface modified cellulose and filter aid filters.**#(92)
2) 25-cm-long, 0.20-µm-porosity surface modified thin-sheet media pleated filter
cartridge.††#(93)
4. Reagents
a. Hydrochloric acid, HCl, 0.06, 1,‡‡#(94) and 6N.
b. Sodium hydroxide, NaOH, 10N.
c. Aluminum chloride, AlCl3⋅6H2O, 0.15 and 6N.‡‡
d. Magnesium chloride, MgCl2⋅6H2O, 10N.‡‡
e. Sodium thiosulfate, Na2S2O3⋅5H2O, 0.5% (w/v).
f. Sodium hypochlorite, 5.25% available chlorine (household bleach).
g. Eluent: Use either:
1) Glycine-NaOH, pH 10.5 or 11.5: See Section 9510B.4 f1). Use within 2 h of pH
adjustment.
2) Beef extract, 3%, pH 9.0.§§#(95) See Section 9510B.4 f2).
h. Eluate neutralizing solution: Use either:
1) Glycine-HCl, pH 1.5: Prepare 0.05M glycine solution and adjust to pH 1.5 with 6N HCl.
Add phenol red, 0.0005%, as a pH indicator. Use within 2 h of pH adjustment.
2) HCl, 1.0N.
i. Nutrient broth, 10X, pH 7.5: Dissolve 8.0 g nutrient broth in 90 mL distilled water, adjust
to pH 7.5 with 10N NaOH, dilute to 100 mL with distilled water, and sterilize by autoclaving.
j. Disodium phosphate, 0.45N: Dissolve 40.2 g Na2HPO4⋅7H2O in 1 L distilled water and
sterilize by autoclaving.
k. Antibiotics: See Section 9510B.4i.
l. Sodium chloride, 0.14N: Dissolve 8.18 g NaCl in 1 L distilled water (necessary only with
electronegative filters).
m. Hanks balanced salt solution, 10X: See Section 9510B.4 j.
5. Procedure
When using electronegative filters, follow ¶s a–f below for production of primary eluate.
When using electropositive filters, first see ¶ g for procedural modifications.
a. Sterilization of apparatus, materials, and reagents: See Section 9510B.5a.
b. Sample size: For drinking water use a minimum sample of 400 L, although 2000 L or more
may have to be processed to detect viruses at a concentration of 1 to 2 infectious units/400 L.
c. Preparation of feed solutions for electronegative filters: Use an HCl additive solution to
adjust sample pH to 3.5 for virus adsorption to filters. If acidification to pH 3.5 is inadequate for
obtaining maximum virus adsorption, add either AlCl3 or MgCl2 solution.
When only HCl is used, prepare additive solution as follows: Determine concentration of
HCl additive solution by titrating a 1-L sample of dechlorinated water to pH 3.5 with 0.06N HCl
and noting volume required. The volume, in milliliters, of titrant required is equal to the volume
of 6N HCl needed/L distilled water for making the additive solution. Make at least 5 L additive
solution for 400 L of sample.
When AlCl3 is used to enhance virus adsorption use pH 3.5 and a final concentration of
added AlCl3 of 0.0015N. Because AlCl3 is an acid salt, titrate a 1-L sample to about pH 4.0 with
0.06N HCl, add AlCl3 to a concentration of 0.0015N and continue titration to pH 3.5, noting
volume of titrant used. Prepare additive solution by adding titrant volume (mL) of 6.0N HCl/L of
0.15N AlCl3.
When MgCl2 is used to enhance virus adsorption, use a pH between 3.5 and 6.0 and a final
concentration of added MgCl2 of 0.1N. To prepare additive solution titrate a 1-L sample to
desired pH with 0.06N HCl as previously described and note volume of titrant used. Add the
titrant volume of 6.0N HCl/L 10N MgCl2 to make the additive solution.
d. Preparation of chemical additive system:
1) When using a fluid proportioner, operate at a pressure of 100 to 700 kPa and a water flow
rate of 4 to 40 L/min. Adjust each of the four chemical additive pumps of the proportioner for a
ratio of 1 to 200 (1 part chemical additive to 200 parts water). Use two pumps, operating
reciprocally, for each additive so that the overall dilution for each additive is 1 to 100. One
additive is either HCl, HCl-AlCl3 or HCl-MgCl2; the other additive, 0.5% Na2S2O3, is needed
only when processing samples containing chlorine. Place lines from the two pumps of each
additive solution into the additive containers and manually operate the pump metering rods to fill
feed lines and purge them of air. Connect fluid proportioner to source water and operate briefly
without a virus adsorbent in place. Sample conditioned water from proportioner outlet and check
pH. The pH should be 3.5 ± 0.3.
When using a Venturi-type proportioning injector, connect injector assembly to water source
and to adsorbent filter inlet, and place additive feed line(s) into additive container(s). Position
valve on injector outlet to drain line position (away from adsorbent filter). Begin flow of sample.
Adjust screw-operated control valve on chemical feed of proportioning injector until water
collected from drain line is at the desired pH as measured with a pH meter. If Na2S2O3 is used to
neutralize chlorine, check to insure that chlorine is absent. Connect virus concentrator assembly
to source water by attaching concentrator inlet hose to valved outlet of a pressurized water
source or to outlet of a water pump, the inlet of which has been placed in the source water.
Operate for several minutes without a virus adsorbent in place to purge the unit of chlorine
solution. Collect a sample from outlet of meter to insure absence of chlorine.
e. First-stage concentration: After preparing concentration apparatus and additive solutions
and checking conditioned water for proper pH and absence of chlorine, attach a virus adsorbent
filter to outlet of chemical additive system. Attach water meter and effluent hose to virus
adsorbent outlet. Record initial meter reading and add to this value the desired volume to be
processed plus an additional 1 or 2% (to account for volume of either 1 or 2 additive solutions,
respectively). This gives meter reading at which sampling is to be stopped. Turn on water and
start a timer (or record starting time). Shortly after filtration begins collect a sample from filter
outlet and check for absence of chlorine and for appropriate pH value. Also check flow rate. Do
not use a flow rate above 40 L/min. Recheck pH and chlorine residual several times during
sample processing, or monitor continuously. When desired volume has been processed, turn
water off. Purge filter holder of excess water with positive pressure from an air or nitrogen gas
source.
f. Washing and virus elution: If AlCl3 or MgCl2 has been used, wash excess Al3+ or Mg2+
from filter with 4 L 0.14N NaCl. Omit washing if only HCl was used. Place wash solution in a
4-L pressure vessel and pass through filter with positive pressure. Purge filter of excess wash
solution with positive pressure and discard entire filtrate.
Using aseptic technique, elute virus from filter as soon as possible in the field or after
returning to the laboratory. If filter holders with adsorbed viruses must be returned to the
laboratory, seal filter holder openings, place filter holder in a sterile plastic bag, and chill.
Use pH 10.5 or 11.5 glycine-NaOH or 3% beef extract, pH 9.0, to elute viruses from
first-stage adsorbent filters. Because some viruses are inactivated when pH 11.5 glycine-NaOH
is used, alternatively elute with pH 10.5 glycine-NaOH or 3% beef extract, pH 9.0.1,3,4
To elute, place eluent in a pressure vessel. Use minimum eluent volumes of 1 L and 300 mL
for cartridge and 293-mm-diam disk filters, respectively. To elute with pH 11.5 glycine-NaOH,
connect pressure vessel to inlet of filter holder and with pressure relief valve on filter holder
open, apply a small positive pressure to the system so that eluent fills void volume of filter
holder. When eluent begins to discharge from pressure relief valve, quickly close it. Filter
remaining eluent slowly through filter within 1 to 2 min and collect filtrate (eluate) in a sterile 2or 4-L beaker. When filtrate no longer appears, slowly increase pressure to force additional fluid
from filter. If using pH 11.5 glycine-NaOH eluent, immediately check filtrate pH and if it is less
than 11.0, elute with 1 L more of pH 11.5 glycine-NaOH. Immediately after checking pH, adjust
filtrate to a pH between 7.5 and 9.5 with pH 1.5 glycine-HCl while mixing vigorously. Complete
elution and pH adjustment to 7.5 to 9.5 in 5 min or less to avoid possibility of appreciable virus
inactivation.
To elute with pH 10.5 glycine-NaOH, use either batch or continuous-flow eluent
recirculation. For the batch method, begin elution as with pH 11.5 glycine-NaOH. Collect the
filtrate, measure pH, and readjust to pH 10.5 with 1.0 or 0.1N NaOH while mixing vigorously.
Then, using this eluate, elute filters four more times, readjusting filtrate to pH 10.5 before each
elution. After the fifth elution, adjust filtrate to pH 7.4 with pH 1.5 glycine-HCl or 1.0N HCl
while mixing vigorously.
Alternatively, elute with pH 10.5 glycine-NaOH by continuous recirculation. Place eluent in
a sterile beaker. Attach short lengths of sterile vinyl or rubber tubing to inlet and outlet openings
of filter holder and place free ends of tubing in eluent beaker; slip midsection of filter inlet
tubing into a peristaltic or roller pump. Open pressure relief valve on filter holder and operate
pump at slow speed so that eluent fills void volume of filter holder. When eluent begins to
discharge from pressure relief valve, quickly close it. Increase pump speed so that eluent
recirculates through filter assembly and beaker at a minimum flow rate of 100 mL/min. After 5
min recirculation, remove filter inlet tube from beaker and pump remaining fluid from filter
assembly. Connect filter inlet to positive pressure source to force additional eluent from filter.
Adjust eluate to pH 7.4 with pH 1.5 glycine-HCl or 1.0N HCl while mixing vigorously.
To elute with 3% beef extract, pH 9.0, follow the procedure described above for pH 11.5
glycine-NaOH. Adjust collected filtrate to pH 7.4 with pH 1.5 glycine-HCl or 1N HCl while
mixing vigorously. The 5 min time limit to complete elution with pH 11.5 glycine-NaOH is not
necessary when beef extract is used.
g. Sample processing of electropositive filters: Processing for electropositive filters is
identical to that for electronegative filters except that addition of Al3+ and Mg2+ and sample pH
adjustments are unnecessary; because Al3+ and Mg2+ are not added, it is not necessary to wash
filters with 0.14N NaOH before elution. If the sample pH is greater than 8.0, adjust to less than
pH 8 by adding 1.0 or 0.1N HCl.
h. Reconcentration of primary eluates: Further concentrate (reconcentrate) viruses in primary
eluates either by organic flocculation, Al(OH)3 adsorption-precipitation (Section 9510D),
polyethylene glycol hydroextraction-dialysis (Section 9510E), or adsorption to and elution from
microporous filters. The latter technique can be used only for glycine or other organic buffer
eluates.
To concentrate (reconcentrate) viruses in glycine eluates by adsorption to and elution from
filters, adjust to pH 3.5 with pH 1.5 glycine-HCl and add AlCl3 to a final concentration of
0.0015N while mixing vigorously. Transfer sample to a 4-L pressure vessel. Filter through a
47-mm-diam 3.0-, 0.45-, and 0.25-µm-porosity fiberglass-acrylic resin filter series at a flow rate
of no more than 130 mL/min and discard filtrate. Rinse filters with 25 mL 0.14N NaCl to remove
excess Al3+. Pipet NaCl solution directly into filter inlet or place in a small pressure vessel
connected to the inlet. Use positive pressure to pass NaCl solution through filter and discard
filtrate. Elute adsorbed viruses from filter with 7-mL portions of either pH 10.5 or 11.5
glycine-NaOH or 3% beef extract, pH 9.0. Pipet 7 mL eluent directly into filter holder inlet or
into a small pressure vessel connected to filter inlet and connect to a positive pressure source.
Carefully apply positive pressure so that eluate flows slowly from filter outlet into a sterile
container. When filtrate no longer flows from outlet, increase pressure to force retained fluid
from filters. If using pH 11.5 glycine-NaOH, measure eluate pH and immediately adjust to pH
between 7.5 and 9.5 with pH 1.5 glycine-HCl. Repeat this elution procedure with another 7-mL
portion of pH 11.5 glycine-NaOH. Complete reconcentration within 5 min. If neither eluate
portion had a final pH of 11.0 or more, repeat elution procedure with additional 7-mL portions of
pH 11.5 glycine-NaOH until an eluate portion has a pH of at least 11.0. Combine all eluates.
If using pH 10.5 glycine-NaOH, elute five successive times with 7-mL volumes of eluent.
After each elution, readjust eluate to pH 10.5 with 0.1N NaOH while mixing vigorously. After
the fifth elution, adjust eluate to pH 7.4 with pH 1.5 glycine-HCl or 0.1N HCl while mixing
vigorously.
If using 3% beef extract, pH 9.0, elute with two 7-mL volumes, combine filtrates, and adjust
to pH 7.4 if necessary.
Measure total eluate volume. For glycine eluates, add 1/10th the measured sample volume of
10X Hanks balanced salt solution and 10X nutrient broth. To all eluates add appropriate volumes
of antibiotics (1/10th volume penicillin-streptomycin or 1/100th volume gentamycin-kanamycin,
or both). Store at 4 or −70°C, depending on time until virus assay.
Further concentrate viruses in beef extract eluates by precipitation at pH 3.5 (organic
flocculation). Viruses in glycine eluates also can be reconcentrated by this technique by first
supplementing them with beef extract to a final concentration of 1 to 3%. Use sterile beef extract
paste (about 80% beef extract) or sterile 20% beef extract solution made from powder to bring
glycine eluates to the desired beef extract concentration. While mixing vigorously, adjust eluate
to pH 3.5 by adding 1N HCl dropwise. Continue to mix at slow speed for 30 min and centrifuge
at 3000 × g for 10 min. Decant and discard supernatant. With vigorous mixing, resuspend
sediment in 1/20 the initial sample volume of 0.45N Na2HPO4. Add antibiotics (1/10 final
sample volume penicillin-streptomycin, 1/100 final sample volume gentamycin-kanamycin, or
both) and while mixing vigorously adjust to pH 7.4 with 1.0 or 0.1N NaOH. Check electrical
conductivity of sample. If conductivity is > 13 000 µmhos, dialyze sample against Hanks
balanced salt solution before assay. Store at 4 or −70°C, depending on time until virus assay.
6. References
1. KATZENELSON, E., B. FATTAL & T. HOSTOVESKY. 1976. Organic flocculation: an
efficient second-step concentration method for the detection of viruses in tapwater.
2. BITTON, G., B.N. FELDBERG & S.R. FARRAH. 1979. Concentration of enteroviruses from
seawater and tap water by organic flocculation using non-fat dry milk and casein water.
Air Soil Pollut. 10:187.
3. PAYMENT, P. & M. TRUDEL. 1980. A simple low cost apparatus for conditioning large
volumes of water for virological analysis. Can. J. Microbiol. 26:548.
4. PAYMENT, P. & M. TRUDEL. 1981. Improved method for the use of proportioning
injectors to condition large volumes of water for virological analysis. Can. J.
Microbiol. 27:455.
9510 D.
Virus Concentration by Aluminum Hydroxide
Adsorption-Precipitation
Viruses can be concentrated from small volumes of water, wastewater, and adsorbent filter
eluates by precipitation with aluminum hydroxide.1-4 This process probably involves both
electrostatic interactions between the negatively charged virus surface and the positively charged
aluminum hydroxide [Al(OH)3] surfaces and coordination of the virus surface by
hydroxo-aluminum complexes.5 Viruses are adsorbed to an Al(OH)3 precipitate that is either
added to the sample or formed in the sample from a soluble aluminum salt and a base such as
sodium carbonate (Na2CO3) or sodium hydroxide (NaOH). Viruses are allowed to adsorb to the
Al(OH)3 precipitate and the virus-containing precipitate is collected by filtration or
centrifugation. The recovered precipitate may be inoculated directly into laboratory hosts for
virus assay or the viruses are eluted from the precipitate with an alkaline buffer or a
proteinaceous solution before virus assay.
The major limitations of this method are that sample size is limited to perhaps a few liters,
soluble organic matter can interfere with virus adsorption, and virus recovery from the
precipitate may be incomplete. Virus adsorption may be improved by forming the Al(OH)3
precipitate in the sample instead of adding it preformed. Although virus adsorption can be
maximized by using large amounts of Al(OH)3, the adsorbed viruses become more difficult to
elute. Therefore, some intermediate amount of Al(OH)3 is used to achieve maximum virus
recovery. Also, Al(OH)3 is a relatively nonspecific adsorbent so that other substances may be
concentrated with viruses. The presence of such impurities may cause the concentrated sample to
be toxic for the cell cultures normally used for virus assay.
Several modifications of the Al(OH)3 adsorption-precipitation procedure have been used to
concentrate viruses from water, wastewater, and eluates from adsorbent filters. Initially,
preformed Al(OH)3 precipitates were made by adding Na2CO3 to AlCl3 solutions and the
Al(OH)3 precipitate was resuspended in 0.15N NaCl. This was added to the wastewater and the
mixture was stirred gently for 1 h or more to allow viruses to adsorb to the precipitate. The
precipitate was recovered by filtration, resuspended in cell culture media, and inoculated into
cell cultures.3,4 More recent procedural modifications include: (a) Al(OH)3 precipitate formation
within the sample,1,2,6-8 (b) recovery of the Al(OH)3 precipitate by centrifugation followed by
elution of viruses from the precipitate with alkaline eluents,1,2,6,7 and (c) a large-volume method
in which the precipitate is formed in the sample and collected on a cartridge filter, and viruses
are eluted from the precipitate on the filter with alkaline eluent.9 The method described here is
for relatively small sample volumes and uses Al(OH)3 that is either preformed or generated
within the sample. The latter modification is preferable because some viruses are not adsorbed
efficiently by preformed precipitates.10
a. Centrifuge, with rotor and buckets, capable of operating at about 1900 × g.
b. Centrifuge bottles and tubes.
c. Beaker, 100-mL or larger.
d. pH meter.
e. Magnetic stirrer and stirring bars or alternative mixing device.
f. Graduated cylinders, 100-mL or larger.
g. Pipets, 1-, 5-, and 10-mL.
h. Laboratory balance.
i. Vacuum-type filter holder or Buchner filter funnel,*#(96) 47-mm diam or larger.
j. Filter flask.*#(97)
k. Spatula,*#(98) flat blade, metal or autoclavable plastic.
l. Vacuum source,*#(99) vacuum pump or laboratory vacuum line.
3. Materials
Filter:* Fiberglass-acrylic resin filter†#(100) or microporous filter, 0.45-µm
porosity,‡#(101) 47-mm diam or larger. To prevent virus adsorption, filter 0.1%
polyoxyethylene sorbitan monooleate solution (¶ 4h) through the filters, using about 1 mL
solution/cm2 of filter surface area. Rinse filter with distilled water, using about 10 mL/cm2 of
filter surface area. Sterilize treated filters by autoclaving.
4. Reagents
a. Hydrochloric acid, HCl, 0.1 and 1.0N.
b. Sodium hydroxide, NaOH, 0.1 and 1.0N.
c. Sodium carbonate, Na2CO3, 4N§#(102).
d. Aluminum chloride, AlCl3, 0.075N§ or 0.9N.
e. Sodium chloride, NaCl, 0.14N.
f. Beef extract, 3%, pH 7.4: Dissolve 3 g beef extract paste or 2.4 g beef extract powder in 90
mL distilled water, adjust to pH 7.4 with 1.0 or 0.1N NaOH, dilute to 100 mL with distilled
water, and sterilize by autoclaving.
g. Antibiotics: Use either:
1) Penicillin-streptomycin, 10X, containing 5000 IU penicillin/mL and 5000 µg
streptomycin/mL. Available commercially or prepare by dissolving powdered sodium or
potassium penicillin-G and streptomycin sulfate in distilled water and sterilizing by filtration.
2) Gentamycin-kanamycin, 100X, containing 5000 µg/mL each of gentamycin base and
kanamycin base (Section 9510B.4i).
h. Polyoxyethylene sorbitan monooleate,i#(103) 0.1% (ν/ν) in distilled water.
5. Procedure
b. Preparation of preformed Al(OH)3 precipitate: While mixing 100 mL 0.075N AlCl3 at
room temperature, slowly add 4N Na2CO3 solution to form precipitate and adjust to pH 7.2.
Continue mixing for 15 min and, if necessary, add more Na2CO3 to maintain pH 7.2. Centrifuge
at 1100 × g for 15 min and discard supernatant. Resuspend sediment in 0.14N NaCl and
recentrifuge. Discard supernatant, resuspend sediment in 0.14N NaCl, and sterilize by
autoclaving. Cool, centrifuge again, decant supernatant, and resuspend Al(OH)3 sediment in 50
mL sterile 0.14N NaCl. Store at 4°C.
c. Sample size, collection, and storage: Process samples of no more than several liters
because the method is too cumbersome and time-consuming for larger volumes. See Section
9510B.5d for sample collection and storage procedures.
d. Sample processing: Do not prefilter sample11,12 because substantial virus losses can
occur. Adjust sample to pH 6.0 with 1.0 or 0.1N HCl while mixing vigorously. Form Al(OH)3
precipitate in sample by adding 1 part 0.9N AlCl3 solution to 100 parts sample to give a final
0.009N Al3+ concentration. Check sample pH and re-adjust to 6.0 with 1.0 or 0.1N NaOH or
HCl, if necessary. Mix slowly for 15 min at room temperature.
Alternatively, use preformed Al(OH)3 precipitate by adding 1 part stock Al(OH)3
suspension/100 parts sample and mix slowly for 2 h at 4 to 10°C to allow for virus adsorption.
Collect virus-containing Al(OH)3 precipitate by centrifugation or filtration. To collect
precipitate by centrifugation, centrifuge at 1700 × g for 15 to 20 min, discard supernatant, and
resuspend sediment in 1/1000 to 1/20 original sample volume of 3% beef extract, pH 7.4.
To collect precipitate by filtration, vacuum filter sample through a treated filter (¶ 3 above)
held in a vacuum-type filter holder or Buchner funnel, using additional filters if filter clogs
before entire sample is filtered. Carefully scrape precipitate from filter(s) with a sterile spatula
and resuspend in 1/1000 to 1/20 original sample volume of 3% beef extract, pH 7.4.
Regardless of collection method, vigorously mix the Al(OH)3 beef extract suspension and, if
necessary, adjust to pH 7.4 with 0.1N HCl or NaOH. Continue mixing for a total of 10 min.
Centrifuge at 1900 × g for 30 min. Decant supernatant, add 1/10 the volume of the concentrate of
penicillin-streptomycin solution or 1/100 volume of gentamycin-kanamycin and store at 4 or
−70°C.
6. References
1. PAYMENT, P., C.P. GERBA, C. WALLIS & J.L. MELNICK. 1976. Methods for concentrating
viruses from large volumes of estuarine water on pleated membranes. Water Res.
10:893.
2. FARRAH, S.R., S.M. GOYAL, C.P. GERBA, C. WALLIS & J.L. MELNICK. 1977. Concentration
of enteroviruses from estuarine water. Appl. Environ. Microbiol. 33:1192.
3. WALLIS, C. & J.L. MELNICK. 1967. Concentration of viruses on aluminum hydroxide
precipitates. In G. Berg, ed. Transmission of Viruses by the Water Route. Interscience
Publ., New York, N.Y.
4. WALLIS, C. & J.L. MELNICK. 1967. Virus concentration on aluminum and calcium salts.
Amer. J. Epidemiol. 85:459.
5. COOKSON, J.T., JR. 1974. The chemistry of virus concentration by chemical methods.
Develop. Ind. Microbiol. 15:160.
6. LYDHOLM, B. & A.L. NIELSEN. 1979. Methods for detection of virus in wastewater
applied to samples from small scale treatment systems. Water Res. 14:169.
7. SELNA, M.W. & R.P. MIELE. 1977. Virus sampling in wastewater-field experiences. J.
Environ. Eng. Div., Proc. Amer. Soc. Civil Eng. 103:693.
8. DOBBERKAU, H.J., R. WALTER & S. RUDIGER. 1981. Methods for virus concentration
from water. In M. Goddard & M. Butler, eds. Viruses and Wastewater Treatment.
Pergamon Press, New York, N.Y.
9. FARRAH, S.R., C.P. GERBA, C. WALLIS & J.L. MELNICK. 1978. Concentration of poliovirus
from tapwater onto membrane filters with aluminum chloride at ambient pH levels.
10. FARRAH, S.R., G.M. GOYAL, C.P. GERBA, R.H. CONKLIN & E.M. SMITH. 1978. Comparison
between adsorption of poliovirus and rotavirus by aluminum hydroxide and activated
sludge flocs. Appl. Environ. Microbiol. 35:360.
11. SOBSEY, M.D., C.P. GERBA, C. WALLIS & J.L. MELNICK. 1977. Concentration of
enteroviruses from large volumes of turbid estuary water. Can. J. Microbiol. 23:770.
12. HOMMA, A., M.D. SOBSEY, C. WALLIS & J.L. MELNICK. 1973. Virus concentration from
sewage. Water Res. 7:945.
9510 E.
Hydroextraction-Dialysis with Polyethylene Glycol
Polyethylene glycol (PEG) hydroextraction is an ultrafiltration process in which the sample
is placed in a cellulose dialysis bag and exposed to PEG, a hygroscopic material. Water and
microsolutes leave the sample by passing across the semipermeable dialysis membrane into the
hygroscopic PEG.1 Viruses and other macrosolutes, including PEG, cannot cross the dialysis
membrane. The sample volume in the dialysis bag is reduced by water loss to the PEG, thereby
concentrating viruses and other macrosolutes. The viruses retained in the dialysis bag are
recovered by opening the bag, collecting the remaining sample, and eluting any viruses possibly
adsorbed to the inner walls of the bag with a small volume of slightly alkaline proteinaceous
solution such as 3% beef extract, pH 9.0. The collected concentrate and eluate are combined and
assayed for viruses.
The main limitations of this method are that only small samples (less than 1 L) can be
processed conveniently, virus elution from the walls of the dialysis bag may be incomplete
unless the elution is done painstakingly, and other macrosolutes in the sample that are
concentrated with viruses may interfere with virus assays by being cytotoxic.
Initial investigations of this method reported low and highly variable virus recoveries from
wastewater.2,3 The type of dialysis tubing and eluent solution as well as the thoroughness of the
elution step have been found to influence virus recovery efficiency. More recently, with
modified procedures, efficient and consistent virus recoveries have been obtained from
wastewater and from adsorbent filter eluates.4,5
a. Beakers, 100-mL or larger.
b. Graduated cylinders, 100-mL or larger.
c. Dialysis tubing clamps.*#(104)
d. Pan, approximately 30 × 30 × 12 cm, autoclavable.
e. Magnetic stirrer and stirring bars or alternative mixing device.
f. Centrifuge, with rotor and buckets, capable of operating at about 1900 × g.
g. pH meter.
h. Pipets, 1-, 5-, and 10-mL.
i. Tape roller†#(105) or similar device to aid in washing the inside walls of dialysis bags
with eluting fluid.
j. Ultrasonic disruptor-emulsifier,‡#(106) probe type, capable of generating 100 W of
acoustical output.
3. Materials
a. Dialysis tubing, seamless, regenerated cellulose, 4.8-nm average pore diameter.§#(107)
b. Polyethylene glycol (PEG),i#(108) dry flakes.
4. Reagents
See Section 9510D.4.
5. Procedure
a. Sterilization of apparatus, materials, and reagents: See Section 9510B.5a. Do not sterilize
PEG.
b. Sample size, collection, and storage: Process samples of no more than a few hundred
milliliters. See Section 9510B.5d for sample collection and storage procedures.
c. Preparation of dialysis tubing: Cut a length of dialysis tubing long enough to
accommodate entire sample. Close one end with a clamp. Do not tie knots to close dialysis
tubing. Fill tubing bag with distilled water, sterilize by autoclaving, and let cool.
d. Sample processing: Aseptically remove dialysis bag from distilled water and drain. Fill
bag with sample and close open end with a second clamp. Place bag in a pan containing a 5-cm
layer of PEG, making sure that bag does not touch pan walls. Cover tubing with an additional 5
cm PEG and store at 4°C (for about 18 h) until sample volume has been reduced to no more than
a few milliliters. (If PEG 6000 is used the process time is reduced to 4 to 6 h.) Although sample
may be allowed to dewater completely, do not let it remain in this state.
Remove dialysis bag from PEG and quickly wash PEG from outside of bag with sterile
distilled water. Remove clamp from one end of bag and carefully collect sample concentrate.
Add about 1/200 to 1/20 the original sample volume of 3% beef extract, pH 9.0, and clamp
closed. Thoroughly wash inside walls of bag with beef extract by rubbing fluid from one end to
the other several times using either fingers or a roller device. Remove clamp from one end of bag
and collect fluid, kneading or squeezing to recover the last traces. Add recovered fluid to
previously collected sample concentrate.
Adjust to pH 7.5 with 1.0 or 0.1N HCl while mixing vigorously. To disperse
solids-associated viruses in sample, stir overnight (about 18 h) in the cold (about 4°C) or treat
with ultrasonics at 100 W for 1 to 2 min. Prevent sample temperature from rising above 37°C
during ultrasonic treatment by chilling in an ice bath. Centrifuge at 1900 × g for 30 min. Decant
supernatant, add 1/10 the volume of the concentrate of penicillin-streptomycin solution or 1/100
volume of gentamycin-kanamycin, and store at 4 or −70°C.
6. References
1. SOBSEY, M.D. 1976. Methods for detecting enteric viruses in water and wastewater. In
G. Berg, H.L. Bodily, E.H. Lennette, J.L. Melnick & T.G. Metcalf, eds. Viruses in
Water. American Public Health Assoc., Washington, D.C.
2. CLIVER, D.O. 1967. Detection of enteric viruses by concentration with polyethylene
glycol. In G. Berg., ed. Transmission of Viruses by the Water Route. Interscience
Publ., New York, N.Y.
3. SHUVAL, H.I., S. CYMBALISTA, B. FATTAL & N. GOLDBLUM. 1967. Concentration of
enteric viruses in water by hydro-extraction and two-phase separation. In G. Berg, ed.
Transmission of Viruses by the Water Route. Interscience Publ., New York, N.Y.
4. WELLINGS, F.M., A.L. LEWIS, C.W. MOUNTAIN & L.V. PIERCE. 1975. Demonstration of
virus in groundwater after effluent discharge onto soil. Appl. Microbiol. 29:751.
5. RAMIA, S. & S.A. SATTAR. 1979. Second-step concentration of viruses in drinking and
surface waters using polyethylene glycol hydroextraction. Can. J. Microbiol. 25:587.
9510 F.
Recovery of Viruses from Suspended Solids in Water and Wastewater
Viruses in the aquatic environment often are associated with solids or particulate matter,
either adsorbed to particulate surfaces or embedded within the solid.1-3 Both freely suspended
and solids-associated viruses are concentrated from water by the methods described above.
There is evidence that solids-associated viruses are not eluted efficiently from adsorbent filters
or from Al(OH)3 precipitates and organic flocs. Recovery of solids-associated viruses by
microporous filter methods employing in-situ elution is inconsistent.2 Solids-associated viruses
on adsorbent filters are eluted more efficiently by disrupting filters in elution fluid than by in-situ
elution,2 but this is cumbersome and time-consuming, especially for large-diameter disk filters
and cartridge filters.
For small volumes of water and wastewater, solids-associated viruses can be recovered
expediently by separating the solids by centrifuging, decanting the supernatant, and eluting
viruses from the solids by resuspending in a small volume of eluent.4 Viruses in the supernatant
can be concentrated by one of the procedures described in Section 9510B, Section 9510C,
Section 9510D, or Section 9510E. Viruses eluted from the resuspended solids are separated from
the solids by centrifuging and are assayed directly or concentrated further by organic
flocculation.5,6 Major limitations of these methods are incomplete virus elution and poor virus
recoveries due to interferences from sample constituents.
a. Centrifuge, with rotor and buckets for 250- to 1000-mL-capacity bottles, capable of
operating at about 1250 × g.
b. Centrifuge bottles, 250- to 1000-mL.
c. pH meter.
d. Laboratory balance.
e. Graduated cylinder, 250-mL or larger.
f. Beaker, 250-mL or larger.
g. Sample bottles, 250-mL or larger.
h. Magnetic stirrer and stirring bars, or alternative mixing device.
i. Pipets, 1-, 5-, and 10-mL.
3. Reagents
a. Hydrochloric acid, HCl, 0.1 and 1.0N.
b. Sodium hydroxide, NaOH, 0.1 and 1.0N.
c. Eluent: Dissolve 10 g beef extract, 1.34 g disodium phosphate heptahydrate,
Na2HPO4⋅7H2O, and 0.12 g citric acid in 90 mL distilled water, adjust to pH 7.0 with 1N HCl or
NaOH, dilute to 100 mL with distilled water, and sterilize by autoclaving.
d. Antibiotics: See Section 9510B.4i.
4. Procedure
b. Sample size, collection, and storage: Collect and process samples of no more than 10 L,
depending on capacity of centrifuge. See Section 9510B.5d for sample collection and storage
procedures.
c. Sample processing: Aseptically transfer 250- to 1000-mL sample volumes to centrifuge
bottles and centrifuge at 1250 × g for 20 min. Decant and pool supernatants for subsequent
processing for viruses by one of the methods for water or wastewater described previously.
Elute viruses from the sedimented solids by resuspending in eluent. Use 40 mL eluent per
quantity of sediment from 250 mL of original sample. Pool resuspended sediments from multiple
centrifuge bottles in a sterile beaker. Alternatively, keep resuspended sediments from small
numbers of centrifuge bottles in the bottles and process them individually. While vigorously
mixing with a magnetic stirrer, adjust to pH 7.0 by slowly adding 1N NaOH or HCl, if necessary.
Reduce mixing speed and continue mixing for 30 min. During this period, check sample pH and
readjust to pH 7.0 as necessary. As an alternative to mixing for 30 min, sonicate samples at 100
W for 15 min in a rosette cooling cell maintained at 4°C. Return sample to centrifuge bottles.
Centrifuge at 1250 × g and 4°C for 15 min, collect supernatant for subsequent assay or further
concentration, and discard the sediment.
If desired, further concentrate viruses from this supernatant by organic flocculation (see
Section 9510E). For supernatants that will be assayed directly for viruses with no further
concentration, adjust to pH 7.4, add 1/10 the volume of sample of penicillin-streptomycin or
1/100 volume of gentamycin-kanamycin and store at 4 or −70°C.
5. References
1. SCHAUB, S.A. & B.P. SAGIK. 1975. Association of enteroviruses with natural and
artificially introduced colloidal solids in water and infectivity of solids-associated
virions. Appl. Microbiol. 30:212.
2. WELLINGS, F.M., A.L. LEWIS & C.W. MOUNTAIN. 1976. Viral concentration techniques
for field sample analysis. In L.B. Baldwin, J.M. Davidson & J.F. Gerber, eds. Virus
Aspects of Applying Municipal Waste to Land. Univ. Florida, Gainesville.
3. WELLINGS, F.M., A.L. LEWIS & C.W. MOUNTAIN. 1974. Virus survival following
wastewater spray irrigation of sandy soils. In J.F. Malina, Jr. & B.P. Sagik, eds. Virus
Survival in Water and Wastewater Systems. Univ. Texas, Austin.
4. BERG, G. & D.R. DAHLING. 1980. Method for recovering viruses from river water solids.
5. GERBA, C.P. 1982. Detection of viruses in soil and aquatic sediments. In C.P. Gerba &
S.M. Goyal, eds. Methods in Environmental Virology. Marcel Dekker, Inc., New York,
N.Y.
6. FARRAH, S.R. 1982. Isolation of viruses associated with sludge particles. In C.P. Gerba
& S.M. Goyal, eds. Methods in Environmental Virology. Marcel Dekker, Inc., New
York, N.Y.
9510 G.
Assay and Identification of Viruses in Sample Concentrates
1. Storage of Sample Concentrates
Because it often is impossible to assay sample concentrates immediately, store them at room
temperature (about 25°C) for up to 2 h or at refrigerator temperatures (4 to 10°C) for up to 48 h
to minimize virus losses. Freeze samples requiring storage longer than 48 h at −70°C or less. Do
not freeze samples at −10 to −20°C because extensive inactivation of some enteric viruses may
occur. Store sample concentrates from finished waters in separate freezers or physically
separated from other virus-containing material in common freezers.
2. Decontamination of Sample Concentrates
Sample concentrates, especially those from wastewater, are likely to be contaminated with
bacteria and fungi that can overgrow cell cultures and interfere with virus detection and assay.
Do not decontaminate by centrifugation or filtration because virus losses are likely to occur. For
many samples, especially those from finished waters, contamination is controlled adequately by
antibiotics such as penicillin-streptomycin or gentamycin-kanamycin that are added immediately
after the sample is obtained. To provide additional protection against fungal contamination, add
amphotericin B or nystatin at concentrations of 2.5 and 50 µg/mL, respectively.1 If
penicillin-streptomycin or gentamycin-kanamycin are inadequate, use one or more additional
antibiotics such as aureomycin, neomycin, or polymyxin B. To maximize the antibiotic effects,
incubate samples for 1 to 3 h at 25 to 37°C after adding the antibiotics. Bacterial destruction is
further enhanced by freezing at −70°C after incubation with antibiotics. Keep samples frozen
until assayed for viruses. To determine if antibiotic treatment has been effective, plate a small
subsample on a general-purpose medium such as plate count agar by the spread plate technique
and incubate at 37°C for 24 to 48 h.
If extensive bacterial contamination persists after antibiotic treatment, treat with chloroform.
Add 1/10 volume of sample of chloroform (CHCl3) and mix vigorously for 30 min at room
temperature or homogenize 1 to 2 min at 4 to 10°C. For phase separation, centrifuge at ≥ 1000 ×
g or store overnight in a refrigerator. Separate sample (upper layer) from CHCl3 (bottom layer)
by aspirating with a pipet and bubble with filter-sterilized air for about 15 min to remove
dissolved CHCl3. It may be necessary to place sample in a sterile, shallow container and expose
it to the atmosphere in a sterile air environment (laminar air flow clean bench or biological safety
cabinet) for up to several hours to remove remaining traces of CHCl3. Do not use ether to
decontaminate samples because of the hazard of explosion or fire.
3. Laboratory Facilities and Host Systems for Virus Assay
Because viruses are obligate, intracellular parasites, they grow (multiply) only in living host
cells. This ability to multiply in, and thereby destroy, their host cells is the basis for virus
detection and assay. The two major host cell systems for human enteric viruses are whole
animals (usually mice) and mammalian cell cultures of primate origin.
A complete description of facilities, equipment, materials, and methods for conducting virus
assays is beyond the scope of this book; see standard handbooks on virology and cell culture.1-4
Virus assay is beyond the capability of most water and wastewater microbiology laboratories. It
should be done only by a trained virologist working in specially equipped virology laboratory
facilities. Take particular care to prevent samples or inoculated hosts from becoming
contaminated with viruses from other sources and to prevent virus cross-contamination arising
from sample concentrates or inoculated hosts. Process and handle samples in a Class II Type I
biological safety cabinet5 or in a ‘‘sterile’’ room or cubicle. The use of such cabinets or facilities
is mandatory for testing drinking water or other finished water samples.
There is no single, universal host system for all enteric viruses. Some enteric viruses, notably
hepatitis A virus, human rotaviruses, and Norwalk-type gastroenteritis viruses, cannot be assayed
routinely in any convenient laboratory host systems. However, most of the known enteric viruses
can be detected by using two or more cell culture systems and perhaps suckling mice. The latter
previously were considered essential for the detection of group A coxsackie-viruses, but recent
studies indicate that the RD cell line may be nearly as sensitive as suckling mice for the isolation
of these viruses as well as other enteroviruses.6,7 In general, the more different host systems
used, the greater the enteric virus recovery rate. However, the number of different host systems
used is limited by practical and economic considerations.
There have been numerous comparative studies on relative sensitivities of various cell
culture systems for enteric virus detection,6-26 but no systematic, comprehensive study has been
reported for enteric virus recoveries from water and wastewater. Primary or secondary human
embryonic kidney (HEK) cell cultures appear to be the single most sensitive host system for
enteric virus isolations, but they are becoming increasingly more difficult to obtain regularly
and, when available from commercial sources, they are expensive. Primary or secondary African
green, cynomolgus, or rhesus monkey or baboon kidney cells are sensitive hosts for many
enteroviruses and reoviruses, but are not particularly suitable for recovering adenoviruses or
group A coxsackieviruses. BGM, a continuous line derived from African green monkey kidney
cells, may be comparable in sensitivity to primary monkey kidney cells for enteric virus
recovery.7,18,21,25,26 A number of other continuous cell lines as well as human fetal diploid cell
strains have been evaluated for enteric virus recoveries. Some human fetal diploid cell strains
give virus isolation rates comparable to primary monkey kidney cells, but plentiful supplies of
specific human fetal diploid cell strains are not readily available and many are difficult to
maintain. Furthermore, each different cell strain must be characterized for virus susceptibility.
Most continuous cell lines generally are less effective than primary cells, but comparable
isolation rates for some enteric virus groups have been obtained with Hep-211 and HeLa17,25
cells.
Assay the entire sample concentrate for enteric viruses, using at least two different host
systems and dividing entire sample equally among the hosts. Preferably use primary (or
secondary) HEK cells with either primary (or secondary) monkey kidney or BGM cells for the
recovery of most enteroviruses, adenoviruses, and reoviruses. Additional use of either suckling
mice or RD cells provides for enhanced recovery of group A coxsackieviruses. Different host
systems may be substituted for these if it is demonstrated that they have equivalent sensitivity.
4. Virus Quantitation Procedures for Sample Concentrates
a. Advantages and disadvantages of different quantitation procedures: Virus assays in
suckling mice or other animals are quantal assays and in cell cultures they can be done either by
quantal (most probable number or 50% endpoint) or enumerative (plaque) methods. Selection
between cell culture assay methods depends on the sample and the choice between achieving
either maximum virus sensitivity or maximum precision and accuracy in estimating virus
concentration. The plaque technique generally is more precise and accurate than the quantal
assay because relatively large numbers of individual infectious units can be counted directly as
discrete, localized areas of infection (plaques). Quantal assays are more sensitive than monolayer
plaque assays, but are less sensitive than an agar cell suspension plaque assay.26
Because virus plaques are discrete areas of infection arising from a single infectious virus
unit, it is relatively easy to recover viruses from individual plaques and then to inoculate them
into additional cell cultures to obtain a pure virus culture for identification. However, large
proportions of so-called ‘‘false-positive’’ plaques that do not confirm as virus-positive when
material from these plaques is further passaged in cell cultures have been reported.27,28 Whether
this problem is due to nonviral, plaque-like areas of cytotoxicity from the sample or to technical
inability to passage viruses successfully from the initial plaques remains uncertain.27,28
The use of specific plaque assay conditions for optimizing the recovery of certain enteric
virus groups may preclude efficient recovery of other enteric groups requiring different plaque
assay conditions. Furthermore, some viruses, such as adenoviruses, do not form plaques
efficiently under any conditions. Cytotoxicity due to water or wastewater constituents in sample
concentrates is difficult to control in plaque assay systems because the agar overlay medium is
difficult to remove and replace.
A potential limitation of quantal assays is the possibility that two or more different virus
types will be inoculated into the same cell culture and thus produce a simple positive culture.
This not only results in an underestimation of virus concentration but also requires separation of
the individual virus types by further passage in cell culture. Such mixed cultures may go
undetected unless virus isolates are identified serologically. Recent results indicate that mixed
positive cultures are encountered rarely when samples are divided into small portions for
inoculation into a series of replicate cell cultures.7,25
Cytotoxicity due to constituents of sample concentrates usually can be controlled in quantal
assay cell cultures by replacing the culture medium before the cells die.
b. Cell culture procedures for virus isolation and assay: To assay sample concentrates in cell
cultures by quantal or plaque methods, drain the medium from newly confluent cultures. To
reduce toxicity, rinsing with buffered saline solution may be helpful. Inoculate with unit volumes
of sample. Use no more than 0.06 mL sample/cm2 of cell layer surface, e.g., maximum volumes
of 1.0, 3.0, and 6.0 mL in cell culture flasks with areas of 25, 75, and 150 cm2, respectively.29 If
samples are expected to contain such large quantities of viruses that it would be difficult to make
reliable estimates of concentration, inoculate cell cultures with small sample volumes or
dilutions of concentrates. Allow viruses to adsorb to cells for 2 h at 37 ± 0.5°C. Redistribute
inoculum over the cell layer manually every 15 min or keep cultures on a mechanical rocker
during the adsorption period. Add liquid maintenance medium to cultures for quantal assays or
agar-containing medium for plaque assays. Invert plaque assay cultures so that cell (agar) side of
culture faces up and incubate at 37°C.
Microscopically examine quantal assay cultures for the appearance of cytopathic effects
(CPE) daily during the first 3 d and then periodically for a total of at least 14 d. Do not change
cell culture medium unless cytotoxicity or cell deterioration occurs. Freeze cultures developing
CPE at −70°C when more than 75% of the cells become involved. After 14 or more days, freeze
at −70°C all remaining cultures, including those remaining negative for CPE as well as controls.
Thaw cultures and clarify culture fluid-cell lysate by slow-speed centrifugation or filtration
through sterile 0.22- or 0.45-µm porosity filters. Inoculate clarified material from each initial
(first-passage) culture into a second (second-passage) culture by transferring 20% of the total
initial culture into newly confluent cell cultures of the same type. Microscopically examine
second-passage cultures for development of CPE periodically over a period of 14 or more days.
Consider second-passage cultures developing CPE as confirmed virus-positive. Freeze and store
at −70°C for virus identification. Discard as negative any virus cultures negative for CPE after
this second incubation period of 14 or more days.
Periodically examine plaque assay cultures for appearance of plaques over a 14-d period.
Mark and tally plaques as they appear. Transfer viruses from each plaque directly to at least two
newly confluent, liquid-medium cell cultures of the same type27 before plaques become too large
and grow together or before the entire cell layer deteriorates. Do not store material obtained from
plaques before transfer to new cell cultures, as this may result in loss of virus titer and
unsuccessful transfers. Microscopically examine these second-passage cultures periodically over
14 d for development of CPE. Freeze cultures developing CPE at −70°C for virus identification.
c. Virus isolation and assay in mice: To detect group A and B coxsackieviruses in mice,
inoculate samples into animals no older than 24 h using standard procedures.2,3,8 Use either the
intracerebral or intraperitoneal route, inoculating 0.02 and 0.05 mL, respectively. Observe mice
daily over a 14-d period for development of weakness, tremors, and either flaccid (due to group
A coxsackieviruses) or spastic (due to group B coxsackieviruses) paralysis. Sacrifice animals
developing symptoms, and using sterile technique, prepare 20% tissue suspensions in Hanks
balanced salt solution of the entire skinned, eviscerated torso or just the brain and legs. Store
suspensions at −70°C until used for further passage and identification. For second passage in
mice, follow general procedures used for the initial inoculations. However, making a second
passage in cell cultures is preferable to making a second passage in mice because it is easier to
do subsequent virus identification by neutralization tests.
d. Estimating virus concentration: Determining the amount of virus in a sample concentrate
depends on the assay used. If a sample concentrate is assayed in cell cultures by the plaque
technique, count all plaques and calculate the virus concentration, expressed as plaque-forming
units (PFU).
If a sample concentrate is assayed by the quantal method, estimate the virus concentration
by the most probable number (MPN) method and express as most probable number of infectious
units (MPNIU), or by a 50% end point method and express as 50% infectious or lethal dose
(ID50 or LD50).2,4,30-32 If the undiluted sample concentrate or a single sample dilution was
inoculated into a series of replicate cell cultures (or mice), calculate the MPNIU from the
number of confirmed CPE-negative cultures (or mice), q, per total number of cultures (or mice)
inoculated, n, according to the formula
MPN = −ln(q/n)
If more than one sample dilution was inoculated into cell cultures (or mice), calculate the
MPNIU from the formula developed by Thomas:33
where:
P = total number of positive cultures (or mice) from all dilutions,
N = total mL sample inoculated for all dilutions, and
Q = total mL sample in all negative cultures (or mice).
In using this formula, exclude from the computation all dilutions containing only positive
cultures (or mice).
For MPN values obtained from a single sample dilution, the 95% confidence interval is
based on the standard error of the binomial distribution when more than 30 cultures (or mice) are
inoculated or from the confidence coefficient table of Crow32,34 when 30 or fewer cultures (or
mice) are inoculated.
Make 50% end-point estimates arithmetically by either the Reed-Muench or Karber
method.2,4 These methods require results from several equally spaced sample dilutions,
preferably with about the same number of dilutions above and below the 50% end point, and
may not be useful for sample concentrates containing relatively low virus levels.
e. Identification of virus isolates: Identify enteric viruses isolated from sample concentrates
by standard serological techniques, although preliminary identification of genus (enterovirus,
reovirus, or adenovirus) sometimes can be made on the basis of information obtained from the
isolation procedure. Enteric viruses recovered in suckling mice are likely to be either group A or
B coxsackieviruses. For enteric viruses isolated in cell cultures, preliminary identification of
genus often can be made from the characteristic appearance of cytopathic effects (CPE) in
infected cell cultures.
Confirm preliminary identification of suspected adenovirus and reovirus isolates by
detecting their respective group specific antigens by complement fixation tests using clarified,
second-passage cell-culture lysate as the antigen. Identify specific reovirus serotypes by
hemagglutination-inhibition (HI) or neutralization (Nt) tests. Adenovirus serotypes can be
separated into four groups on the basis of their ability (or inability) to hemagglutinate rhesus
monkey or rat erythrocytes.2,3,8 Except for type 18, the first 28 numbered adenoviruses can be
identified as to specific serotype by HI. Alternatively, identify all adenovirus serotypes by Nt
tests using either individual type-specific antisera or intersecting antisera pools. Also identify
specific enterovirus serotypes by neutralization tests in cell cultures using intersecting pools of
hyperimmune sera.2,3,8,35 Use mice for Nt tests for group A and B coxsackieviruses only if the
virus isolates fail to propagate in cell cultures.36 Because polioviruses often are the most
prevalent enteroviruses in water and wastewater, test enterovirus isolates for neutralization by an
antisera pool against the three types of poliovirus before making neutralization tests with
intersecting antisera pools.
5. References
1. PAUL, J. 1975. Cell and Tissue Culture, 5th ed. Churchill Livingstone, New York, N.Y.
2. LENNETTE, E.H. & N.J. SCHMIDT, eds. 1979. Diagnostic Procedures for Viral and
Rickettsial Infections, 5th ed. American Public Health Assoc., Washington, D.C.
3. LENNETTE, E.H., A. BALOWS, W.J. HAYSLER & J.P. TRUANT, eds. 1980. Manual of
Clinical Microbiology, 3rd ed. American Soc. Microbiology, Washington, D.C.
4. FRESHNEY, R.I. 1987. Culture of Animal Cells. A Manual of Basic Technique, 2nd ed.
Alan R. Liss, Inc., New York, N.Y.
5. U.S. PUBLIC HEALTH SERVICE. 1976. Guidelines for Research Involving Recombinant
DNA Molecules. Appendix D-I, Biological Safety Cabinets. National Inst. Health,
Bethesda, Md.
6. SCHMIDT, N.J., H.H. HO & E.H. LENNETTE. 1975. Propagation and isolation of group A
coxsackieviruses in RD cells. J. Clin. Microbiol. 2:183.
7. SCHMIDT, N.J., H.H. HO, J.L. RIGGS & E.H. LENNETTE. 1978. Comparative sensitivity of
various cell culture systems for isolation of viruses from wastewater and fecal samples.
8. HSIUNG, G.D. 1982. Diagnostic Virology, 3rd ed. Yale Univ. Press, New Haven, Conn.
9. KELLY, S., J. WINSSER & W. WINKELSTEIN. 1957. Poliomyelitis and other enteric viruses
in sewage. Amer. J. Pub. Health 47:72.
10. KELLY, S. & W.W. SANDERSON. 1962. Comparison of various tissue cultures for the
isolation of enteroviruses. Amer. J. Pub. Health 52: 455.
11. PAL, S.R., J. MCQUILLIN & P.S. GARDNER. 1963. A comparative study of susceptibility of
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
primary monkey kidney cells, Hep 2 cells and HeLa cells to a variety of faecal viruses.
J. Hyg., Camb. 61:493.
LEE, L.H., C.A. PHILLIPS, M.A. SOUTH, J.L. MELNICK & M.D. YOW. 1965. Enteric virus
isolations in different cell cultures. Bull. World Health Org. 32:657.
SCHMIDT, N.J., H.H. HO & E.H. LENNETTE. 1965. Comparative sensitivity of human fetal
diploid kidney cell strains and monkey kidney cell cultures for isolation of certain
human viruses. Amer. J. Clin. Pathol. 43:297.
BERQUIST, K.R. & G.J. LOVE. 1966. Relative efficiency of three tissue culture systems
for the primary isolation of viruses from feces. Health Lab. Sci. 3:195.
HERRMANN, E.C. 1967. The usefulness of human fibroblast cell lines for the isolation of
viruses. Amer. J. Epidemiol. 85:200.
FAULKNER, R.S. & C.E. VAN ROOYEN. 1969. Studies on surveillance and survival of
viruses in sewage in Nova Scotia. Can. J. Pub. Health 60:345.
LUND, E. & C.E. HEDSTROM. 1969. A study on sampling and isolation methods for the
detection of virus in sewage. Water Res. 3:823.
SHUVAL, H., B. FATTAL, S. CYMBALISTA & N. GOLDBLUM. 1969. The phase-separation
method for the concentration and detection of viruses in water. Water Res. 3:225.
SCHMIDT, N.J. 1972. Tissue culture in the laboratory diagnosis of virus infections.
Amer. J. Clin. Pathol. 57:820.
COONEY, M.K. 1973. Relative efficiency of cell cultures for detection of viruses. Health
Lab. Sci. 4:295.
DAHLING, D.R., G. BERG & D. BERMAN. 1974. BGM: A continuous cell line more
sensitive than primary rhesus and African green kidney cells for the recovery of viruses
from water. Health Lab. Sci. 11:275.
SCHMIDT, N.J., H.H. HO & E.H. LENNETTE. 1976. Comparative sensitivity of the BGM
cell line for the isolation of enteric viruses. Health Lab. Sci. 13:115.
HATCH, M.H. & G.E. MARCHETTI. 1971. Isolation of echoviruses with human embryonic
lung fibroblast cells. Appl. Microbiol. 22:736.
RUTALA, W.A., D.F. SHELTON & D. ARBITER. 1977. Comparative sensitivities of viruses
to cell cultures and transport media. Amer. J. Clin. Pathol. 67:397.
IRVING, L.G. & F.A. SMITH. 1981. One-year survey of enteroviruses, adenoviruses and
reoviruses isolated from effluent at an activated-sludge purification plant. Appl.
MORRIS, R. & W.M. WAITE. 1980. Evaluation of procedures for recovery of viruses from
water—II detection systems. Water Res. 14: 795.
LEONG, L.Y.C., S.J. BARRETT & R.R. TRUSSELL. 1978. False-positives in testing of
secondary sewage for enteric viruses. Abs. Annu. Meeting, American Soc.
KEDMI, S. & B. FATTAL. 1981. Evaluation of the false-positive enteroviral plaque
29.
30.
31.
32.
33.
34.
35.
36.
phenomenon occurring in sewage samples. Water Res. 15:73.
PAYMENT, P. & M. TRUDEL. 1985. Influence of inoculum size, incubation temperature,
and cell culture density on virus detection in environmental samples. Can. J.
Microbiol. 31:977.
CHANG, S.L., G. BERG, K.A. BUSCH, R.E. STEVENSON, N.A. CLARKE & P.W. KABLER. 1958.
Application of the Most Probable Number method for estimating concentrations of
animal viruses by tissue culture technique. Virology 6:27.
CHANG, S.L. 1965. Statistics of the infective units of animal viruses. In G. Berg, ed.
Transmission of Viruses by the Water Route. Interscience Publ., New York, N.Y.
SOBSEY, M.D. 1976. Field monitoring techniques and data analysis. In L.B. Baldwin,
J.M. Davidson & J.F. Gerber, eds. Virus Aspects of Applying Municipal Waste to
Land. Univ. Florida, Gainesville.
THOMAS, H.A., JR. 1942. Bacterial densities from fermentation tube tests. J. Amer.
CROW, E.L. 1956. Confidence intervals for a proportion. Biometrika 43:423.
MELNICK, J.L., V. RENNICK, B. HAMPIL, N.J. SCHMIDT & H.H. HO. 1973. Lyophilized
combination pools of enterovirus equine antisera: Preparation and test procedures for
the identification of field strains of 42 enteroviruses. Bull. World Health Org. 48:263.
MELNICK, J.L., N.J. SCHMIDT, B. HAMPIL & H.H. HO. 1977. Lyophilized combination
pools of enterovirus equine antisera: Preparation and test procedures for the
identification of field strains of 19 group A coxsackievirus serotypes. Intervirology
8:1720.
9610
DETECTION OF FUNGI*#(109)
9610 A.
Introduction
1. Significance
Fungi, including yeasts and filamentous species or molds, are ubiquitously distributed,
achlorophyllous, heterotrophic organisms with organized nuclei and usually with rigid walls.
They may be found wherever nonliving organic matter occurs, although some species are
pathogenic and others are parasitic. In spring water near the source, the number of fungus spores
usually is minimal. Unpolluted stream water has relatively large numbers of species representing
the true aquatic fungi (species possessing flagellated zoospores and gametes), aquatic
Hyphomycetes, and soil fungi. Moderately polluted water may carry cells or spores of the three
types; however, it has fewer true aquatic fungi and aquatic Hyphomycetes, and soil fungi are
more numerous. Heavily polluted water has large numbers of soil fungi. The group designated as
soil fungi includes yeast-like fungi, many species of which have been isolated from polluted
waters.
The association between fungal densities and organic loading suggests that fungi may be
useful indicators of pollution. Unfortunately, no single species or group of fungi has been
identified as important in this role. There may be some exceptional special cases; for example,
the principal distinction between the yeasts Candida lambica and C. krusei is the ability to use
pentose sugars. Because the former species grows well on pentoses, it could be used as an
indicator of pulp and paper mill wastes, which are high in such sugars. Certain species of yeasts
and filamentous fungi are characteristic of warmer waters and may be useful indicators of
thermal pollution.
Because fungi possess broad enzymatic capabilities, they can degrade actively most complex
natural substances and certain synthetic compounds, including some pesticides. Most fungi are
aerobic or microaerophilic, although a few species show limited anaerobic metabolism and a
very few are capable of totally anaerobic growth.1 Some species do not require light.
2. Occurrence and Survival
Fungi are present in, and have been recovered from, diverse, remote, and extreme aquatic
habitats including lakes, ponds, rivers, streams, estuaries, marine environments, wastewaters,
sludge, rural and urban stormwater runoff, well waters, acid mine drainage, asphalt refineries, jet
fuel systems, and aquatic sediments.
a. Fungi in potable water: Fungi have been found in potable water2-7 and on the inner
surface of distribution system pipes.8 Either they survive water treatment or they enter the
system after treatment and remain viable. Tuberculate macroconidia of Histoplasma capsulatum9
can pass through a 0.75-m rapid sand filter. Plain sedimentation or alum flocculation and settling
removed 80 to 99% of the spores. If these relatively large (8- to 14-µm) globose, tuberculate
macroconidia pass through treatment, it is not surprising that other fungi, typically with smaller
spores, are found in treated water.
Having survived treatment or having been introduced after treatment, fungal spores can
remain viable for extended periods of time. Pathogenic fungi have been stored effectively in
sterile distilled water for relatively long periods.10 Spores of H. capsulatum, stored in raw Ohio
River water and sterile tap water remained highly infective for mice after 400 d.11
Tastes and odors in potable water are associated with the presence of procaryotic organisms
such as bacteria, actinomycetes, and cyanobacteria. However, fungi may be involved.6,7
Propagules from 19 genera of filamentous fungi have been isolated from a chlorinated
surface water system and a nonchlorinated groundwater distribution system.2 The mean number
of colony-forming units (CFU) was 18/100 mL in the groundwater system and 34/100 mL in the
surface system.
In Finland,3 fungi were isolated from rivers, lakes, and ponds supplying nine communities
with sand-filtered water, three with artificially recharged groundwater, of which two used
chemical coagulation, and three with chemically coagulated and disinfected water. Mesophilic
fungi were common in all raw water samples; however, thermotolerant fungi were more
abundant in river than in lake water. Chemical coagulation and disinfection proved far more
efficient in removing fungi than sand filtration and disinfection. Aspergillus fumigatus was the
most common fungus.
Five chlorinated groundwater systems4 in the U.S. yielded an average count per positive
sample of about 5.5 CFU/100 mL. In France5 yeasts were recovered from 50% of 38 samples
and filamentous fungi from 81%.
Except for Aspergillus fumigatus3,4, A. flavus,4,5 and A. niger,4 the fungi isolated from
potable water usually are not considered to be medically important. Fungus infections may be
significant for individuals with compromised immune systems. Most of the fungi are common
soil saprobes.
b. Fungi in recreation waters: Some fungi pathogenic to humans may be expected in
recreational waters such as pools and beaches and in accompanying washing facilities such as
shower stalls.
Trichophyton mentagrophytes, the cause of tinea pedis or athlete’s foot, has been isolated12
from the wooden flooring of a shower stall. Seven species of pathogenic and potentially
pathogenic fungi were isolated from 361 samples of beach sand in Hawaii.13 Beach sand on the
German Baltic coast, in Portugal, and on the Adriatic coast yielded Epidermophyton.14
c. Survival on chlorination: An unidentified yeast was isolated from other organisms that
survived chlorination of wastewater effluents.15 This yeast survived 1 mg free chlorine/L for 20
min in contrast with E. coli, which failed to survive 5 min contact with 0.03 mg free chlorine/L.
The amount of chlorine for fungus control is known at least for C. albicans. It has been shown16
that cells of C. albicans were inactivated effectively with 4 mg chlorine/L in 30 min when the
initial cell count was 105 cells/mL. In an Illinois study with C. parapsilosis,17,18 a commonly
isolated yeast known to cause health problems in the tropics, greater amounts of chlorine were
required to inactivate the organism than coliform bacteria. Mechanisms of inactivation by
chlorine on assimilative stages of yeasts and other microorganisms have been suggested.19
Fungal cells, especially conidia, can survive much higher doses of chlorine than coliform
bacteria,20 including a 10-min exposure to 10 mg chlorine/L when the initial spore count was
approximately 106/mL.
3. Growth Patterns and Identification
In water there are two basic patterns of fungal growth. True aquatic fungi produce zoospores
or gametes that are motile by means of flagella, either of the whiplash or tinsel type. Some fungi,
particularly the Trichomycetes (fungi that inhabit the hind gut of certain worms, mosquito
larvae, etc.), have amoeboid stages. Aquatic fungi typically are collected by exposing suitable
baits (solid foodstuffs) in the habitat being examined or in a sample within the laboratory.
Relatively little work on these fungi in polluted water has been done in the U.S. They have been
studied more extensively in polluted waters in England, Germany, and Japan.
The second fungal growth form is nonmotile in all stages of the life cycle. Growth and
reproduction usually are asexual (anamorphic). Three growth processes have been recognized:
(a) filamentous growth with blastic spores or spores produced in special structures; (b)
filamentous growth with the filaments breaking up in an arthric (fragmenting) manner to form
separate spores called arthroconidia as in Geotrichum and related genera; and (c) single-celled
growth produced on each parent cell, called budding, typical of the yeasts.
Identification of fungi, which are considerably larger than bacteria, is dependent on colonial
morphology on a solid medium, growth and reproduction morphology, and, for yeasts,
physiological activity in laboratory cultures. Increasing numbers of fungi usually indicate
increasing organic loadings in water or soil. Large numbers of similar fungi suggest excessive
organic load while a highly diversified mycobiota indicates populations adjusted to the
environmental organics. Despite their wide occurrence, little attention has been given to the
presence and ecological significance of fungi in aquatic habitats. The relevance of fungi and
their activities in water is emphasized by increasing knowledge of their pathogenicity for
humans, animals, and plants; their role as food or energy sources; their activity in natural
purification processes; and their function in sediment formation.
A survey of the literature of fungi occurring in water, wastewater, and related organically
polluted substrata listed 984 species21: 133 species were assigned to the Mastigomycotina (fungi
with flagellated zoospores); 79 to the Zygomycotina, mostly mucoraceous fungi; 161 to the
Ascomycotina, including perfect (teleomorphic) states of some of those assigned to the Fungi
Imperfecti; 18 to the Basidiomycotina, including perfect states of several yeasts; and 593 to the
Deuteromycotina or Fungi Imperfecti. Of the total, 133 species were zoosporic, 131 species were
yeast-like, and 718 species were filamentous. Most zoosporic species were recovered from
mildly polluted or unpolluted waters; of the remaining species fewer than half were recovered in
numbers large enough to indicate membership in a population for even a brief period of time.
The significance of fungi in both aquatic and terrestrial environments has been discussed in
detail.22-38
Quantitative enumeration of fungi is not equivalent to that of unicellular bacteria because a
fungal colony may develop from a single cell (spore), an aggregate of cells (a cluster of spores or
a single multi-celled spore), or from a mycelial or pseudomycelial fragment (containing more
than one viable cell). It is assumed that each fungal colony developing in laboratory culture
originates from a single colony-forming unit (CFU), which may or may not be a single cell.
4. References
1. TABAK, H. & W.B. COOKE. 1968. Growth and metabolism of fungi in an atmosphere of
nitrogen. Mycologia 60:115.
2. NAGY, L.A. & B.H. OLSON. 1982. The occurrence of filamentous fungi in water
distribution systems. Can. J. Microbiol. 28:667.
3. NIEMI, R.M., S. KUNTH & K. LUNDSTROM. 1982. Actinomycetes and fungi in surface
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
waters and potable water. Appl. Environ. Microbiol. 43:378.
ROSENZWEIG, W.D., H. MINNIGH & W.O. PIPES. 1986. Fungi in potable water distribution
systems. J. Amer. Water Works Assoc. 78: 53.
HINZELIN F. & J.C. BLOCK. 1985. Yeast and filamentous fungi in drinking water.
Environ. Tech. Letters 6:101.
BURMAN, N.P. 1965. Symposium on consumer complaints. 4. Taste and odour due to
stagnation and local warming in long lengths of piping. Proc. Soc. Water Treat. Exam.
14:125.
BAYS, L.R., N.P. BURMAN & W.M. LEWIS. 1970. Taste and odour in water supplied in
Great Britain. A survey of the present position and problems for the future. Proc. Soc.
Water Treat. Exam. 19:136.
NAGY, L.A. & B.H. OLSON. 1985. Occurrence and significance of bacteria, fungi, and
yeasts associated with distribution pipe surfaces. Proc. American Water Works Assoc.,
Water Quality Technology Conf., p. 213.
METZLER, D.F., C. RITTER & R.L. CULP. 1956. Combined effect of water purification
processes on the removal of Histoplasma capsulatum from water. Amer. J. Pub. Health
46:1571.
CASTELLANI, A. 1963. The cultivation of pathogenic fungi in sterile distilled water.
Commentarii 1(10):1.
COOKE, W.B. & P.W. KABLER. 1953. The survival of Histoplasma capsulatum in water.
Lloydia 16:252.
AJELLO, L. & M.E. GETZ. 1954. Recovery of dermatophytes from shoes and shower
stalls. J. Invest. Derm. 22:17.
KISHIMOTO, R.A. & G.E. BAKER. 1969. Pathogenic and potentially pathogenic fungi
isolated from beach sands and selected soils of Oahu, Hawaii. Mycologia 61:539.
MULLER, G. 1973. Occurrence of dermatophytes in the soils of European beaches. Sci.
Total Environ. 2:116.
ENGELBRECHT, R.S., D.H. FOSTER, E.O. GREENING & S.H. LEE. 1974. New microbial
indicators of waste water efficiency. Environ. Protect. Technol. Ser. No. 670/2-73-082.
JONES, J. & J.A. SCHMITT. 1978. The effect of chlorination on the survival of cells of
Candida albicans. Mycologia 70:684.
ENGELBRECHT, R.S. & C.N. HAAS. 1977. Acid-fast bacteria and yeasts as disinfection
indicators: Enumeration methodology. Proc. American Water Works Assoc. Water
Quality Technology Conf. 1977, p. 1.
HAAS, C.N. & R.S. ENGELBRECHT. 1980. Chlorine dynamics during inactivation of
coliforms, acid-fast bacteria, and yeasts. Water Res. 14:1749.
HAAS, C.N. & R.S. ENGELBRECHT. 1980. Physiological alterations of vegetative
microorganisms resulting from chlorination. J. Water Pollut. Control Fed. 52:1976.
ROSENZWEIG, D.W., H.A. MINNIGH & W.O. PIPES. 1983. Chlorine demand and
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
37.
38.
inactivation of fungal propagules. Appl. Environ. Microbiol. 45:182.
COOKE, W.B. 1986. The Fungi of ‘‘Our Mouldy Earth’’. Beihefte zur Nova Hedwigia
85:1.
BARLOCHER, F. & B. KENDRICK. 1976. Hyphomycetes as intermediates of energy flow
in streams. In E. B. Jones, ed. Recent Advances in Aquatic Mycology. Elek Science,
London, England.
COOKE, W.B. 1961. Pollution effects on the fungus population of a stream. Ecology
42:1.
COOKE, W.B. 1965. The enumeration of yeast populations in a sewage treatment plant.
Mycologia 57:696.
COOKE, W.B. 1970. Our Mouldy Earth. FWPCA Res. Contract Ser. Publ. No. CWR,
Cincinnati, Ohio.
COOKE, W.B. 1971. The role of fungi in waste treatment. CRC Critical Rev. Environ.
Control. 1:581.
COOKE, W.B. 1976. Fungi in sewage. In E.B.G. Jones, ed. Recent Advances in Aquatic
Mycology. Elek Science, London, England.
COOKE, W.B. 1979. The Ecology of Fungi. CRC Press, Boca Raton, Fla.
DICK, M.W. 1971. The ecology of Saprolegniales in the lentic and littoral muds with a
general theory of fungi in the lake ecosystem. J. Gen. Microbiol. 65:325.
HARLEY, J.L. 1971. Fungi in ecosystems. J. Appl. Ecol. 8:627.
MEYERS, S.P., D.G. AHEARN & W.L. COOK. 1970. Mycological studies of Lake
Champlain. Mycologia 62:504.
NOELL, J. 1973. Slime-inhabiting geofungi in a polluted stream (winter/spring).
Mycologia 65:57.
PARK, D. 1972. Methods of detecting fungi in organic detritus in water. Trans. Brit.
Mycol. Soc. 58:281.
QURESHI, A.A. & B.J. DUTKA. 1974. A preliminary study on the occurrence and
distribution of geofungi in Lake Ontario near the Niagara River. Proc. 17th Conf. Great
Lakes Research, International Soc. Great Lakes Research.
SHERRY, J.P. & A.A. QURESHI. 1986. Isolation and enumeration of fungi using
membrane filtration. In B.J. Dutka, ed. Membrane Filtration Applications, Techniques,
and Problems. Marcel Dekker, New York, N.Y.
SIMARD, R.E. 1971. Yeasts as an indicator of pollution. Marine Poll. Bull. 12:123.
SPARROW, F.K. 1968. Ecology of fresh water fungi. In G.C. Ainsworth & A.S.
Sussman, eds. The Fungi, An Advanced Treatise. Vol. 3. The Fungal Population.
Academic Press, New York, N.Y.
TOMLINSON, T.G. & I.L. WILLIAMS. 1975. Fungi, In C.R. Curds & H.A. Hawkes, eds.
Ecological Aspects of Used-Water Treatment. Vol. 1. The Organisms and their
Ecology. Academic Press, New York, N.Y.
5. Bibliography
EMERSON, R. 1958. Mycological organization. Mycologia 50:589.
COOKE, W.B. 1958. CONTINUOUS SAMPLING OF TRICKLING FILTER POPULATIONS. I.
PROCEDURES. Sewage Ind. Wastes 30:21.
COOKE, W.B. & A. HIRSCH. 1958. Continuous sampling of trickling filter populations. II.
Populations. Sewage Ind. Wastes 30:139.
COOKE, W.B. 1959. Trickling filter ecology. Ecology 40:273.
SPARROW, F.K. 1959. Fungi. (Ascomycetes, Phycomycetes); including W.W. Scott, Key to
genera, Fungi Imperfecti (Aquatic Hyphomycetes only). In W.T. Edmondson, ed. Ward &
Whipple’s Fresh Water Biology, 2nd ed. John Wiley & Sons, New York, N.Y.
COOKE, W.B. 1963. A Laboratory Guide to Fungi in Polluted Waters, Sewage, and Sewage
Treatment Systems, Their Identification and Culture. USPHS Publ. 999-WP-1, Cincinnati,
Ohio.
FULLER, M.S. & R.O. PAYTON. 1964. A new technique for the isolation of aquatic fungi.
BioScience 14:45.
WILLOUGHBY, L.C. & V.G. COLLINS. 1966. A study of fungal spores and bacteria in Blelham Tarn
and its associated streams. Nova Hedwigia 12:150.
COOKE, W.B. & G.S. MATSUURA. 1969. Distribution of fungi in a waste stabilization pond system.
Ecology 50:689.
BROCK, T.D. 1970. Biology of Microorganisms. Prentice-Hall, Englewood Cliffs, N.J.
COOKE, W.B. 1970. Fungi in the Lebanon sewage treatment plant and in Turtle Creek, Warren
Co., Ohio. Mycopathol. Mycol. Appl. 42:89.
PATERSON, R.A. 1971. Lacustrine fungal communities. In J. Cairns, ed. Structure and Function
of Microbial Communities. Symposium, American Microscopical Soc., Burlington, Vt. 1969.
Virginia Polytechnic Inst. & State Univ. Res. Div. Mono. 3:209.
JONES, E.B.G. 1971. Aquatic Fungi. In C. Booth, ed. Methods in Microbiology 4:335. Academic
Press, New York, N.Y.
FARR, D.F. & R.A. PATERSON. 1974. Aquatic fungi in rivers: Their distribution and response to
pollutants. VPI-WRRC Bull. 68, Virginia Water Resources Research Center, Virginia
Polytechnic Inst. & State Univ., Blacksburg.
GARETH JONES, E.B., ed. 1976. Recent Advances in Aquatic Mycology. Elek Science, London,
England.
FULLER, M.S., ed. 1978. Lower Fungi in the Laboratory. Palfrey Contrib. in Botany 1:1-212.
Athens, Ga.
ALEXOPOULOS, C.J. & C.W. MIMS. 1979. Introductory Mycology, 3rd ed. John Wiley & Sons,
New York, N.Y.
9610 B.
Pour Plate Technique
1. Samples
a. Containers: Collect samples as directed in Section 9060A. Alternatively, use cylindrical
plastic vials with snap-on caps. These vials usually are sterile as received. Transport them in an
upright position to minimize the chance of leakage and discard after use.
b. Storage: Hold samples for not more than 24 h. If analysis is not begun promptly after
sample collection, refrigerate.
2. Media
For counting, neopeptone-glucose-rose bengal-aureomycin® agar is the usual medium of
choice, although experience may indicate that Czapek agar (for Aspergillus, Penicillium, and
related fungi) and yeast extract-malt extract-glucose agar or Diamalt agar (for yeasts) may be
preferable. For inventory, use neopeptone-glucose agar.
a. Neopeptone-glucose-rose bengal aureomycin® agar: Add 5.0 g neopeptone, 10.0 g
glucose, 3.5 mL rose bengal solution (1 g/100 mL reagent-grade water), and 20.0 g agar to 1 L
reagent-grade water. Because this medium is used for making pour plates, prepare and store
basal agar either in bulk, or more conveniently, in tubes in 10-mL amounts. Sterilize by
autoclaving; the final pH should be about 6.5.
Separately prepare a solution of chlortetracycline or tetracycline (1.0 g water-soluble
antibiotic/150 mL reagent-grade water) and refrigerate. Before use, sterilize by filtration. To
complete the medium, add 0.05 mL sterile solution to 10 mL melted basal agar at about 45°C.
This medium may not be available in dehydrated form and may require preparation from the
basic ingredients. Dehydrated Cooke’s rose bengal agar may be used in place of the agar base.
This medium is useful for isolating a broad spectrum of fungal species.
b. Czapek (or Czapek-Dox) agar: Dissolve 30.0 g sucrose, 3.0 g sodium nitrate (NaNO3), 1.0
g dipotassium hydrogen phosphate (K2HPO4), 0.5 g magnesium sulfate (MgSO4), 0.5 g
potassium chloride (KCl), 0.01 g ferrous sulfate (FeSO4), and 15.0 g agar in 1 L reagent-grade
water. The pH should be 7.3 after sterilization.
This medium is useful for isolating species of Aspergillus, Penicillium, Paecilomyces, and
some other fungi with similar physiological requirements.
c. Yeast extract-malt extract-glucose agar: Dissolve 3.0 g yeast extract, 3.0 g malt extract,
5.0 g neopeptone (or equivalent), 10.0 g glucose, and 20.0 g agar in 1 L reagent-grade water. No
adjustment of pH is required.
This medium is useful for isolating yeasts.
d. Diamalt agar: Dissolve 150 g diamalt and 20.0 g agar in 1 L reagent-grade water. No
adjustment of pH is required. The medium will be turbid but filtration is unnecessary.
This medium is useful in the purification of yeast isolates and for study of yeast species in
various specified tests.
e. Neopeptone-glucose agar: Dissolve 5.0 g neopeptone (or equivalent), 10.0 g glucose, and
20.0 g agar in 1 L reagent-grade water. The pH should be about 6.5 after sterilization. (This
medium is known also as Emmons’ Sabouraud Agar or Emmons’ Sabouraud Dextrose Agar.)
This medium is useful for maintaining stock cultures. It is comparable to
neopeptone-glucose-rose bengal aureomycin® agar but contains neither rose bengal nor an
antibiotic.
3. Procedure
As many as 40 samples can be analyzed simultaneously by a single analyst by the following
procedure; however, 20 samples represents the optimum number.
a. Preparation and dilution: To a sterile 250-mL erlenmeyer flask add 135 mL sterile
reagent-grade water and 15 mL sample to obtain a 1:10 sample dilution. Use a sterile measuring
device for each sample, or, less preferably, rinse the measure with sterile reagent-grade water
between samples. Mix sample well before withdrawing the 15-mL portion. Shake flask on a
rotary shaker at about 120 to 150 oscillations/min for about 30 min or transfer flask contents to a
blender jar, cover and blend at low speed for 1 min or high speed for 30 s. Preferably use a
sterile blender jar and appurtenances for each sample or wash jar thoroughly between samples
and rinse with sterile water. Further dilutions may be made by adding 45 mL sterile water to 5
mL of a 1:10 diluted suspension.
For stream water samples a dilution of 1:10 usually is adequate. Dilute samples with large
amounts of organic material, such as sediments, to 1:100 or 1:1000. Dilute stream bank or soil
samples to 1:1000 or 1:10 000.
b. Plating: Prepare five plates for each dilution to be examined. To use
neopeptone-glucose-rose bengal-aureomycin® agar, aseptically transfer 10 mL of medium at
45°C to a 9-cm petri dish. Add 1 mL of appropriate sample dilution and mix thoroughly by
tilting and rotating dish (see plating procedure under heterotrophic plate count, Section 9215).
Alternatively add to petri dish 1 mL sample, 0.05 mL antibiotic solution, and 10 mL liquefied
agar medium at 43 to 45°C. Solidify agar as rapidly as possible. (In arid areas use more medium
to prevent dehydration during incubation.)
c. Incubation: Stack plates but do not invert. Incubate at room conditions of temperature (20
to 24°C) and lighting but avoid direct sunlight. Examine and count plates after 3, 5, and 7 d.
d. Counting and inventory: The fungus plate count will provide the basis for rough
quantitative comparisons among samples; the inventory will give relative importance of at least
the more readily identifiable species or genera.
In preparing plates, use sample portions that will give about 50 to 60 colonies on a plate.
Determine this volume by trial and error. When first examining a new habitat plate at least two
sample dilutions. Estimates of up to 300 colonies may be made, but discard more crowded
plates. The medium containing rose bengal tends to produce discrete colonies and permits
slow-growing organisms to develop.
The inventory includes the direct identification of fungi based on colonial morphology and
the counting of colonies assignable to various species or genera. When discrete colonies cannot
be identified and identification is important, use a nichrome wire with its tip bent in an L-shape
to pick from each selected colony and streak on a slant of neopeptone-glucose agar (¶ 2e). If five
plates are used per sample, the average number of colonies on all plates (total number of
colonies counted/5) times the reciprocal of the dilution (10/1, 100/1, 1000/1, etc.) equals the
fungus colony count per milliliter of original sample. For solid or semisolid samples, use a
correction for the water content to report fungus colonies per gram dry weight. Determine water
content by drying paired 15-mL portions of original sample at 100°C overnight; the difference
between wet and dry weights is the amount of water lost from the sample.
9610 C.
Spread Plate Technique
The spread plate technique is an alternative procedure for obtaining quantitative data on
colony-forming units.
1. Samples
2. Media
Use any of the following media. Aureomycin®-rose bengal-glucose-peptone agar (ARGPA)
(e) and streptomycin-terramycin®-malt extract agar (STMEA) () are useful in analyzing sewage
and polluted waters.1
a. Neopeptone-glucose-rose bengal aureomycin® agar: See Section 9610B.2a.
b. Czapek (or Czapek-Dox) agar: See Section 9610B.2b.
c. Yeast extract-malt extract-glucose agar: See Section 9610B.2c.
d. Diamalt agar: See Section 9610B.2d.
e. Aureomycin®-rose bengal-glucose-peptone agar: Dissolve 10.0 g glucose, 5.0 g peptone,
1.0 g potassium dihydrogen phosphate (KH2PO4), 0.5 g magnesium sulfate (MgSO4⋅7H2O),
0.035 g rose bengal, and 20.0 g agar in 800 mL reagent-grade water and sterilize. Dissolve 70.0
mg aureomycin® hydrochloride in 200 mL reagent-grade water, sterilize by filtration, and add to
the cooled (42 to 45°C) agar base. The pH should be 5.4. Pour 25-mL portions into sterile petri
dishes (100 × 15 mm) and let agar harden. Poured plates may be held up to 4 weeks at 4°C.
f. Streptomycin-terramycin®-malt extract agar: Dissolve 30.0 g malt extract, 5.0 g peptone,
and 15.0 g agar in 800 mL reagent-grade water and sterilize. Dissolve 70.0 mg each of
streptomycin and terramycin® in separate 100-mL portions reagent-grade water, sterilize by
filtration, and add to the cooled (42 to 45°C) agar base. The pH should be 5.4. Pour about 20-mL
portions into sterile petri dishes (60 × 15 mm) and let agar harden. Poured plates may be held up
to 4 weeks at 4°C.
3. Procedure
a. Preparation and dilution: See Section 9215A.5 and Section 9610B.3a. Make dilutions
with buffered water (Section 9050C.1) and select dilutions that yield 20 to 150 colonies per
plate.
b. Plating: Pre-dry plates separately with lids off in a laminar-flow hood at room temperature
and about 30% relative humidity for 1 to 1.5 h. Prepare at least three plates, or five plates if data
are to be analyzed statistically, per sample or dilution. Using a sterile pipet, transfer 1.0 mL of
sample or dilution onto surface of a pre-dried agar plate. Spread sample over entire agar surface
using a sterile L-shaped glass rod or use a mechanical device to rotate plate and ensure proper
sample distribution.
c. Incubation: With dish covers on, let plates dry at room temperature, invert plates, and
incubate at 15°C for 7 d in an atmosphere of high humidity (90 to 95%). Alternatively incubate
at 20°C for 5 to 7 d. Slow-growing fungi may not produce noticeable colonies until 6 or 7 d.
d. Counting and recording: Using a Quebec colony counter, count all colonies on each
selected plate. If counting must be delayed temporarily, hold plates at 4°C for not longer than 24
h. Depending on colony size, plates with as many as 150 colonies can be counted but the optimal
maximum number is 100 colonies.
Record results as colony-forming units (CFU)/100 mL original sample. For solid or
semisolid samples report CFU/g wet or dry, preferably dry. If three or more plates are used per
sample, use average number of colonies times the reciprocal of the dilution (see Section 9610B)
to give colony count. If no plates have colonies, record count as < 1 for the highest dilution. If
there are more than 150/plate, record as Too Numerous To Count (TNTC) but indicate a count of
> 150 for the appropriate dilution. If colonies are crowded and overlapping with spreaders,
record as ‘‘obscured’’ (OBSC) and repeat analysis with higher dilution or earlier observations.
4. Reference
1. EL-SHAARAWI, A., A.A. QURESHI & B.J. DUTKA. 1977. Study of microbiological and
physical parameters in Lake Ontario adjacent to the Niagara River. J. Great Lakes Res.
3:196.
9610 D.
Membrane Filter Technique
For general information on the membrane filter technique and apparatus needed see Section
9222. However, except for comparisons of different manufacturers’ membranes, no critical tests
have been reported for membrane filters for fungal isolation efficiency. Media components, pH
levels, and antibiotics have been used in routine plating procedures. It appears that the reported
procedures are satisfactory.
1. Samples
See Section 9610B.1 and Section 9060A.
2. Media
Use unmodified or modified aureomycin®-rose bengal-glucose-peptone agar (MARGPA) or
modified streptomycin-terramycin®-malt extract agar (MSTMEA).1 These media are prepared
identically to the unmodified media described in 9610C.2e and f except that the concentration of
each antibiotic is increased from 70 mg/L to 200 mg/L. Dispense media in portions of 5 to 7 mL
in glass or plastic petri dishes (60 × 15 mm); plastic dishes with tight-fitting lids are preferred.
3. Procedure
a. Preparation and dilution: See Section 9215A.5 and Section 9610B.3a. Select dilutions to
yield 20 to 100 colonies per membrane.
b. Filtration: Filter appropriate volumes of well-shaken sample or dilution, in triplicate,
through membrane filters with pore diameter of 0.45 or 0.8 µm. See Section 9222.
c. Incubation: Transfer filters to dishes, invert dishes or not, and incubate at 15°C for 5 d in a
humid atmosphere. Alternatively incubate at 20°C for 3 d, or longer depending on fungi present.
d. Counting and recording: Using a binocular dissecting microscope at a magnification of
10×, count all colonies on each selected plate. If counting must be delayed temporarily, hold
plates at 4°C for not longer than 24 h. Ideal plates have 20 to 80 colonies per filter. See Section
9610C.
4. Reference
1. QURESHI, A.A. & B.J. DUTKA. 1978. Comparison of various brands of membrane filter
for their ability to recover fungi from water. Appl. Environ. Microbiol. 32:445.
9610 E.
Technique for Yeasts
Of the total number of fungal colonies obtained from polluted waters, as many as 50% may
be yeast colonies. Solid media such as those described above do not permit growth of all yeasts;
thus, a quantitative enrichment technique may be useful in addition to the plate count (see also
Fungi Pathogenic to Humans, Section 9610H).
1. Media
For enrichment, use yeast nitrogen base-glucose broth; for isolation, use yeast extract-malt
extract-glucose agar or diamalt agar.
a. Yeast nitrogen base-glucose broth: Dissolve 13.4 g yeast nitrogen base in 1 L
reagent-grade water; sterilize by filtration. Prepare 500 mL each of 2% and 40% aqueous
glucose solutions and sterilize separately by filtration. To make final medium, aseptically add to
a sterile 250-mL erlenmeyer flask 25 mL yeast nitrogen base solution and 25 mL of either 2% or
40% glucose solutions to make 1% or 20% final glucose concentrations. Stopper flask with a
gauze-wrapped cotton stopper and store until used.
b. Yeast extract-malt extract-glucose agar: See Section 9610B.2c.
c. Diamalt agar: See Section 9610B.2d.
2. Procedure
a. Sample preparation and dilution: Prepare as directed in Section 9610B.
b. Enrichment: In 250-mL erlenmeyer flasks prepare one flask each of yeast nitrogen base
medium containing 1% and 20% glucose. Inoculate with 1 mL of appropriate sample dilution
and incubate at room temperature on a rotary shaker operating at 120 to 150 oscillations/min for
at least 64 h. Shaken cultures are necessary to prevent overgrowth by filamentous fungi.
c. Isolation: Remove flasks from shaker and let settle 4 to 5 h. Yeast cells, if present, will
settle to the bottom, bacteria and filamentous fungi will remain in suspension, and filamentous
fungi will float on the surface or will be attached to the glass surface at or above the meniscus.
With a nichrome wire loop remove a loopful of sediment at the sediment-supernatant interface
from a tilted flask and smear-streak on yeast extract-malt extract-glucose agar. Use three plates
per flask. Incubate at room temperature but out of direct sunlight for 2 to 3 d. It is not necessary
to invert dishes. To obtain pure cultures, pick from reasonably isolated colonies and restreak on
the same medium or on diamalt agar plates. Obtain pure cultures of as many different colonies as
can be recognized.
d. Counting: It is impossible to obtain a meaningful plate count after this type of enrichment
isolation. If it is assumed that one cell in the original sample will produce one or more colonies
on the plates after enrichment, it can be stated that yeasts, or specific types of yeasts, occur at a
minimal number dependent on the highest positive dilution. The reciprocal of this dilution is the
indicated number of yeasts in the sample.
3. Bibliography
LODDER, J., ed. 1970. Th e Yeasts, A Taxonomic Study, 2nd ed. North Holland Publ. Co.,
Amsterdam.
BUCK, J.D. 1975. Distribution of aquatic yeasts—effect of inoculation temperature and
chloramphenicol concentration on isolation. Mycopathologia 56:73.
9610 F.
Zoosporic Fungi
1. Occurrence and Significance
Most fungi found in lacustrine (lake) and lotic (river) habitats that reproduce asexually by
motile, uniflagellate spores and have determinate growth of the fungal body belong to the class
Chytridiomycetes. Fungi with indeterminate growth, asexual reproduction by motile, biflagellate
spores, and sexual reproduction involving oogonia and antheridia, are members of the class
Oomycetes. A reduction in numbers of species of both classes tends to occur in polluted areas of
rivers but more species of Oomycetes than of Chytridiomycetes can be found in polluted
situations. Species of the Oomycete genera Saprolegnia (notably S. ferax) and Leptomitus appear
to be more tolerant than other forms. Bioassay studies indicate that Oomycetes are more tolerant
to zinc, cyanide, and mannitol than are Chytridiomycetes. The latter appear to be more tolerant
to treatment with surfactants than do the Oomycetes.
Some Chytridiomycetes may parasitize planktonic and other algae. In the case of epidemic
fungal infections of phytoplankton species, the activities of fungi may affect the composition of
phytoplankton communities by delaying the time of algal maxima and by reducing the
population of certain algae so that other phytoplankters will replace the infected algal
populations. In the case of nonepidemic infections, fungi may not influence algal populations;
instead, they may infect only phytoplankters during periods of decline and thus only hasten
decomposition of the algae.
Filamentous Oomycetes, particularly members of the Saprolegniaceae and Pythiaceae, are
found in virtually all types of freshwater habitats and damp-to-wet soils. Most of the nearly 250
species involved occur as saprobes on dead and decaying organic matter such as insect exuviae,
algae, and submerged vascular plant remains. A few occur as parasites of algae, aquatic
invertebrates, fish, and vascular plants; none are associated with human disease.
Rarely do any of these fungi develop in sufficient numbers to be observed or collected
directly. Consequently, various techniques have been devised for their collection and isolation.
2. Sampling and Baiting
Collect samples in sterile 35-mL plastic vials, refrigerate, and start analysis within 6 to 8 h.
Place each sample in a sterile plate (20 × 100 mm) and dilute with 10 to 15 mL sterile
reagent-grade water. Add three to four split hemp seed halves (Cannabis sativa), or whole seeds
of mustard (Brassica) or sesame (Sesamum) as bait to each culture. Incubate at 18 to 23°C and
examine daily for fungal growth on the bait. As growth becomes evident, usually within 72 h,
remove the infected bait, wash it thoroughly with water from a wash bottle, and transfer to a
fresh plate of water containing two to three halves of hemp or other seed. Genera may be
identified from spore arrangement within the sporangium and the manner in which spores are
released. Specific determination requires microscopic examination of the sexual reproductive
structures.
To collect the few naturally occurring parasites or pathogens, place the host organisms in a
plate containing sterile water and hemp seed.
3. Isolation
Although most filamentous Oomycetes can be cultivated on plain cornmeal agar, selective
media for isolating Saprolegnia from fresh water have been developed.1
Obtain axenic cultures by drawing spores into a micropipet as they emerge from the
sporangium. Less preferably, use hyphal tips, but note that several different genera and species
frequently occur on a single piece of bait. Transfer the spore suspension or hyphal tip to a plate
of cornmeal agar. When growth on the agar has occurred, remove bacteria-free hyphal tips
aseptically by cutting out a small block of agar. Transfer to fresh medium or water. If growth is
not free from contamination after one transfer, make additional transfers to insure pure cultures.
Other methods have been outlined.2
4. Dilution Plating
Make serial dilutions with sterile reagent-grade water (1:100 000 to 1:700 000) and spread 1
mL over surface of a freshly prepared cornmeal agar plate. Remove each developing colony and
transfer to water for identification. This method permits numerical estimation and determination
of composition of the Oomycete community but requires at least 10 plates.
5. References
1. HO, H.H. 1975. Selective media for the isolation of Saprolegnia spp. from fresh water.
2. SEYMOUR, R.L. 1970. The Genus Saprolegnia. Nova Hedwigia Beihefte 19:1.
6. Bibliography
WILLOUGHBY, L.G. 1962. The occurrence and distribution of reproductive spores of
Saprolegniales in fresh water. J. Ecol. 50:733.
9610 G.
Aquatic Hyphomycetes
Freshwater Hyphomycetes are a very specialized group of conidial fungi that usually occur
on partially decayed, submerged leaves and occasionally wood of angiosperms. The mycelium,
which is branched and septate, ramifies through the leaf tissue, especially in petioles and veins.
The conidiophores project into the water and the conidia that usually develop are liberated under
water. Mature conidia also can be found in the surface foam of most rivers, streams, and lakes.
The conidia of the majority of these fungi are hyaline, thin-walled, and either tetraradiately
branched, that is, with four divergent arms, or sigmoid (S-shaped) with the curvature in more
than one plane. A special feature of the conidia is that while suspended in water, even for long
periods, they do not germinate. However, if they come to rest on a solid surface, germ tubes are
produced within a few hours. The size and morphology of these spores make them potentially
more prominent in plankton analysis work than the spores of other fungi.
Ecological investigations of freshwater Hyphomycetes have been limited to substrate,
habitat, dispersal, and their role in the enhancement of leaf substrates as food for aquatic
invertebrates. The most common substrates of these organisms are submerged, decaying leaves
of angiosperms such as alder (Alnus), oak (Quercus), hazelnut (Corylus), elm (Ulmus), maple
(Acer), chestnut (Castanea), blackberry (Rubus), ash (Fraxinus), and willow (Salix). Submerged
gymnosperm leaves usually are free of aquatic Hyphomycetes. The usual habitat of these fungi is
well-oxygenated water, such as alpine brooks, mountain streams, and fast-flowing rivers.
However, they also have been found in slow-running, often contaminated, rivers, stagnant or
temporary pools, melting snow, and soil. There is often an increase in the numbers of species and
individuals of aquatic Hyphomycetes from autumn until spring, with a decline between April and
June.
2. Sample Collection and Storage
For most freshwater environments, collect foam or partially decayed, submerged,
angiosperm leaves in sterile bottles. Refrigerate sample until it is examined.
3. Sample Treatment and Analysis
Wash the leaf samples in sterile distilled water and place one to three leaves in a sterile petri
dish about 1 cm deep containing sterile pond, river, or lake water. Incubate at room temperature.
Within 1 to 2 d, the mycelium and conidia develop. Conidiophores and conidia can be observed
with a dissecting microscope on any portion of a leaf surface, but most frequently are seen on
petioles and veins. When released, the conidia either remain suspended in the water or settle to
the dish bottom. Using a dissecting microscope, pick up single conidia with a micropipet.
Transfer each conidium to a microscope slide in a drop of water for identification. The conidium
may be transferred with a sterile needle to a plate of 2% malt extract agar for colony production.
Search for conidia in foam samples with a dissecting microscope and isolate single conidia as
described above. Submerge mycelial plugs from stock culture isolates of aquatic Hyphomycetes
in autoclaved pond water in deep petri dishes; conidiogenesis usually occurs within 2 to 10 d.
Conidia in all stages of development can be preserved on slides with lactophenol mounting
medium in which either acid fuchsin or cotton blue is dissolved and sealed with clear fingernail
polish. To permit good adherence of the nail polish, avoid excessive amounts of mounting
medium.
9610 H.
Fungi Pathogenic to Humans
Routine isolations of fungi from polluted streams and wastewater treatment plants usually
have yielded relatively few species pathogenic to human and other higher animals. Geotrichum
candidum, an arthroconidium-producing fungus, for which there is, in the United States,
presumptive evidence of an association with disease, is isolated almost universally. When its
teleomorphic stage (an ascus) develops, it is known as Endomyces candidus. Rhinocladiella
mansonii, now called Exophiala mansonii, a causal agent of one form of chromomycosis, usually
in the tropics, is equally widespread. It has been listed also as Phialophora jeanselmei and
Trichosporium heteromorphum. Aspergillus fumigatus, a causal agent of pulmonary
aspergillosis, is commonly isolated. Pseudallescheria (Petriellidium, Allescheria) boydii is a
causal agent of eumycotic mycetomas and other eumycotic conditions grouped1 under the
heading ‘‘Pseudallescheriasis.’’ Infection may follow from a puncture wound with contaminated
materials or breathing such contaminated materials as sprays at wastewater treatment plants or
contaminated air. It usually is recovered in its anamorphic state, Scedosporium (Monosporium)
apiospermum. The presence of these fungi in stream water probably represents soil runoff
because virtually all zoopathogenic fungi exist saprobically in soil as their natural reservoir.
Other zoopathogenic fungi occasionally are recovered in low frequencies from streams, polluted
or not. Another fungus, the yeast Candida albicans, can be recovered in varying numbers from
wastewater treatment plant effluents, streams receiving such effluents, and recreational waters.
In humans this fungus is usually a commensal organism, like Geotrichum candidum, coexisting
in harmony with its host organism; up to 80% of normal, healthy adults have detectable levels of
C. albicans in their feces, while about 35% harbor it in their oral cavities in the absence of any
overt disease. A very large proportion of the female population has vaginal candidiasis in
varying degrees of severity. The presence of C. albicans in raw wastewater, wastewater
treatment plant effluent, or contaminated water is not surprising. C. albicans has been isolated
from these habitats on routine media heavily supplemented with antibacterial drugs, but not on
media or with techniques described in Section 9610B or Section 9610E. It also has been isolated
from estuarine and marine habitats on a maltose-yeast nitrogen
base-chloramphenicol-cycloheximide medium.
2. Identification of C. albicans
C. albicans can be identified among the white and pink yeasts growing on an 0.8-µm black
membrane filter on maltose-yeast nitrogen base-chloramphenicol-cycloheximide medium. From
each colony, inoculate a 0.5-mL portion of calf or human blood serum, incubate at 37°C for 2 to
3 h, transfer a drop or two to a slide, and examine microscopically for the production of germ
tubes from a majority of the cells. Of the white yeasts, only C. albicans produces these short
hyphae from the parent cell within 2 to 3 h incubation.2
3. References
1. RIPPON, J. 1982. Medical Mycology. W.B. Saunders Co., Philadelphia, Pa.
2. BUCK, J.D. & B.M. BUBACIS. 1978. Membrane filter procedure for enumeration of
Candida albicans in natural waters. Appl. Environ. Microbiol. 35:237.
4. Bibliography
PAGAN, E.F. 1970. Isolation of human pathogenic fungi from river water. Ph.D. dissertation,
Botany Dep., Ohio State Univ., Columbus.
9711
PATHOGENIC PROTOZOA*#(110)
9711 A.
Introduction
1. Significance
Pathogenic intestinal protozoa are significant problems in drinking water supplies. These
organisms cause diarrhea or gastroenteritis of varying severity; numerous outbreaks have
occurred. During the 1970s, waterborne outbreaks due to Giardia lamblia were noted with
increasing frequency, especially in communities using unfiltered surface water sources.1 By the
mid 1980s, waterborne outbreaks due to Cryptosporidium parvum began to appear, and in 1993,
this organism was responsible for the largest waterborne outbreak in U.S. history.2 Recreational
waterborne outbreaks also have been reported.3 Cryptosporidium can cause a severe diarrhea
that is self-limiting in immunocompetent individuals but may be prolonged and life-threatening
in the immunocompromised.4
Microscopic antibody-based methods generally have been used for detecting and quantifying
the environmentally resistant cyst stages of the pathogenic intestinal protozoa in water samples.
Using these methods, surveys have demonstrated the wide distribution and occurrence of
Giardia cysts and Cryptosporidium oocysts in raw and treated water supplies.5-7 Recently,
another protozoan intestinal pathogen, Cyclospora cayetanensis, has been associated with
waterborne8 and foodborne illness.9 Another group of protozoan organisms, Microsporidia,
while not yet associated with a waterborne outbreak, appear to be widely distributed in nature.
Microsporidia have caused intestinal illness and conjunctivitis, primarily in the
immunocompromised.10 Although Cyclospora have been found in drinking water,11 the methods
for detecting both Cyclospora and Microsporidia in environmental samples are developmental.
Giardia and Cryptosporidium occur in domestic and feral animals as well as in humans. The
environment may become contaminated through direct deposit of human and animal feces or
through sewage and wastewater discharges to receiving water. Ingestion of water containing
these organisms may cause disease.
2. References
1. CRAUN, G.F. 1990. Waterborne giardiasis. In E.A. Meyer, ed. Giardiasis. Elsevier,
New York, N.Y.
2. MACKENZIE, W.R., N.J. HOXIE, M.E. PROCTOR, M.S. GRADUS, K.A. BLAIR, D.E. PETERSON,
J.J. KAZMIERCZAK, D.G. ADDISS, K.R. FOX, J.B. ROSE & J.P. DAVIS. 1994. A massive
outbreak in Milwaukee of Cryptosporidium infection transmitted through the public
water supply. N. England J. Med. 331(3):161.
3. ROSE, J.B., J.T. LISLE & M. LECHEVALLIER. 1997. Waterborne cryptosporidiosis:
4.
5.
6.
7.
8.
9.
10.
11.
incidence, outbreaks and treatment strategies. In R. Fayer, ed. Cryptosporidium and
Cryptosporidiosis. CRC Press, Boca Raton, Fla.
UNGAR, B.L.P. 1994. Cryptosporidium and cryptosporidiosis. In S. Broder, T.C.
Merigan, Jr. & D. Bolognesi, eds. Textbook of AIDS Medicine. Williams & Wilkins,
Baltimore, Md.
LECHEVALLIER, M.W., W.D. NORTON & R.G. LEE. 1991. Occurrence of Giardia and
Cryptosporidium spp. in surface water supplies. Appl. Environ. Microbiol. 57:2610.
LECHEVALLIER, M.W., W.D. NORTON & R.G. LEE. 1991. Giardia and Cryptosporidium
spp. in filtered drinking water supplies. Appl. Environ. Microbiol. 57:2617.
ROSE, J.B., C.P. GERBA & W. JAKUBOWSKI. 1991. Survey of potable water supplies for
Cryptosporidium and Giardia. Environ. Sci. Technol. 25:1393.
SOAVE, R. & W.D. JOHNSON, JR. 1995. Cyclospora: conquest of an emerging pathogen.
Lancet 345:667.
CENTERS FOR DISEASE CONTROL AND PREVENTION. 1996. Update: outbreaks of
Cyclospora cayetanensis infection—United States and Canada, 1996. Morbid. Mortal.
Weekly Rep. 45:611.
WEBER, R., R.T. BRYAN, D.A. SCHWARTZ & R.L. OWEN. 1994. Human microsporidial
infection. Clin. Microbiol. Rev. 7:426.
RABOLD, J.G., C.W. HOGE, D.R. SHLIM, C. KEFFORD, R. RAJAH & P. ECHEVERRIA. 1994.
Cyclospora outbreak associated with chlorinated drinking water. Lancet 344:1360.
9711 B.
Detection and Enumeration Methods
Methods for the simultaneous detection and enumeration of Giardia cysts and
Cryptosporidium oocysts in water have appeared in previous editions of Standard Methods.
These methods were developed to assist in the investigation of suspected waterborne disease
outbreaks. They were applied subsequently to occurrence and distribution studies and to the
determination of drinking water treatment effectiveness. The need for quantitative methods for
regulatory purposes1 resulted in method evaluation studies that underscored the deficiencies of
the existing antibody-based immunofluorescence methods.2 These deficiencies included
requiring analysts with a high degree of experience, lengthy analysis time, high expense, lack of
specificity, erratic efficiency, low precision, and difficulty in determining viability.3
The regulatory requirements for precise, sensitive, and quantitative methods for these
organisms have stimulated interest and research on methods for their detection. No method is
included in this edition of Standard Methods because the methods for Giardia and
Cryptosporidium are evolving rapidly. A regulatory method based on the widely used
immunofluorescence assay is available.4 In addition, a draft method that addresses some of the
limitations of the regulatory method has been published.5 The reader with a need to analyze
water samples for pathogenic intestinal protozoa is advised to consult the current literature for
the latest methodology.
2. References
1. U.S. ENVIRONMENTAL PROTECTION AGENCY. 1996. Monitoring requirements for
public drinking water supplies: Cryptosporidium, Giardia, viruses, disinfection
byproducts, water treatment plant data and other information requirements. 40 CFR
Part 141; Federal Register 61: 24353.
2. Development of Performance Evaluation (PE) Sample Preparation Protocols for
Giardia Cysts and Cryptosporidium Oocysts. 1996. EPA Contract No. 68-C3-0365,
available from EPA Water Docket, 202/ 260-3027.
3. JAKUBOWSKI, W., S. BOUTROS, W. FABER, R. FAYER, W. GHIORSE, M. LECHEVALLIER, J.
ROSE, S. SCHAUB, A. SINGH & M. STEWART. 1996. Status of environmental methods for
Cryptosporidium. J. Amer. Water Works Assoc. 88(9):107.
4. FOUT, G.S., F.W. SCHAEFER, J.W. MESSER, D.R. JAHLING & R.E. STETLER. 1996. ICR
Microbial Laboratory Manual. EPA 600/R-95-178, U.S. Environmental Protection
Agency, Washington, D.C. [available online]
5. U.S. ENVIRONMENTAL PROTECTION AGENCY. 1997. Method 1622: Cryptosporidium in
Water by Filtration/IMS/FA (draft). EPA-821/R-97-023, U.S. Environmental
Protection Agency, Washington, D.C. [available online]
Figures
Figure 9020:1. Frequency curve (positively skewed distribution).
Figure 9215:1. Preparation of dilutions.
Figure 9215:2. Drying weight loss of 15-mL agar plates stored separately, inverted with
lids on. Source: Unpublished data. Water Purification Lab., Chicago Dep. Water.
Figure 9215:3. Weight loss of 25-mL agar plates (100 × 15 mm) dried separately in a
laminar-flow hood at room temperature (24 to 26°C), relative humidity (30 to 33%),
and air velocity 0.6 m/s. Source: Unpublished data. Alberta Environmental Centre,
Vegreville, Alta.
Figure 9221:1. Schematic outline of presumptive, confirmed, and completed phases for
total coliform detection.
Figure 9240:1. Filaments of Crenothrix polyspora showing variation of size and shape of
cells within the sheath. Note especially the multiple small round cells, or ‘‘conidia,’’ found
in one of the filaments. This distinctive feature is the reason for the name polyspora. Young
growing colonies usually are not encrusted with iron or manganese. Older colonies often
exhibit empty sheaths that are heavily encrusted. Cells may vary considerably in size:
Rod-shaped cells average 1.2 to 2.0 µm in width by 2.4 to 5.6 µm in length; coccoid cells of
‘‘conidia’’ average 0.6 µm in diameter.
Figure 9240:2. Filaments of Sphaerotilus natans, showing cells within the filaments and
some free ‘‘swarmer’’ cells. Filaments show false branching and areas devoid of cells.
Individual cells within the sheath may vary in size, averaging 0.6 to 2.4 µm in width by 1.0
to 12.0 µm in length; most strains are 1.1 to 1.6 µm wide by 2.0 to 4.0 µm long.
Figure 9240:3. Laboratory culture of Gallionella ferruginea, showing cells, stalks excreted
by cells, and branching of stalks where cells have divided. A precipitate of inorganic iron
on and around the stalks often blurs the outlines. Cells at tip of stalk average 0.4 to 0.6 µm in
width by 0.7 to 1.1 µm in length.
Figure 9240:4. Mixture of fragments of stalks of Gallionella ferruginea and inorganic
iron-manganese precipitate found in natural samples from wells. Fragmented stalks
appear golden yellow to orange when examined under the microscope.
Figure 9240:5. Single-celled iron bacterium Siderocapsa treubii. Cells are surrounded by a
deposit of ferric hydrate. Individual cells average 0.4 to 1.5 µm in width by 0.8 to 2.5 µm in
length.
Figure 9240:6. Photosynthetic purple sulfur bacteria. Large masses of cells have
brown-orange to purple color—may appear chalky if there is a large amount of sulfur within
the cells. Left: cells of Chromatium okenii (5.0 to 6.5 µm wide by 8 to 15 µm long)
containing sulfur globules. Right: Thiospirillum jenense (3.5 to 4.5 µm wide by 30 to 40 µm
long); cell contains sulfur globules and polar flagellum is visible.
Figure 9240:7. Colorless filamentous sulfur bacteria: Beggiatoa alba trichomes, containing
globules of sulfur. Filaments are composed of a linear series of individual rod-shaped cells
that may be visible when not obscured by light reflecting from sulfur granules. Trichomes are
2 to 15 µm in diameter and may be up to 1500 µm long; individual cells, if visible, are 4.0 to
16.0 µm long.
Figure 9240:8. Thiothrix nivea in a rosette formation. Filaments are 1 to 1.5 µm in diameter
and of varying length. Individual cells are 1 to 1.5 µm wide and 2 to 4 µm long.
Figure 9240:9. Colorless filamentous sulfur bacteria: portion of a colony, showing
branching of the mucoid filament, identified as Thiodendron mucosum.1 Because the
name Thiodendron previously had been used in bacterial taxonomy, its use here is
illegitimate and this organism remains unnamed. Individual cells (1.0 to 2.5 µm wide by 3 to
9 µm long) have been found within the jelly-like material of the filaments. The long axis of
the cells runs parallel to the long axis of the filaments.
Figure 9240:10. Colorless nonfilamentous sulfur bacteria: dividing cell of Thiovolum majus,
containing sulfur globules. Cells may measure 9 to 17 µm in width by 11 to 18 µm in
length and are generally found in nature in a marine littoral zone rich in organic matter and
hydrogen sulfide.
Figure 9250:1. Bacterial colonies—typical colony type vs. actinomycete colony type, 50×.
Left: A typical bacterial colony characterized by a smooth mucoid appearance and a
relatively distinct smooth border. Right: An actinomycete colony characterized by the mass
of branching filaments that result in the fuzzy appearance of its border and by the dull
powdery appearance of the spore-laden, aerial hyphae.
Figure 9510:1. Two-stage microporous filter adsorption-elution method for concentrating
viruses from large volumes of water with electronegative filters.
Figure 9510:2. Schematic of apparatus for first-stage concentration with negatively
charged filters.
Tables
TABLE 9020:I. KEY QUALITY CONTROL PRACTICES
Further
Information in
Section 9020B,
¶
Item
Action
Frequency
Reagent water
Bench surface
Air in workplace
Thermometers
Balances and weights
Balances
pH meter
Monitor quality
Monitor for contamination
Monitor bacterial density
Check accuracy
Check accuracy
Service and recalibrate
Standardize
Check against another meter
Check volume accuracy
See Table 9020:II
Weekly
2e
Monthly
2e
Semiannually
3a
Monthly
3b
Annually
3b
Each use
3c
Monthly
3c
Each use
3f
Check performance
Check performance
Check temperature
Check temperature
Defrost
Check for leaks and surface
scratches
Test with UV meter
Monthly
Each use
Daily
Daily
Semiannually
Each use
3g
3h
3i
3j
3j
3k
Quarterly
3l
Media-dispensing
apparatus
Hot-air oven
Autoclave
Refrigerator
Freezer
Membrane filtration
equipment
UV lamps
Further
Information in
Section 9020B,
¶
Item
Action
Frequency
Biohazard hood
Monitor air and UV lamps
Inspect for airflow
Check temperature
Clean optics and stage
Inspect for cleanliness, chips,
and etching
Check pH
Conduct inhibitory residue test
Check pH and volume
Check pH and appearance
Check performance
Perform duplicate analyses
Repeat counts
Monthly
Quarterly
Twice daily
Each use
Each use
3m
3m
3n and o
3p
4a
Each batch
Annually
Each use
Each use
Weekly
Weekly
Monthly
4a1)
4a2)
4c
4i1)
4i2)
8a4)
8a2)
Incubator
Microscope
Glassware
Dilution water bottles
Media
Autoclave
Plate counts
TABLE 9020:II. QUALITY OF REAGENT WATER USED IN MICROBIOLOGY TESTING
Test
Chemical tests:
Conductivity
pH
Total organic carbon
Heavy metals, single (Cd, Cr,
Cu, Ni, Pb, and Zn)
Heavy metals, total
Ammonia/organic nitrogen
Total chlorine residual
Bacteriological tests:
Heterotrophic plate count
(See Section 9215)
Use test (see 4e)
Monitoring Frequency
Maximum Acceptable Limit
Continuously or with
each use
With each use
Monthly
Annually*
>0.5 megohms resistance
or <2 µmhos/cm at 25°C
5.5–7.5
<1.0 mg/L
<0.05 mg/L
Annually*
Monthly
Monthly or with each use
<0.10 mg/L
<0.10 mg/L
<0.01 mg/L
Monthly
< 1000 CFU/mL
Quarterly and for a new
source
Student’s t ≤ 2.78
* Or more frequently if there is a problem.
TABLE 9020:III. TIME AND TEMPERATURE FOR AUTOCLAVE STERILIZATION
Material
Time at 121°C*
Membrane filters and pads
Carbohydrate-containing media (lauryl
tryptose, BGB broth, etc.)
Contaminated materials and discarded
cultures
Membrane filter assemblies (wrapped),
sample collection bottles (empty)
Buffered dilution water, 99 mL in
screw-cap bottle
Rinse water, volume > 100 mL
10 min
12–15 min
30 min
15 min
15 min
Adjust for
volume
* Except for media, times are guidelines; check for sterility.
TABLE 9020:IV. HOLDING TIMES FOR PREPARED MEDIA
Medium
Holding Time
Membrane filter (MF) broth in screw-cap flasks
at 4°C
MF agar in plates with tight-fitting covers at 4°C
Agar or broth in loose-cap tubes at 4°C
Agar or broth in tightly closed screw-cap tubes or
other sealed containers
Poured agar plates with loose-fitting covers in
sealed plastic bags at 4°C
Large volume of agar in tightly closed screw-cap
flask or bottle at 4°C
96 h
2 weeks
2 weeks
3 months
2 weeks
3 months
TABLE 9020:V. CONTROL CULTURES FOR MICROBIOLOGICAL TESTS
Control Culture
Group
Total coliforms
Positive
Escherichia coli
Negative
Staphylococcus aureus
Control Culture
Group
Positive
Negative
Fecal coliforms
Enterobacter aerogenes
E. coli
Escherichia coli
Fecal streptococci
E. coli
Enterococcus faecalis
Enterococci
S. faecalis
Pseudomonas sp.
E. aerogenes
Streptococcus faecalis
E. aerogenes
Staphylococcus aureus
E. coli
S. mitis/salivarius
TABLE 9020:VI. CALCULATION OF PRECISION CRITERION
Duplicate
Analyses
Sample
No.
Logarithms
of Counts
Range of
Logarithms (Rlog)
D1
D2
L1
L2
(L1 − L2)
1
2
3
89
38
58
71
34
67
1.9494
1.5798
1.7634
1.8513
1.5315
1.8261
0.0981
0.0483
0.0627
⋅
⋅
⋅
⋅
⋅
⋅
⋅
⋅
⋅
⋅
⋅
⋅
⋅
14
15
⋅
⋅
7
110
6
121
⋅
0.8451
2.0414
⋅
0.7782
2.0828
⋅
0.0669
0.0414
Calculations:
1) ∑ of Rlog = 0.0981 + 0.0483 + 0.0627 + . . . + 0.0669 + 0.0414= 0.718 89
3) Precision criterion = 3.27 î = 3.27 (0.0479) = 0.1566
TABLE 9020:VII. DAILY CHECKS ON PRECISION OF DUPLICATE COUNTS*
Duplicate
Analyses
Logarithms
of Counts
TABLE 9020:VII. DAILY CHECKS ON PRECISION OF DUPLICATE COUNTS*
Duplicate
Analyses
Date of
Analysis
8/29
8/30
8/31
D1
Logarithms
of Counts
L1
L2
Range of
Logarithms
Acceptance
of Range†
1.8513
2.0414
1.8633
1.8129
2.0828
1.6990
0.0383
0.0414
0.1643
A
A
U
D2
71
65
110 121
73
50
* Precision criterion = (3.27î) = 0.1566.
† A = acceptable; U = unacceptable.
TABLE 9020:VIII. COLIFORM COUNTS AND THEIR LOGARITHMS
MPN
Coliform Count
No./100 mL
log MPN
11
27
36
48
80
85
120
130
136
161
317
601
760
1020
3100
1.041
1.431
1.556
1.681
1.903
1.929
2.079
2.114
2.134
2.207
2.501
2.779
2.881
3.009
3.491
x = 442
xg = antilog 2.1825 = 152
TABLE 9020:IX. COMPARISON OF FREQUENCY OF MPN DATA
TABLE 9020:IX. COMPARISON OF FREQUENCY OF MPN DATA
Class Interval
Frequency (MPN)
0 to 400
400 to 800
800 to 1200
1200 to 1600
1600 to 2000
2000 to 2400
2400 to 2800
2800 to 3200
11
2
1
0
0
0
0
0
TABLE 9020:X. COMPARISON OF FREQUENCY OF LOG MPN DATA
Class Interval
Frequency
(log MPN)
1.000 to 1.300
1.300 to 1.600
1.600 to 1.900
1.900 to 2.200
2.200 to 2.500
2.500 to 2.800
2.800 to 3.100
3.100 to 3.400
3.400 to 3.700
1
2
1
5
1
2
2
0
1
TABLE 9211:I. SPECIAL RAPID TECHNIQUES
Microbial Group
Nonspecific microflora
Rapid Method
Bioluminescence
Chemiluminescence
Impedance
Colorimetric
Epifluorescence/fluorometric
Test
Time
h
Sensitivity
cells/mL
1
1
3–12
0.02
<1–several
100 000
500 000
100 000
10 000
—
Microbial Group
Test
Time
h
Sensitivity
cells/mL
4–5
10–13
1–7
6–12
9–12
8–20
3.5–15
2–20
0.01–500 000
1 000 000
200–100 000
>50
5–130 000
0.1–>10 000 000
2
2–3
500–3000
—
Rapid Method
Fecal coliforms
Radiometric
Glutamate decarboxylase
Electrochemical
Impedance
Gas chromatographic assay
Colorimetric
Potentiometric
Gram-negative bacteria
Limulus assay
Fluorescent antibody
TABLE 9215:I. EFFECT OF TEMPERATURE OF DRYING ON WEIGHT LOSS OF 15-ML AGAR
PLATES STORED SEPARATELY*
Time for Plates to Lose 1 to 4 g of Water
(Avg. for 5 Plates)
h
Plates Inverted
with Lids On
Plates Inverted
with Lids Removed
Temp.
°C
1g
2g
3g
4g
1g
2g
3g
4g
24
37
50
60
32
17
6
4
64
35
12
8
95
51
18
12
125
67
24
16
3.7
1.7
0.7
—
7.0
3.5
1.3
—
10.5
5.3
1.9
—
14.0
7.0
2.7
—
* Referenced in Canada Centre for Inland Waters Manual, Burlington, Ont.
TABLE 9221:I. PREPARATION OF LAURYL TRYPTOSE BROTH
Inoculum
mL
Amount of
Medium in Tube
mL
Volume of
Medium + Inoculum
mL
Dehydrated Lauryl
Tryptose Broth
Required
g/L
Inoculum
mL
Amount of
Medium in Tube
mL
Volume of
Medium + Inoculum
mL
Dehydrated Lauryl
Tryptose Broth
Required
g/L
1
10
10
20
100
100
100
10 or more
10
20
10
50
35
20
11 or more
20
30
30
150
135
120
35.6
71.2
53.4
106.8
106.8
137.1
213.6
TABLE 9221:II. MPN INDEX AND 95% CONFIDENCE LIMITS FOR VARIOUS COMBINATIONS OF
POSITIVE AND NEGATIVE RESULTS WHEN FIVE 20-ML PORTIONS ARE USED
No. of Tubes
Giving Positive
Reaction Out of
5 of 20 mL Each
MPN
Index/
100 mL
0
1
2
3
4
5
<1.1
1.1
2.6
4.6
8.0
>8.0
95% Confidence
Limits
(Approximate)
Lower
Upper
0
0.05
0.3
0.8
1.7
4.0
3.0
6.3
9.6
14.7
26.4
Infinite
TABLE 9221:III. MPN INDEX AND 95% CONFIDENCE LIMITS FOR VARIOUS COMBINATIONS OF
POSITIVE AND NEGATIVE RESULTS WHEN TEN 10-ML PORTIONS ARE USED
No. of Tubes
Giving Positive
Reaction Out of
10 of 10 mL Each
0
1
MPN
Index/
100 mL
<1.1
1.1
95% Confidence
Limits
(Approximate)
Lower
Upper
0
0.03
3.0
5.9
No. of Tubes
Giving Positive
Reaction Out of
10 of 10 mL Each
MPN
Index/
100 mL
2
3
4
5
6
7
8
9
10
2.2
3.6
5.1
6.9
9.2
12.0
16.1
23.0
>23.0
95% Confidence
Limits
(Approximate)
Lower
Upper
0.26
8.1
0.69 10.6
1.3
13.4
2.1
16.8
3.1
21.1
4.3
27.1
5.9
36.8
8.1
59.5
13.5 Infinite
TABLE 9221:IV. MPN INDEX AND 95% CONFIDENCE LIMITS FOR VARIOUS COMBINATIONS
OF POSITIVE RESULTS WHEN FIVE TUBES ARE USED PER DILUTION (10 ML, 1.0 ML, 0.1 ML)
95% Confidence
Limits
95% Confid
Limits
Combination
of Positives
MPN Index/
100 mL
Lower
Upper
Combination
of Positives
MPN Index/
100 mL
0-0-0
0-0-1
0-1-0
0-2-0
<2
2
2
4
—
1.0
1.0
1.0
—
10
10
13
4-2-0
4-2-1
4-3-0
4-3-1
4-4-0
22
26
27
33
34
9.0
12
12
15
16
1-0-0
1-0-1
1-1-0
1-1-1
1-2-0
2
4
4
6
6
1.0
1.0
1.0
2.0
2.0
11
15
15
18
18
5-0-0
5-0-1
5-0-2
5-1-0
5-1-1
5-1-2
23
30
40
30
50
60
9.0
10
20
10
20
30
Lower
95% Confidence
Limits
Combination
of Positives
MPN Index/
100 mL
2-0-0
2-0-1
2-1-0
2-1-1
2-2-0
2-3-0
4
7
7
9
9
12
3-0-0
3-0-1
3-1-0
3-1-1
3-2-0
3-2-1
8
11
11
14
14
17
4-0-0
4-0-1
4-1-0
4-1-1
4-1-2
Upper
Combination
of Positives
1.0
2.0
2.0
3.0
3.0
5.0
17
20
21
24
25
29
5-2-0
5-2-1
5-2-2
5-3-0
5-3-1
5-3-2
50
70
90
80
110
140
20
30
40
30
40
60
3.0
4.0
4.0
6.0
6.0
7.0
24
29
29
35
35
40
5-3-3
5-4-0
5-4-1
5-4-2
5-4-3
5-4-4
170
130
170
220
280
350
80
50
70
100
120
160
38
45
46
55
63
5-5-0
5-5-1
5-5-2
5-5-3
5-5-4
5-5-5
240
300
500
900
1600
≥1600
100
100
200
300
600
—
Lower
13
17
17
21
26
95% Confid
Limits
5.0
7.0
7.0
9.0
12
MPN Index/
100 mL
Lower
TABLE 9222:I. SUGGESTED SAMPLE VOLUMES FOR MEMBRANE FILTER TOTAL COLIFORM
TEST
Volume (X) To Be Filtered
mL
Water Source
Drinking water
Swimming pools
Wells, springs
Lakes, reservoirs
Water supply intake
Bathing beaches
100
50
10
1
0.1
X
X
X
X
X
X
X
X
X
X
X
X
X
X
0.01
0.001
0.0001
mL
Water Source
100
50
River water
Chlorinated sewage
Raw sewage
10
1
0.1
0.01
0.001
X
X
X
X
X
X
X
X
X
X
0.0001
X
TABLE 9222:II. CONFIDENCE LIMITS FOR MEMBRANE FILTER COLIFORM RESULTS USING
100-ML SAMPLE
95% Confidence Limits
Number of Coliform
Colonies Counted
Lower
Upper
0
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
0.0
0.1
0.2
0.6
1.0
1.6
2.2
2.8
3.4
4.0
4.7
5.4
6.2
6.9
7.7
8.4
9.2
9.9
10.7
11.5
12.2
3.7
5.6
7.2
8.8
10.2
11.7
13.1
14.4
15.8
17.1
18.4
19.7
21.0
22.3
23.5
24.8
26.0
27.2
28.4
29.6
30.8
TABLE 9222:III. SUGGESTED SAMPLES VOLUMES FOR MEMBRANE FILTER FECAL COLIFORM
TEST
mL
Water Source
100
50
Lakes, reservoirs
Wells, springs
Water supply intake
Natural bathing waters
Sewage treatment plant
Farm ponds, rivers
Stormwater runoff
Raw municipal sewage
Feedlot runoff
Sewage sludge
X
X
X
X
X
X
10
1
X
X
X
X
X
X
X
X
0.1
X
X
X
X
X
0.01
0.001
0.0001
X
X
X
X
X
X
X
X
X
TABLE 9223:I. COLOR CHANGES FOR VARIOUS MEDIA
Substrate
Total Coliform
Positive
ONPG-MUG Yellow
CPRG-MUG Red or
magenta
E. coli Positive
Negative Result
Blue
fluorescence
Blue
fluorescence
Colorless/no
fluorescence
Yellow/no
fluorescence
TABLE 9225:I. BIOCHEMICAL REACTIONS OF KEY SPECIES OF THE FAMILY
ENTEROBACTERIACEAE*
Perc
Species
Citrobacter diversus
Citrobacter freundii
Lactose
Fermentation
35
50
ONPG
Hydrolysis
96
95
Indole
Production
Methyl
Red
99
5
100
100
VogesProskauer
0
0
Perc
Species
Enterobacter aerogenes
Enterobacter agglomerans
Enterobacter cloacae
Escherichia coli
Escherichia coli, inactive
Escherichia fergusonii
Escherichia hermannii
Escherichia vulneris
Hafnia alvei
Klebsiella oxytoca
Klebsiella ozaenae
Klebsiella pneumoniae
Klebsiella rhinoscleromatis
Serratia fonticola
Serratia marcescens
Lactose
Fermentation
ONPG
Hydrolysis
Indole
Production
Methyl
Red
VogesProskauer
95
40
93
95
25
0
45
15
5
100
30
98
0
97
2
100
90
99
95
45
83
98
100
90
100
80
99
0
100
95
0
20
0
98
80
98
99
0
0
99
0
0
0
0
1
5
50
5
99
95
100
100
100
40
20
98
10
100
100
20
98
70
100
0
0
0
0
0
85
95
0
98
0
9
98
* Modified after Farmer, J.J. III, 1985. Clinical identification of new species and biogroups of Enterobacteriaceae. J. Clin. Microbiol.
21:46.
† Reactions that become positive after 2 d are not considered.
TABLE 9230:I. SELECTED KEY BIOCHEMICAL CHARACTERISTICS OF THE STREPTOCOCCUS
SPECIES WITHIN THE FECAL STREPTOCOCCUS AND ENTEROCOCCUS GROUPS*
Fecal Streptococcus Group
Enterococcus Group
Test
S. faecalis
S. faecium
S. avium
S. gallinarum
S. bovis
−
+
+
−
+
+
−
+
+
−
+
+
−
+
+
+
+
+
+
+
+
+
+
+
Growth at 10°C
+
+
+
+
−
Pyruvate utilization7
+
−
−
−
−
Phosphatase activity8
+
−
+
+
−
Catalase
40% Bile
Esculin6
Growth at 45°C
Growth in 6.5% NaCl
−
S
Fecal Streptococcus Group
Enterococcus Group
Test
S. faecalis
S. faecium
S. avium
S. gallinarum
S. bovis
Arginine hydrolysis8
+
+
−
−
L-Sorbose fermentation9
−
−
−d
+
−
−
Lactose fermentation8
+
+
+
+
+
n-Acetyl-β-glucoseaminidase
activity9
−d
−
−
+
−
Starch6
−
−
−
−
+
Arabinose6
−
+
+
−
−
S
* + = 90% or more of strains are positive
− = 90% or more of strains are negative
d = reactions variable
TABLE 9250:I. GENERAL MACROSCOPIC PROPERTIES OF BACTERIAL COLONIES ON SOLID
MEDIUM
Characteristic
Typical Colony Type
Actinomycete Colony Type*
Appearance
Shiny or opalescent
Texture
Degree of adherence
to solid medium
Edge of colony
Soft
Weak
When young it is composed of hyphae, b
some species these may later fragme
Substrate and surface hyphae have n
distinctive color. As the colony mature
fluffy aerial hyphae that carry spores f
and give to colonies of different speci
various colors and sometimes a chalk
appearance. Soluble pigments, either
melanin or brightly colored type, that
diffuse into the medium, also are com
Strong and leathery
Strong
Regular, continuous, and
not different from
colony as a whole
Irregular, intermittent, slightly less dense
colony as a whole, and of hyphal
appearance
* Actinomycetes are authentic bacteria by all modern criteria, except for their hyphal character and mode of spore formation.
TABLE 9260:I. SUMMARY DATA FROM WATERBORNE BACTERIAL DISEASE OUTBREAKS,
1985–94
Type of Water
Variable
Drinking water
Total outbreaks
Agent:
Shigella
Campylobacter
Salmonella
E. coli O157:H7
System:
Noncommunity
Community
Individual
Source:
Well
Lake
Spring
Cistern
Cause:
Untreated groundwater
Distribution system deficiency
Treatment deficiency
Unknown
Total outbreaks
Agent:
Pseudomonas
Shigella
Legionella
Leptospira
E. coli O157:H7
Location:
Hotel/motel
Recreational
water
Number
21
12
6
2
1
10
8
3
17
2
1
1
9
7
4
1
71
44
17
6
2
2
23
Type of Water
Variable
Number
Outdoor recreation area (surface
water)
Home
Spa or public swimming pool
Resort
Apartment complex/condominum
Source:
Whirlpool/hot tub
Lake/pond
Swimming pool
Stream
21
14
5
4
4
47
20
3
1
TABLE 9260:II. REACTIONS OF COMMON BACTERIA ON TSI AND LIA MEDIA
Organism
Shigella
TSI*
LIA*
K/A−
K/Ag+
K/A−
K/A+
A/Ag−
A/Ag+ or K/Ag+
A/Ag+
K/K−
R/A+
K/A+
A/Ag−
K/A−
Aeromonas
A/A−
K/A−
Yersinia
A/A− or K/A−
K/A−
Plesiomonas
K/A−
K/A−
Salmonella
Escherichia
Proteus
Citrobacter
Enterobacter
* Fermentation reactions = slant/butt; H2S production = + or −; K = alkaline,A = acid, R = red (deaminase reaction); g = gas
produced.
TABLE 9260:III. REACTIONS OF ENTERIC BACTERIA ON TSI AND LIA MEDIA
Organism
TSI Reactions*
LIA
Reactions*
Shigella
K/A−
K/Ag+
K/A−
K/K+
Salmonella
Organism
Escherichia
Proteus
Citrobacter
Enterobacter
TSI Reactions*
LIA
Reactions*
A/Ag−
A/Ag+ or K/Ag+
A/Ag+
K/A−
R/A+
K/A+
A/Ag−
K/A−
A/A−
K/A−
A/A− or K/A−
K/A−
A/Ag−
K/A−
Aeromonas
Yersinia
Klebsiella
* Fermentation reactions = slant/butt, H2S production = + or −, K = alkaline,A = acid, R = red (deaminase reaction), g = gas
produced.
TABLE 9260:IV. REACTIONS OF AEROMONAS AND ENTERIC BACTERIA ON KAPER’S MEDIUM
Fermentation Pattern*
Motility
H 2S
Indole
Aeromonas hydrophila
K/A
+
−
+
Klebsiella pneumoniae
A/A
−
−
Klebsiella oxytoca
A/A
−
−
−
+
Escherichia coli
K/K or K/A
K/K, K/A, A/K or A/A
−
+
+
Salmonella spp.
+ or −
+
K/K, K/N or N/N
+
−
R/K or R/A
+
+ or −
−
+
K/K, K/N or N/N
−
+
+ or −
K/K or K/A
−
+
K/K, K/N, or N/N
+
−
−
Organism
Enterobacter spp.
Proteus spp.
Yersinia enterocolitica
Citrobacter spp.
Serratia spp.
−
−
* K = alkaline; A = acid; N = neutral; R = red (deamination reaction).
TABLE 9260:V. MYCOBACTERIA OF WATERBORNE OR UNKNOWN ORIGIN
Mycobacterium
Species
Environmental
Contaminant
Reservoir
M. kansasii
M. marinum
M. simiae
Rarely
Rarely
No
Water, swine, cattle
Fish, water
Primates, possibly water
Mycobacterium
Species
Environmental
Contaminant
Reservoir
M. scrofulaceum
M. szulgai
M. avium-intracellulare
M. xenopi
M. ulcerans
M. fortuitum
M. chelonae
Possibly
No
Possibly
Possibly
No
Yes
Yes
Soil, water, foodstuffs
Unknown
Soil, water, swine, cattle, birds
Water
Unknown
Soil, water, animals, marine life
Soil, water, animals, marine life
TABLE 9260:VI. PHENOTYPIC CHARACTERISTICS OF CLINICALLY SIGNIFICANT
ENVIRONMENTAL MYCOBACTERIA*
Mycobacterium
Species
Nitrate
Reduction
Hydrolysis of
Polyoxyethylen
Sorbitan
Monooleate†
Growth
Rate
Pigmentation
Urease
M. kansasii
M. marinum
S
S
P
P
±
+
+
−
+
+
M. simiae
S
P
±
−
−
M. scrofulaceum
S
S
±
M. szulgai
M. xenopi
S
S
S/P
S
+
−
+
−
±
−
−
−
M. avium-intracellulare
S
N
−
−
M. ulcerans
S
N
−
±
M. fortuitum
M. chelonae
R
R
N
N
+
+
−
+
−
±
±
−
* S = slow (3 to 8 weeks), R = rapid (7 d or longer), P = photochromogenic, S = scotochromogenic, N = nonphotochromogenic, S/P
= scotochromogenic at 37°C andphotochromogenic at 24°C.
† Tween 80.
10010
INTRODUCTION*#(111)
Physical and chemical characteristics of water bodies affect the abundance, species
composition, stability, productivity, and physiological condition of aquatic organism
populations. Biological methods used for assessing water quality include the collection,
counting, and identification of aquatic organisms; biomass measurements; measurements of
metabolic activity rates; measurements of the toxicity, bioconcentration, and bioaccumulation of
pollutants; and processing and interpretation of biological data.
Information from these methods may serve one or more of the following purposes:
1. To explain the cause of color, turbidity, odor, taste, or visible particulates in water;
2. To aid in the interpretation of chemical analyses, for example, in relating the presence or
absence of certain biological forms to oxygen deficiency or supersaturation in natural waters;
3. To identify the source of a water that is mixing with another water;
4. To explain the clogging of pipes, screens, or filters, and to aid in the design and operation
of water and wastewater treatment plants;
5. To determine optimum times for treatment of surface water with algicides and to monitor
treatment effectiveness;
6. To determine the effectiveness of drinking water treatment stages and to aid in
determining effective chlorine dosage within a water treatment plant;
7. To identify the nature, extent, and biological effects of pollution;
8. To indicate the progress of self-purification in bodies of water;
9. To aid in determining the condition and effectiveness of unit processes and biological
wastewater treatment methods in a wastewater treatment plant;
10. To document short- and long-term variability in water quality caused by natural
phenomena and/or human activities;
11. To provide data on the status of an aquatic system on a regular basis;
12. To correlate the biological mass or components with water chemistry or conditions.
The specific nature of a problem and the reasons for collecting samples will dictate which
communities of aquatic organisms will be examined and which sampling and analytical
techniques will be used.
The following communities of aquatic organisms are considered in specific sections that
follow:
1. PLANKTON (Section 10200): A community of plants (phytoplankton) and animals
(zooplankton), usually drifting or suspended in water, nonmotile or insufficiently motile to
overcome transport by currents. In fresh water they generally are small or microscopic in size; in
the marine or estuarine environment, larger forms are observed more frequently.
2. PERIPHYTON (Section 10300): A community of microscopic plants and animals associated
with the surfaces of submersed objects. Some are attached, some move about. Many of the
protozoa and other minute invertebrates and algae found in the plankton also occur in the
periphyton.
3. MACROPHYTON (Section 10400): The larger plants of all types. They are sometimes
attached to the bottom (benthic), sometimes free-floating, sometimes totally submersed, and
sometimes partly emergent. Complex types usually have true roots, stems, and leaves; the
macroalgae are simpler but may have stem- and leaf-like structures.
4. MACROINVERTEBRATES (Section 10500): The invertebrates defined here are those
retained by the US Standard No. 30 sieve. They are generally bottom-dwelling organisms
(benthos).
5. FISH (Section 10600): Vertebrates of diverse morphology, ecology, and behavior,
inhabiting (and generally limited to) aquatic systems. They have fins and gills.
6. AMPHIBIANS, AQUATIC REPTILES, BIRDS, AND MAMMALS: These vertebrates also may be
affected directly or indirectly by spills or other discharges of pollutants and may be useful in
monitoring the presence of toxic substances or long-term changes in water quality. Discussion of
these organisms is not included.
Large numbers of bacteria and fungi are present in the plankton and periphyton and
constitute an essential element of the total aquatic ecosystem. Although their interactions with
living and dead organic matter profoundly affect the larger aquatic organisms, techniques for
their investigation are not included herein (see Part 9000).
Field observations are indispensable for meaningful biological interpretations, but many
biological factors cannot be evaluated directly in the field. These must be analyzed as field data
or field samples within the laboratory. Because the significance of the analytical result depends
upon the representativeness of the sample, attention is given to field methods as well as to
associated laboratory procedures.
Before sampling begins, clearly define study objectives. For example, the frequency of a
repetitive sampling program may vary from hourly, for a detailed study of diel variability, to
every third month (quarterly) for a general assessment of seasonal conditions, depending on
objectives. The scope of the study must be adjusted to limitations in personnel, time, and budget.
Before the development of a study plan, examine historic data for the study area and conduct a
literature search of work by previous investigators.
Whenever practicable, biologists should collect their own samples. Much of the value of an
experienced biologist lies in personal observations of conditions in the field and in the ability to
recognize signs of environmental changes as reflected in the various aquatic communities.
The primary orientation of Part 10000 is toward field collection and associated laboratory
analyses to aid in determining the status of aquatic communities under field conditions and to aid
in interpreting the influence of past and present environmental conditions. The methods selected
are necessary for the appraisal of water quality. Principal emphasis is on methods and
equipment, rather than on interpretation or application of results. The complex interrelationships
existing in an aquatic environment often require many different field and laboratory procedures;
consequently, frequent cross-references between sections have been made.
Many other types of studies may be, and are being, conducted that are oriented more toward
laboratory research. Such laboratory studies will develop further basic knowledge of community
and/ or organism responses under controlled conditions and will aid in predicting effects of
future changes in environmental conditions on the aquatic communities. However, such studies
are not within the scope of this book.
10200
PLANKTON*#(112)
10200 A.
Introduction
The term ‘‘plankton’’ refers to those microscopic aquatic forms having little or no resistance
to currents and living free-floating and suspended in natural waters. Planktonic plants,
‘‘phytoplankton,’’ and planktonic animals, ‘‘zooplankton,’’ are covered in this section. The
phytoplankton (microscopic algae) occur as unicellular, colonial, or filamentous forms. Many are
photosynthetic and are grazed upon by zooplankton and other aquatic organisms. Other
organisms occurring in the same environment are dealt with elsewhere: zoosporic fungi in
Section 9610F; aquatic hyphomycetes in Section 9610G; and bacteria in Part 9000. The
zooplankton in fresh water comprise principally protozoans, rotifers, cladocerans, and copepods;
a greater variety of organisms occurs in marine waters.
1. Significance
Plankton, particularly phytoplankton, long have been used as indicators of water quality.1-4
Some species flourish in highly eutrophic waters while others are very sensitive to organic
and/or chemical wastes. Some species develop noxious blooms, sometimes creating offensive
tastes and odors5 or anoxic or toxic conditions resulting in animal deaths or human illness.6 The
species assemblage of phytoplankton and zooplankton also may be useful in assessing water
quality.7
Because of their short life cycles, plankters respond quickly to environmental changes, and
hence their standing crop and species composition are more likely to indicate the quality of the
water mass in which they are found. They strongly influence certain nonbiological aspects of
water quality (such as pH, color, taste, and odor), and in a very practical sense, they are a part of
water quality. Certain taxa often are useful in determining the origin or recent history of a given
water mass. Because of their transient nature, and often patchy distribution, however, the utility
of plankters as water quality indicators may be limited. Information on plankton as indicators is
interpreted best in conjunction with concurrently collected, physicochemical and other biological
data.
Planktonic organisms predominate in ponds, lakes, and oceans. Potamoplankton develop in
large rivers with slow-moving waters that approach lentic conditions. Because their origin can be
uncertain and the duration of their exposure to pollutants unknown, plankters generally are less
valuable as water quality indicators in lotic than in lentic environments.
2. References
1. PALMER, C.M. 1969. A composite rating of algae tolerating organic pollution. J.
2.
3.
4.
5.
6.
7.
Phycol. 5:78.
PALMER, C.M. 1963. The effect of pollution on river algae. Bull. N.Y. Acad. Sci.
108:389.
RAWSON, D.S. 1956. Algal indicators of trophic lake types. Limnol. Oceanogr. 1:18.
STOERMER, E.F. & J.J. YANG. 1969. Plankton Diatom Assemblages in Lake Michigan.
Spec. Rep. No. 47, Great Lakes Research Div., Univ. Michigan, Ann Arbor.
PRESCOTT, G.W. 1968. The Algae: A Review. Houghton Mifflin Co., Boston, Mass.
CARMICHAEL, W., ed. 1981. The Water Environment, Algal Toxins and Health.
Plenum Press, New York, N.Y.
GANNON, J.E. & R.S. STEMBERGER. 1978. Zooplankton (especially crustaceans and
rotifers) as indicators of water quality. Trans. Amer. Microsc. Soc. 97:16.
10200 B.
Sample Collection
The frequency and location of sampling is dictated by the purpose of the study.1 Locate
sampling stations as near as possible to those selected for chemical and bacteriological sampling
to insure maximum correlation of findings. Establish a sufficient number of stations in as many
locations as necessary to define adequately the kinds and quantities of plankton in the waters
studied. The physical nature of the water (standing, flowing, or tidal) will influence greatly the
selection of sampling stations. The use of sampling sites selected by previous investigators
usually will assure the availability of historical data that will lead to a better understanding of
current results and provide continuity in the study of an area.
In stream and river work, locate stations upstream and downstream from suspected pollution
sources and major tributary streams and at appropriate intervals throughout the reach under
investigation. If possible, locate stations on both sides of the river because lateral mixing of river
water may not occur for great distances downstream. In a similar manner, investigate tributary
streams suspected of being polluted but take care in the interpretation of data from a small
stream because much of the plankton may be periphytic in origin, arising from scouring of
natural substrates by the flowing water. Plankton contributions from adjacent lakes, reservoirs,
and backwater areas, as well as soil organisms carried into the stream by runoff, also can
influence data interpretation. The depth from which water is discharged from upstream stratified
reservoirs also can affect the nature of the plankton.
Because water of rivers and streams usually is well mixed vertically, subsurface sampling,
i.e., the upper meter or a composite of two or more strata, often is adequate for collection of a
representative sample. There may be problems caused by stratification due to thermal discharges
or mixing of warmer or colder waters from tributaries and reservoirs. Always sample in the main
channel of a river and avoid sloughs, inlets, or backwater areas that reflect local habitats rather
than river conditions. In rivers that are mixed vertically and horizontally, measure plankton
populations by examining periodic samples collected at midstream 0.5 to 1 m below the surface.
If it can be determined or correctly assumed that the plankton distribution is uniform and
normal, use a scheme of random sampling to accomodate statistical testing. Include both random
selection of sampling sites and transects as well as the random collection of samples at each
selected site. On the other hand, if it is known or assumed that plankton distribution is variable
or patchy, include additional sampling sites, collect composite samples, and increase sample
replication. Use appropriate statistical tests to determine population variability.
In sampling a lake or reservoir use a grid network or transect lines in combination with
random procedures. Take a sufficient number of samples to make the data meaningful. Sample a
circular lake basin at strategic points along a minimum of two perpendicular transects extending
from shore to shore; include the deepest point in the basin. Sample a long, narrow basin at
several points along a minimum of three regularly spaced parallel transects that are
perpendicular to the long axis of the basin, with the first near the inlet and the last near the outlet.
Sample a large bay along several parallel transects originating near shore and extending to the
lake proper. Because many samples are required to appraise completely the plankton
assemblage, it may be necessary to restrict sampling to strategic points, such as the vicinity of
water intakes and discharges, constrictions within the water body, and major bays that may
influence the main basin.
In lakes, reservoirs, and estuaries where plankton populations can vary with depth, collect
samples from all major depth zones or water masses. The sampling depths will be determined by
the water depth at the station, the depth of the thermocline or an isohaline, or other factors. In
shallow areas of 2 to 3 m depth, subsurface samples collected at 0.5 to 1 m may be adequate. In
deeper areas, collect samples at regular depth intervals. In estuaries sample above and below the
pyncocline. Depth intervals for sampling vary for estuaries of different sizes and depths, but use
depths representative of the vertical range. Composite sampling above and below the pyncocline
often is used. In marine sampling, the intent and scope of the study will determine the collection
extent.
Over the continental shelf, take samples at stations approximately equidistant from the shore
seaward. Take a vertical series from surface to near bottom at each station, gradually adding
more stations across the shelf. It is important to sample the entire vertical range over a
continental shelf. Benthic grab samples may be taken to collect dormant resting cells or cysts.
Beyond the shelf in pelagic waters, sample in the photic zone from the surface to the thermocline
for phytoplankton and to deeper depths for zooplankton. Sampling depths vary, but often are at
10- to 25-m intervals above the thermocline, then at 100- to 200-m intervals below the
thermocline to 1000 m, and thereafter at 500- to 1000-m intervals.
Samples usually are referred to as ‘‘surface’’ or ‘‘depth’’ (subsurface) samples. The latter are
samples taken from some stated depth, whereas surface samples may be interpreted as samples
collected as near the water surface as possible. A ‘‘skimmed’’ sample of the surface film
plankton (neuston)2 can be revealing; however, ordinarily do not include a disproportionate
quantity of surface film in a surface sample because a neustonic flora3 as well as plankton often
are trapped on top or at the surface film together with pollen, dust, and other detritus. Various
methods have been used for sampling surface organisms.
Sampling frequency depends on the intent of the study as well as the range of seasonal
fluctuations, the immediate meteorological conditions, adequacy of equipment, and availability
of personnel. Select a sampling frequency at some interval shorter than community turnover
time. This requires consideration of life-cycle length, competition, predation, flushing, and
current displacement. Frequent plankton sampling is desirable because of normal temporal
variability and migratory character of the plankton community. Daily vertical migrations occur
in response to sunlight, and random horizontal migrations or drifts are produced by winds,
shifting currents, and tides. Ideally, collect daily samples and, when possible, sample at different
times during the day and at different depths. When this is not possible, weekly, biweekly,
monthly, or even quarterly sampling still may be useful for determining major population
changes.
In river, stream, and estuarine regions subject to tidal influence, expect fluctuations in
plankton composition over a tidal cycle. A typical sampling pattern at a station within an estuary
includes a vertical series of samples taken from the surface, across the pyncocline, to near
bottom, collected at 3-h intervals, over at least two complete tidal cycles. Once a characteristic
pattern is recognized the sampling routine may be modified.
A useful series of monographs on oceanographic methodology has been published.4-7
Representative taxonomic references for estuarine and marine phytoplankton include
diatoms,8-11 dinoflagellates,12-14 coccolithophores,15 and cyanophyceae16 (cyanobacteria).
2. Sampling Procedures
Once sampling locations, depths, and frequency have been determined, prepare for field
sampling. Label sample containers with sufficient information to avoid confusion or error. On
the label indicate date, cruise number, sampling station, study area (river, lake, reservoir), type of
sample, and depth. Use waterproof labels. When possible, enclose collection vessels in a
protective container to avoid breakage. If samples are to be preserved immediately after
collection, add preservative to container before sampling. Sample size depends on type and
number of determinations to be made; the number of replicates depends on statistical design of
the study and statistical analyses selected for data interpretation. Always design a study around
an objective with a statistical approach rather than fit statistical analyses to data already
collected.
In a field record book note sample location, depth, type, time, meteorological conditions,
turbidity, water temperature, salinity, and other significant observations. Engineer’s field
notebooks with waterproof paper are very suitable. Field data are invaluable when analytical
results are interpreted and often help to explain unusual changes caused by the variable character
of the aquatic environment. Collect coincident samples for chemical analyses to help define
environmental variations having a potential effect on plankton.
a. Phytoplankton: In oligotrophic waters or where phytoplankton densities are expected to be
low collect a sample of up to 6 L. For richer, eutrophic waters collect a sample of 0.5 to 1 L.
Because of their small size, nannoplankton and picoplankton can pass through collection
nets, making nets unsuitable for most phytoplankton sampling.
For qualitative and quantitative evaluations collect whole (unfiltered and unstrained) water
samples with a water collection bottle consisting of a cylindrical tube with stoppers at each end
and a closing device. Lower the open sampler to the desired depth and close by dropping a
weight, called a messenger, which slides down the supporting wire or cord and trips the closing
mechanism. If possible, obtain composite samples from several depths or pool samples from one
depth from several casts. The most commonly used samplers that operate on this principle are
the Kemmerer,17 Van Dorn18 (Figure 10200:1), Niskin, and Nansen samplers.
Because these samplers collect whole water samples, all size classes of phytoplankton are
collected. Different size categories of phytoplankton can be separated by subsequently filtering
these whole water samples through netting of the appropriate mesh size. Select appropriate mesh
sizes for concentrating the various size categories of phytoplankton typical of the aquatic system
under study.19,20
The Van Dorn usually is the preferred sampler for standing crop, primary productivity, and
other quantitative determinations because its design offers no inhibition to free flow of water
through the cylinder. In deep-water situations, the Niskin bottle is preferred. It has the same
design as the Van Dorn sampler except that the Niskin sampler can be cast in a series on a single
line for simultaneous sampling at multiple depths with the use of auxiliary messengers. Because
the triggering devices of these samplers are very sensitive, avoid rough handling. Always lower
the sampler into the water; do not drop. Kemmerer and Van Dorn samplers have capacities of 0.5
L or more. Polyethylene or polyvinyl chloride sampling devices are preferred to metal samplers
because the latter liberate metallic ions that may contaminate the sample. Use polyethylene or
glass sample storage bottles. Metallic ion contamination can lead to significant errors when algal
assays or productivity measurements are made.
For shallow waters use the Jenkins surface mud sampler,21 one of the bottle samplers
modified so that it is held horizontally,22 or an appropriate bacteriological sampler.23
For greater speed of collection and to obtain large, accurately measured quantities of
organisms, use a pump. Diaphragm and peristaltic pumps are less damaging to organisms than
centrifugal pumps.24 Centrifugal pump impellers can damage organisms as can passage through
the hose.25 Lower a weighted hose, attached to a suction pump, to the desired depth, and pump
water to the surface. The pump is advantageous because it supplies a homogeneous sample from
a given depth or an integrated sample from the surface to a particular depth. If a centrifugal
pump is used, draw samples from the line before they reach the impeller. For samples to be
analyzed for organochlorine compounds use TFE tubing.
To examine live samples fill containers partially and store in a refrigerator or ice chest in the
dark, or preferably, hold at ambient temperature. Examine specimens promptly after collection.
If it is impossible to examine living material or if phytoplankton are to be counted later,
preserve the sample. For a sample that will be preserved, fill the container completely. The most
suitable phytoplankton preservative is Lugol’s solution, which can be used for most forms
including the naked flagellates. Unfortunately, acidic Lugol’s solution (or formalin) dissolves the
coccoliths of Coccolithophores, which are common in estuarine and marine waters.
Lugol’s solution: To preserve samples with Lugol’s solution add 0.3 mL Lugol’s solution to
100 mL sample and store in the dark. For long-term storage add 0.7 mL Lugol’s solution per 100
mL sample and buffered formaldehyde to a minimum of 2.5% final concentration after 1 h.
Prepare Lugol’s solution by dissolving 20 g potassium iodide (KI) and 10 g iodine crystals in
200 mL distilled water containing 20 mL glacial acetic acid.26 Utermohl’s27 modification of
Lugol’s solution results in a neutral or slightly alkaline solution. Prepare modified Lugol’s
solution by dissolving 10 g KI and 5 g iodine crystals in 20 mL distilled water, then adding 50
mL distilled water in which 5 g anhydrous sodium acetate has been dissolved. This allows
preservation of Coccolithophores, but would be less effective for other flagellates.
Other acceptable preservatives are:
Formalin: To preserve samples with formalin, add 40 mL buffered formalin (20 g sodium
borate, Na2B2O4, + 1 L 37% formaldehyde) to 1 L of sample immediately after collection. In
estuarine and marine collections, adjust pH to at least 7.5 with sodium borate for samples
containing Coccolithophores.
Merthiolate: To preserve samples with merthiolate add 36 mL merthiolate solution to 1 L of
sample and store in the dark. Prepare merthiolate solution by dissolving 1.0 g merthiolate, 1.5 g
sodium borate, and 1.0 mL Lugol’s solution in 1 L distilled water. Merthiolate-preserved
samples are not sterile, but can be kept effectively for 1 year, after which time formalin must be
added.28
‘‘M3’’ fixative: Prepare by dissolving 5 g KI, 10 g iodine, 50 mL glacial acetic acid, and 250
mL formalin in 1 L distilled water (dissolve the iodide in a small quantity of water to aid in
solution of the iodine). Add 20 mL fixative to 1 L sample and store in the dark.
Glutaraldehyde: Preserve samples by adding neutralized glutaraldehyde to yield a final
concentration of 1 to 2%.
Other commonly used preservatives include 95% alcohol, and 6-3-1 preservative, (6 parts
water, 3 parts 95% alcohol, and 1 part formalin). Use equal volumes of preservative and sample.
To retain color in preserved plankton, store samples in the dark or add 1 mL saturated copper
sulfate (CuSO4) solution/L.
Most preservatives distort and disrupt certain cells,29,30 especially those of delicate forms
such as Euglena, Cryptomonas, Synura, Chromulina, and Mallamonas. Lugol’s iodine solution
usually is least damaging for these phytoflagellates. To become familiar with live specimens and
preservation-caused distortions, use reference collection from biological supply houses or
consult experienced co-workers.
b. Zooplankton: The choice of sampler depends on the type of zooplankton, the kind of study
(distribution, productivity, etc.) and the body of water being investigated. Zooplankton
populations invariably are distributed in a patchy way, making both sampling and data
interpretation difficult.
For collecting microzooplankton (20 to 200 µm) such as protozoa, rotifers, and immature
microcrustacea, use the bottle samplers described for phytoplankton. The small zooplankters
usually are sufficiently abundant to yield adequate samples in 5- to 10-L bottles; however,
composite samples over depth and time are recommended. Water bottle samplers are suitable
especially for discrete-depth samples. If depth-integrated samples are desired, use pumps or nets.
The larger and more robust microzooplankters (e.g., loricate forms and crustacea) may be
concentrated by passing the whole water through a 20-µm mesh net. If quantitative estimates of
other nonloricate, delicate forms are required, do not screen. Fix 0.5 to 5 L of whole water for
enumeration of these forms.
Bottle samplers usually are unsuitable for collecting larger zooplankton, such as mature
microcrustacea, that, unlike the smaller forms, are much less numerous and are sufficiently agile
to avoid capture. Although comparatively large water volumes, and consequently adequate
numbers of microcrustacea, can be sampled with a pump, avoidance by larger, more agile
zooplankters at the pump head can cause sampling error. Consequently, larger trap samplers or
nets are the preferred collection methods.
The Juday trap31 operates on the same principle as the water bottle samplers but is generally
larger (10 L). The larger size makes the Juday trap more suitable for collecting zooplankters,
especially larger copepods. However, it is awkward to use and its 10-L capacity is inadequate for
oligotrophic lakes or other water bodies with few zooplankters. Because it is constructed of
metal it is unsuited if heavy metals analyses are required.
The Schindler-Patalas trap32 (Figure 10200:2) usually is preferred to the Juday trap because
it is constructed of clear acrylic plastic and is transparent. It can be lowered into the water with
minimal disturbance and is suitable for collecting larger zooplankters. Models of 10- to 12-L
capacity are available but the 30-L size is preferred. It has no mechanical closing mechanism and
thus is convenient for cold-weather sampling when mechanical devices tend to malfunction. Like
the Juday trap, it can be fitted with nets of various mesh sizes, but the No. 20 mesh net is used
most often.
Plankton nets are preferred to bottles and traps for sampling where plankters are few or
where only qualitative data or a large biomass is needed for analysis. Because they were
designed originally for qualitative sampling, modifications are required for quantitative work.
The mesh size, type of material, orifice size, length, hauling method, type of tow, and volume
sampled will depend on the particular needs of the study.33,34 Type of netting and mesh size
determine filtration efficiency, clogging tendencies, velocity, drag, and the condition of the
sample after collection. Silk, formerly the common mesh material in plankton nets, is not
recommended because of shrinkage of mesh openings and rotting with age. Nylon monofilament
mesh is preferred because of its mesh size accuracy and durability. Nylon nets of different mesh
sizes still are labelled by the silk rating system: characteristics of commonly used nylon plankton
nets are listed in Table 10200:I. Finer mesh sizes clog more readily than coarser mesh; a
compromise must be made between mesh size small enough to retain desired organisms
effectively and a size large enough to preclude a serious clogging problem. If clogging occurs,
reduce its effects by decreasing the length of tow.
The maximum volume, VM, of water that can be filtered through a net during a vertical tow
can be estimated with the formula,
where:
r = radius of net orifice and
d = depth to which net is lowered.
This volume is a maximum because clogging of the net’s meshes by phytoplankton and other
particles and, for fine netting, even the netting itself can cause some water to be diverted from
the net’s path.35,36 Keep net towing distance as short as practical to alleviate clogging. If the net
has a pronounced green or brown color after towing, clogging probably has occurred.
To estimate sampling volume, VA, mount a calibrated flow meter midway between the net
rims and mouth center (the meter is mounted off-center to avoid flow reduction associated with
the towing bridle).37 Equip meter with lock mechanisms to prevent it turning in reverse or while
in air. Record flow-meter readings before and after collecting sample. Calculate filtration
efficiency, E, from:
If E is less than about 0.8, substantial clogging has occurred. Take steps to increase efficiency.
Clogging not only decreases the volume filtered, but also leads to biased samples because
filtration efficiency is nonuniform during the tow.34
Various types of plankton nets are shown in Figure 10200:3. Simple conical nets have been
used for many years with little modification in design or improvement in accuracy. Their major
source of error is that the filtration characteristics of conical nets usually are unknown. Filtration
efficiency in No. 20 mesh cone nets ranges from 40 to 77%. To improve efficiency, place a
porous cylinder collar or nonporous truncated cone in front of the conical portion of the net. The
Juday net exemplifies a commonly used net with a truncated cone. For good filtration
characteristics the ratio of filtering area of net to orifice area should be at least 3:1. Bridles
attaching the net to the towing line also adversely influence filtration efficiency and increase
turbulence in front of the net, thereby increasing the potential for net avoidance by larger
zooplankters. The tandem, Bongo net design (Figure 10200:3) reduces these influences and
permits duplicate samples to be collected simultaneously.
Three types of tows are used: vertical, horizontal, and oblique. Vertical tows are preferred to
obtain an integrated water column sample. To make a vertical tow, lower the weighted net to a
given depth, then raise vertically at an even speed of 0.5 m/s.
In small water bodies haul the net hand over hand with a steady, unhurried motion
approximating the speed of 0.5 m/s. In large bodies where long net hauls and vessel drifting are
expected, use a davit, meter wheel, angle indicator, and winch. Attach a 3- to 5-kg weight to hold
the net down. Determine depth of the net by multiplying the length of the extended wire by the
cosine of the wire’s angle with the vertical direction. Maintain wire angle as close to the vertical
as possible by controlling the boat’s speed null against the wind drift, or wherever feasible, do
vertical hauls from an anchored boat.
Vertical and oblique tows collect a composite sample, whereas horizontal tows collect a
sample at a discrete depth. Oblique tows usually are preferred over vertical tows in shallow
water or wherever a longer net tow is required. For oblique tows, lower the net or sampler to
some predetermined depth and then raise at a constant rate as the boat moves forward. Oblique
tows do not necessarily sample a true angle from the bottom to the surface. Under best
conditions the pattern is somewhat sigmoid due to boat acceleration and slack in the tow line.
Horizontal tows usually are used to obtain depth distribution information on zooplankton.
Although a variety of horizontal samplers is available (see Figure 10200:4), use the
Clarke-Bumpus sampler38 for quantitative collection of zooplankton because of its built-in
flowmeter and opening-closing device. For horizontal tows use a boat equipped as above and
determine sampler depth as above. Lower sampler to preselected depth, open, tow at that depth
for 5 to 10 min, then close and raise it.
A variety of zooplankton sampling methods can be used in flowing water. The method of
choice depends largely on flow velocity. Properly weighted bottles, traps and pump hoses, and
nets can be used in medium- to slow-flowing waters. In turbulent, well-mixed waters, collect
surface water by bucket and filter it through the appropriate mesh size. Select sample size based
on concentration of zooplankters.
Give plankton nets proper care and maintenance. Do not let particulate matter dry on the net
because it can significantly reduce size of mesh apertures and increase frequency of clogging.
Wash net thoroughly with water after each use. Periodically clean with a warm soap solution.
Because nylon net material is susceptible to deterioration from abrasion and sunlight, guard
against unnecessary wear and store in the dark.
Traps and nets do not work well in shallow areas with growths of aquatic vegetation. To
obtain an integrated sample for the entire water column in such areas, use a length of
light-weight rubber or polyethylene tubing with netting attached over one end and a rope on the
other.39 Attach netting by tape or rubber bands that will stay in place in water, but can be
removed easily after sampling. Use tubing of 5- to 10-cm diam and long enough to reach from
the surface to the bottom. Lower the open end (the end with the rope attached) until it almost
touches the bottom. Then pull this end up using the rope and keep the covered end above the
water surface. When the open end is out of the water, let the end with the netting fall back into
the water, pull the tubing into the boat, open end first, and let the water in the tube drain out
through the netting. When the zooplankton has been concentrated in a small volume, just above
the netting, remove the netting over a container and catch the concentrated sample. Wash netting
and end of tubing into the container to assure that all the zooplankton is collected. This method is
not limited to areas with aquatic vegetation. It provides an excellent method of obtaining an
integrated sample from any shallow area. In standing waters, collect tow samples by filtering 1
to 5 m3 of water.
Preserve zooplankton samples with 70% ethanol or 5% buffered formalin. Ethanol
preservative is preferred for materials to be stained in permanent mounts or stored. Formalin may
be used for the first 48 h of preservation with subsequent transfer to 70% ethanol. Formalin
preservative may cause distortion of pleomorphic forms such as protozoans and rotifers. Make
formalin in sucrose-saturated water to minimize carapace distortion and loss of eggs in
crustaceans, especially cladocerans.40 Bouin’s fixative produces reasonable results for
soft-bodied microzooplankton.41 This fixative is picric acid saturated in calcium
carbonate-buffered formaldehyde containing 5% (v/v) acetic acid. Dilute Bouin’s fixative 1:19
with the sample. Because rapid fixation is necessary, pour the sample onto the fixative or inject
fixative rapidly into the sample.
Use a narcotizing agent such as carbonated water, menthol-saturated water, or neosynephrine
to prevent or reduce contraction or distortion of organisms, especially rotifers, cladocerans, and
many marine invertebrates.42,43 Adding a few drops of detergent prevents clumping of preserved
organisms. Preserve samples as soon as most animal movement has ceased, usually within a half
hour of narcotization. To prevent evaporation, add 5% glycerin to the concentrated sample. In
turbid samples, differentiate animal and detrital material by adding 0.04% rose bengal stain,
which intensely stains the carapace (shell) of zooplankters and is a good general cytoplasmic
stain.
3. References
1. U.S. ENVIRONMENTAL PROTECTION AGENCY. 1982. Handbook for Sampling and
Sample Preservation of Water and Wastewater. EPA-600/4-82-029.
2. PARKER B.C. & R.F. HATCHER. 1974. Enrichment of surface freshwater microlayers with
algae. J. Phycol. 10:185.
3. TAGUCHI, S. & K. NAKAJIMA. 1971. Plankton and seston in the sea surface of three
inlets of Japan. Bull. Plankton Soc. Japan 18:20.
4. UNITED NATIONS EDUCATIONAL, SCIENTIFIC AND CULTURAL ORGANIZATION. 1966.
Determination of Photosynthetic Pigments in Sea-water. Monogr. Oceanogr. Methodol.
No. 1. United Nations Educational, Scientific & Cultural Org., Paris.
5. UNITED NATIONS EDUCATIONAL, SCIENTIFIC AND CULTURAL ORGANIZATION. 1968.
Zooplankton Sampling. Monogr. Oceanogr. Methodol. No. 2. United Nations
Educational, Scientific & Cultural Org., Paris.
6. UNITED NATIONS EDUCATIONAL, SCIENTIFIC AND CULTURAL ORGANIZATION. 1973. A
Guide to the Measurement of Marine Primary Production under Some Special
Conditions. Monogr. Oceanogr. Methodol. No. 3. United Nations Educational,
Scientific & Cultural Org., Paris.
7. SOURNIA, A., ed. 1978. Phytoplankton Manual. Monogr. Oceanogr. Methodol. No. 6.
United Nations Educational, Scientific & Cultural Org., Paris.
8. CUPP, E.E. 1943. Marine plankton diatoms of the west coast of North America. Bull.
Scripps Inst. Oceanogr. 5:1.
9. HUSTEDT, F. 1927–66. Die Kieselalgen Deutschlands, Osterreichs und der Schweiz mit
Berucksichtigung der Ubrigen Lander Europas Sowie der Angrenzenden
Meeresgebiete. In L. Rabenhorst, Kryptogamen-Flora. Vol. 7: Teil 1 (1927–30); Teil 2
(1931–59); Teil 3 (1961–66). Akademie Verlag, Leipzig, Germany.
10. LEBOUR, M.V. 1930. The Planktonic Diatoms of Northern Seas. Ray Soc., London.
11. HENDEY, N.I. 1964. An introductory account of the smaller algae of British coastal
waters, V. Bacillariophyceae (Diatoms). Fish. Invest. Min. Agr. Fish. Food (G.B.), Ser.
IV:1.
12. DODGE, J.D. 1975. The prorocentrales (Dinophyceae), II. Revision of the taxonomy
within the genus Prorocentrum. Bot. Limnol. Soc. 71: 103.
13. LEBOUR, M.V. 1925. The Dinoflagellates of Northern Seas. Marine Biological Assoc.
United Kingdom, Plymouth.
14. SCHILLER, J. 1931–37. Dinoflagellatae (Peridineae) in monographischer Behandlung.
In L. Rabenhorst, Kryptogamen-Flora. Vol. 10; Teil 1 (1931–33); Teil 2 (1935–37).
Akademie Verlag, Leipzig, Germany.
15. SCHILLER, J. 1930. Coccolithineae. In L. Rabenhorst, Kryptogamen-Flora. Vol. 10, p.
89. Akademie Verlag, Leipzig, Germany.
16. GEITLER, L. 1932. Cyanophyceae von Europa unter Berucksichtigung der anderen
Kontinente. In L. Rabenhorst, Kryptogamen-Flora. Vol. 14, p. 1. Akademie Verlag,
Leipzig, Germany.
17. WELCH, P.S. 1948. Limnological Methods. Blakiston Co., Philadelphia, Pa.
18. STRICKLAND, J.D.H. & T.R. PARSONS. 1968. A Practical Manual of Sea Water Analysis.
Fish. Res. Board Can. Bull. No. 167. Queen’s Printer, Ottawa, Ont.
19. DUSSART, B.M. 1965. Les differentes categories de plancton. Hydrobiologia 26:72.
20. SIEBURTH, J.MCN., V. SMETACEK & J. LENZ. 1978. Pelagic ecosystem structure:
Heterotrophic compartments of plankton and their relationship to plankton size
fractions. Limnol. Oceanogr. 23: 1256.
21. MORTIMER, C.H. 1942. The exchange of dissolved substances between mud and water
in lakes. J. Ecol. 30:147.
22. VOLLENWEIDER, R.A. 1969. A Manual on Methods for Measuring Primary Production
in Aquatic Environments. IBP Handbook No. 12. Blackwell Scientific Publ., Oxford,
England.
23. GELDREICH, E.E., H.D. NASH, D.F. SPINO & D.J. REASONER. 1980. Bacterial dynamics in a
water supply reservoir: a case study. J. Amer. Water Works Assoc. 72:31.
24. BEERS, J.R. 1978. Pump sampling. In A. Sournia, ed. Phytoplankton Manual. United
Nations Educational, Scientific and Cultural Org., Paris.
25. EXTON, R.J., W.M. HOUGHTON, W. ESAIAS, L.W. HAAS & D. HAYWARD. 1983. Spectral
differences and temporal stability of phycoerythrin fluorescence in estuaries and
coastal waters due to the domination of labile cryptophytes and stable cyanobacteria.
Limnol. Oceanogr. 28:1225.
26. EDMONDSON, W.T., ed. 1959. Freshwater Biology, 2nd ed. John Wiley & Sons, New
York, N.Y.
27. UTERMOHL, H. 1958. Zur Vervollkommung der quantitativen Phytoplankton-Methodik.
Int. Ver. Theoret. Angewand. Limnol., Commun. No. 9.
28. WEBER, C.I. 1968. The preservation of phytoplankton grab samples. Trans. Amer.
Microsc. Soc. 87:70.
29. PAERL, H.W. 1984. An evaluation of freeze fixation as a phytoplankton preservation
method for microautoradiography. Limnol. Oceanogr. 29:417.
30. SILVER, M.W. & P.J. DAVOLL. 1978. Loss of 14C activity after chemical fixation of
phytoplankton: Error source for autoradiography and other productivity measurements.
31. JUDAY, C. 1916. Limnological apparatus. Trans. Wis. Acad. Sci. 18: 566.
32. SCHINDLER, D.W. 1969. Two useful devices for vertical plankton and water sampling.
J. Fish. Res. Board Can. 26: 1948.
33. SCHWOERBEL, J. 1970. Methods of Hydrobiology. Pergamon Press, Toronto, Ont.
34. TRANTER, D.J., ed. 1980. Reviews on Zooplankton Sampling Methods. United Nations
Educational, Scientific & Cultural Org., Switzerland.
35. GANNON, J.E. 1980. Towards improving the use of zooplankton in water quality
surveillance of the St. Lawrence Great Lakes. Proc. 1st Biol. Surveillance Symp., 22nd
Conf. Great Lakes Research Can. Tech. Rep. Fish. Aquat. Sci. 976, p. 87.
36. ROBERTSON, A. 1968. Abundance, distribution, and biology of plankton in Lake
Michigan with the addition of a Research Ships of Opportunity project. Spec. Rep. No.
35, Great Lakes Research Div., Univ. Michigan, Ann Arbor.
37. EVANS, M.S. & D.W. SELL. 1985. Mesh size and collection characteristics of 50-cm
diameter conical plankton nets. Hydrobiologia 122: 97.
38. CLARKE, G.L. & D.F. BUMPUS. 1940. The Plankton Sampler: An Instrument for
Quantitative Plankton Investigations. Spec. Publ. No. 5, Limnological Soc. America.
39. PENNAK, R.W. 1962. Quantitative zooplankton sampling in littoral vegetation areas.
Limnol. Oceanog. 7:487.
40. HANEY, J.F. & D.J. HALL. 1973. Sugar-coated Daphnia; A preservation technique for
Cladocera. Limnol. Oceanogr. 18:331.
41. COATS, D.W. & J.F. HEINBOKEL. 1982. A study of reproduction and other life cycle
phenomena in plankton protists using an acridine orange fluorescence technique. Mar.
Biol. 67:71.
42. GANNON, J.E. & S.A. GANNON. 1975. Observations on the narcotization of crustacean
zooplankton. Crustaceana 28(2):220.
43. STEEDMAN, H.F. 1976. Narcotizing agents and methods. In H.F. Steedman, ed.
Zooplankton Fixation and Preservation. Monogr. Oceanogr. Methodol. No. 4. United
Nations Educational, Scientific & Cultural Org., Paris.
10200 C.
Concentration Techniques
The organisms contained in water samples sometimes must be concentrated in the laboratory
before analysis. Three techniques for concentrating phytoplankton, namely, sedimentation,
membrane filtration, and centrifugation, are described below. A special technique for
zooplankton also is given.
1. Sedimentation
Sedimentation is the preferred method of concentration because it is nonselective (unlike
filtration) and nondestructive (unlike filtration or centrifugation), although many of the
picoplankton, the smaller nannoplankton, and actively swimming flagellates (in unpreserved
samples) may not settle completely. The volume concentrated varies inversely with the
abundance of organisms and is related to sample turbidity. It may be as small as 1 mL for use
with an inverted microscope or as large as 1 L for general phytoplankton and zooplankton
enumeration.
Allow 1 h settling/mm of column depth. For a treated sample (10 mL liquid detergent/L)
allow about 0.5 h settling/mm depth.1 The sample may be concentrated in a series of steps by
quantitatively transferring the sediment from the initial container to sequentially smaller ones.
Use cylindrical settling chambers with thin, clear glass bottoms. Fill settling chambers without
forming a vortex, keep them vibration-free, and move them carefully to avoid nonrandom
distribution of settled matter. Carefully siphon or decant the supernatants to obtain the desired
final volume (5 mL for diatom mounts). Store the concentrated sample in a closed, labeled glass
vial.
2. Membrane Filtration
The filtration method permits use of high magnification for enumerating small plankters
including flagellates and cyanobacteria. However, delicate forms such as ‘‘naked’’ flagellates
are distorted by even gentle filtration. When populations are dense and the content of detritus is
high, the filter clogs quickly and silt may crush the organisms or obscure them from view.
Pour a measured volume of well-mixed sample into a funnel equipped with a membrane filter
having a pore diameter of 0.45 µm. Apply a vacuum of less than 50 kPa to the filter until about
0.5 cm of sample remains on filter. Break vacuum, then apply low vacuum (about 12 kPa) to
remove remaining water but not to dry the filter.
For samples with a low phytoplankton and silt content the method does not require counting
of individual plankters to assemble enumeration data and it increases the probability of
observing less abundant forms.2 Samples also may be concentrated on a filter, inverted onto a
microscope slide, and quick-frozen, permitting the removal of the filter and transfer of plankton
to the slide.3,4
3. Centrifugation
Plankton can be concentrated by batch or continuous centrifugation. Centrifuge batch
samples at 1000 g for 20 min. The Foerst continuous centrifuge is no longer recommended as a
quantitative device but it may be desirable to continue its use in existing programs to assure
continuity with previously collected data. Although centrifugation accelerates sedimentation, it
may damage fragile organisms.
4. Zooplankton Concentration
Zooplankton samples often need to be concentrated in the field, especially when large water
bottles or pump methods of sampling are used. Moreover, samples obtained by nets or other
methods sometimes need to be concentrated further for storage or preparation for examination.
When only small volume reductions are needed, pour sample back into the bucket of traps or
nets. In processing large volumes of water as with pump sampling, use larger plankton buckets
or funnels with greater water volume retention and filtration surface area. Construct a filter
funnel similar to that shown in Figure 10200:5 of clear acrylic plastic or other suitable material.5
The volume of the apparatus and the mesh size depend on volume of water to be filtered and size
of organisms to be retained. The mesh size of the filter funnel normally is the same as that of the
net or other field sampling device.
5. References
1. FURET, J.E. & K. BENSON-EVANS. 1982. An evaluation of the time required to obtain
sedimentation of fixed algal particles prior to enumeration. Brit. Phycol. J. 17:253.
2. MCNABB, C.D. 1960. Enumeration of freshwater phytoplankton concentrated on the
membrane filter. Limnol. Oceanogr. 5:57.
3. HEWES, C.D. & O. HOLM-HANSEN. 1983. A method for recovering nanoplankton from
filters for identification with the microscope: The filter-transfer-freeze (FTF)
technique. Limnol. Oceanogr. 28:389.
4. HEWES, C.D., F.M.H. REID & O. HOLM-HANSEN. 1984. The quantitative analysis of
nanoplankton: A study of methods. J. Plankton Res. 6:601.
5. LIKENS, G.E. & J.J. GILBERT. 1970. Notes on quantitative sampling of natural
populations of planktonic rotifers. Limnol. Oceanogr. 15:816.
10200 D.
Preparing Slide Mounts
1. Phytoplankton Semi-Permanent Wet Mounts
Agitate the settled sample concentrate and withdraw a subsample with an accurately
calibrated pipet. Clean pipet regularly. To prepare wet mounts transfer 0.1 mL to a glass slide,
place a cover slip over the sample, and ring the cover slip with an adhesive such as clear nail
polish to prevent evaporation. For semipermanent mounts, add a few drops of glycerin to the
slide. As the sample ages the water evaporates, leaving the organisms imbedded in the glycerin.
If the cover slip is ringed with adhesive, the slide can be retained for a few years if stored in the
dark.
2. Phytoplankton Permanent Mounts
a. Membrane filter mounts: Place two drops of immersion oil on a labeled slide. Immediately
after filtering place the filter on top of the oil with a pair of forceps and add two drops of oil on
top of the filter. The oil impregnates the filter and makes it transparent. Impregnation time is 24
to 48 h. This procedure can be completed in 1 to 2 h by applying heat (70°C). Once the filter has
cleared, place a few additional drops of oil on it and cover with a cover slip. The mounted filter
is now ready for microscopic examination. Alternatively, mount membrane filters in mounting
medium.*#(113) Immerse filters in 1-propanol to displace residual water and transfer to xylol for
several minutes to clear filters. Place a section of filter or entire filter on a microscope slide with
the mounting medium, cover with a cover glass, and dry at low temperature.1
b. Sedimented slide mounts: Two techniques are available for making permanent, resin
mounts of natural phytoplankton that has been deposited by sedimentation on a microscope slide
or cover glass and dehydrated by ethanol vapor substitution.2,3
3. Diatom Mounts
Samples concentrated for diatom analysis by settling or centrifugation may contain
dissolved materials, such as marine salts, formalin, and detergents, that will leave interfering
residues. Wash well with distilled water before slide preparation. Transfer several drops of
washed concentrate by means of a large-bore disposable pipet or large-bore dropper to a cover
glass on a hot plate warmed enough to increase the evaporation rate but not enough to cause
boiling (use a large-bore pipet or dropper to prevent possible selective filtration, thus exclusion,
of larger forms or those forming colonies or chains). If the cleaned material is very concentrated,
improve distribution of diatoms by adding the drops to a cover glass already flooded with
distilled water. Evaporate to dryness. Repeat addition and evaporation until a sufficient quantity
of sample has been transferred to the cover glass, but avoid producing a residue so dense that
organisms cannot be recognized. If in doubt about the density, examine under a compound
microscope. After evaporation, incinerate the residue on the cover glass on a hot plate at 300 to
500°C; alternatively, use a muffle furnace. This usually requires 20 to 45 min. Mount as
described below.
Treat samples concentrated for diatom analysis by membrane filtration as described by
Patrick and Reimer.4 Mix equal volumes of conc nitric acid (HNO3) and sample. CAUTION:
When working with conc HNO3 wear safety goggles and an acid-resistant apron and gloves, and
work under a hood. Add a few grains of potassium dichromate (K2Cr2O7)5 to facilitate digestion
of the filter and cellular organic matter. Add more dichromate if solution color changes from
yellow to green. Place sample on a hot plate and boil down to approximately one-third the
original volume. Alternatively, let treated sample stand overnight. This cleaning process destroys
organic matter and leaves only diatom shells (frustules). Cool, wash with distilled water, and
mount as described above. Transfer cleaned frustules to a cover glass and dry as described
above.
Place a drop of mounting medium in the center of a labeled slide. Use 25- by 75-mm slides
with frosted ends. Using a suitable high-refractive-index microscopic mounting medium assures
permanent, easily handled mounts for examination under oil immersion. Heat the slide to near
90°C for 1 to 2 min before applying the heated cover slip with its sample residue to hasten
evaporation of solvent in the mounting medium. Remove the slide to a cool surface and, during
cooling (5 to 10 s), apply firm but gentle pressure to the cover glass with a broad, flat instrument.
4. Zooplankton Mounts
For zooplankton analyses, withdraw a 5-mL subsample from the concentrate and dilute or
concentrate further as necessary. Transfer sample to a counting cell or chamber (see below) for
analysis as a wet mount. Use polyvinyl lactyl phenol†#(114) for preparing semipermanent
zooplankton mounts. The mounts are good for about a year, after which time the clearing agent
causes deterioration of organisms. For long-term storage ring cover slip with clear lacquer
(fingernail polish) to retard mountant crystallization. For permanent mounting, other mountants
are available.‡#(115)
For the protozoan portion of the microzooplankton, a protargol staining procedure6 not only
provides a permanent mount but also reveals the cytological details often necessary for
identification. This procedure is qualitative and is especially important in taxonomic studies of
the ciliated protozoa.
5. References
1. MILLIPORE FILTER CORPORATION. 1966. Biological examination of water, sludge and
bottom materials. Millipore Techniques, Water Microbiology, p. 25.
2. SANFORD, G.R., A. SANDS & C.R. GOLDMAN. 1962. A settle-freeze method for
concentrating phytoplankton in quantitative studies. Limnol. Oceanogr. 14:790.
3. CRUMPTON, W.G. & R.G. WETZEL. 1981. A method for preparing permanent mounts of
phytoplankton for critical microscopy and cell counting. Limnol. Oceanogr. 26:976.
4. PATRICK, R. & C.W. REIMER. 1967. The Diatoms of the United States. Vol. 1. Monogr.
13, Philadelphia Acad. Natur. Sci.
5. HOHN, M.H. & J. HELLERMAN. 1963. The taxonomy and structure of diatom populations
for three eastern North American rivers using three sampling methods. Trans. Amer.
Microsc. Soc. 62:250.
6. SMALL, E.B. & D.H. LYNN. 1985. Phylum Ciliophora Doflein, 1901. In J.J. Lee, S.H.
Hunter & E.C. Bovee, eds. An Illustrated Guide to the Protozoa. Soc. Protozoology,
Lawrence, Kansas.
10200 E.
Microscopes and Calibrations
1. Compound Microscope
Use either a standard or an inverted compound microscope for algal identification and
enumeration. Equip either type with a mechanical stage capable of moving all parts of a counting
cell past the objective lens. Standard equipment is a set of 10× or 12.5× oculars and 10×, 20×,
40×, and 100× objectives. Use objectives to provide adequate working distance for the counting
chamber. Magnification requirements vary with the plankton fraction being investigated, the
type of microscope, counting chamber used, and optics. With standard objectives, the
Sedgwick-Rafter chamber limits magnification to approximately 200× and the Palmer-Maloney
cell limits magnification to approximately 500×. Inverted microscopes are limited in resolution
by their optics. The useful upper limit of magnification for any objective is 1000 times the
numerical aperture (NA). Above this magnification, no greater detail can be resolved. Use
combinations of oculars, intermediate magnifiers, and objectives to obtain the greatest
magnification without exceeding the useful limit of magnification. When the limit is exceeded,
empty magnification results. Empty magnification occurs where the image is larger but no
greater resolution is achieved. Optics providing contrast enhancement such as phase contrast or
differential interference contrast are useful.
2. Stereoscopic Microscope
The stereoscopic microscope is essentially two complete microscopes assembled into a
binocular instrument to give a stereoscopic view and an erect rather than an inverted image. Use
this microscope for the study and counting of large plankters such as mature microcrustacea.
Include 10× to 15× paired oculars in combination with 1× to 8× objectives. This combination of
optics bridges the gap between the hand lens and the compound microscope and provides
magnification ranging from 10× to 120×. Alternatively, use a good-quality zoom-type instrument
with comparable magnification.
3. Inverted Compound Microscope
The inverted compound microscope often is used routinely for plankton counting in many
laboratories.1-3 This instrument is unique in that the objectives are below a movable stage and
the illumination comes from above, thus permitting viewing of organisms that have settled to the
bottom of a chamber. Place samples in a cylindrical settling chamber having a thin, clear glass
bottom. Chambers of various capacities are available; the appropriate size depends on the density
of organisms. After a suitable period of settling (see Section 10200C.1), count organisms in the
settling chamber.
The major advantage of the inverted microscope is that by a simple rotation of the nosepiece
a specimen can be examined (or counted) directly in the settling chamber at any desired
magnification. Although not recommended, oil immersion objectives have some useful
applications. No preparation or manipulation other than settling is required. Generally, examine
a preserved sample. Techniques are available for samples with an abundance of organisms that
tend to float.4
4. Epifluorescence Microscope
An epifluorescence microscope may be either standard or inverted. It uses incident light to
excite electrons in intracellular compounds, such as pigments or absorbed stains, with the energy
emitted during electron return to the ground state being measured as fluorescent light. The
technique has been applied to the microscopic identification of chlorophyll-containing cells
(autotrophs) and nonpigmented heterotrophic plankton; fluorescent stains such as primulin or
proflavin also have been used to differentiate nannoplanktonic primary and secondary
producers.5-7 Excitation and emission wavelengths are unique for each pigment and stain and
require distinct light filter combinations and light sources. Select the filter combinations for the
particular application. Epifluorescence microscopy is particularly useful for the enumeration of
picoplankton and heterotrophic flagellate populations common to most aquatic systems.
Concentrate samples by membrane filtration. Use epifluorescence microscopy as a
complementary procedure to standard light microscope counting techniques.
5. Microscope Calibration
Microscope calibration is essential. The usual equipment for calibration is a Whipple grid
(ocular micrometer, reticle, or reticule) placed in an eyepiece of the microscope and a stage
micrometer that has a standardized, accurately ruled scale on a glass slide. The Whipple disk
(Figure 10200:6) has an accurately ruled grid subdivided into 100 squares. One square near the
center is subdivided further into 25 smaller squares. The outer dimensions of the grid are such
that with a 10× objective and a 10× ocular, it delimits an area of approximately 1 mm2 on the
microscope stage. Because this area may differ from one microscope to another, carefully
calibrate the Whipple grid for each microscope.
With the ocular and stage micrometers parallel and in part superimposed, match the line at
the left edge of the Whipple grid with the zero mark on the stage micrometer scale (Figure
10200:7). Determine the width of the Whipple grid image to the nearest 0.01 mm from the stage
micrometer scale. Should the width of the image of the Whipple grid be exactly 1 mm (1000
µm), the larger squares will be 1/10 mm (100 µm) on a side and each of the smaller squares 1/50
mm (20 µm).
When the microscope is calibrated at higher magnifications, the entire scale on the stage
micrometer will not be seen; make measurements to the nearest 0.001 mm. Additional details for
calibration are available.8
6. References
1. WETZEL, R.G. & G.E. LIKENS. 1991. Limnological Analyses, 2nd ed. Springer-Verlag,
New York, N.Y.
2. LUND, J.W.G., C. KIPLING & E.D. LECREN. 1958. The inverted microscope method of
estimating algal numbers and the statistical basis of estimations by counting.
Hydrobiologia 11:143.
3. SICKO-GOAD, L. & E.F. STOERMER. 1984. The need for uniform terminology concerning
phytoplankton cell size fractions and examples of picoplankton from the Laurentian
Great Lakes. J. Great Lakes Res. 10:90.
4. REYNOLDS, C.S. & G.H.M. JAWORSKI. 1978. Enumeration of natural Microcystis
populations. Brit. Phycol. J. 13:269.
5. DAVIS, P.G. & J. MCN. SIEBURTH. 1982. Differentiation of phototrophic and
heterotrophic nanoplankton populations in marine waters by epifluorescence
microscopy. Ann. Inst. Oceanogr. 58:249.
6. CARON, D.A. 1983. Techniques for enumeration of heterotrophic and phototrophic
nanoplankton, using epifluorescence microscopy, and comparison with other
procedures. Appl. Environ. Microbiol. 46:491.
7. SHERR, E.B. & B.F. SHERR. 1983. Double-staining epifluorescence techniques to assess
frequency of dividing cells and bacteriovory in natural populations of heterotrophic
microprotozoa. Appl. Environ. Microbiol. 46:1388.
8. JACKSON, H.W. & L.G. WILLIAMS. 1962. Calibration and use of certain plankton
counting equipment. Trans. Amer. Microsc. Soc. 81:96.
10200 F.
Phytoplankton Counting Techniques
1. Counting Units
Some phytoplankton are unicellular while others are multicellular (colonial). The variety of
configurations poses a problem in enumeration. For example, should a four-celled colony of
Scenedesmus (Plate 32) be reported as one colony or four individual cells? Listed below are
suggestions for reporting:
suggestions for reporting:
Enumeration
Method
Total cell count
Natural unit count1
(clump count)
Areal standard
unit count*#(116)
Counting
Unit
Reporting
Unit
One cell
One organism
(any unicellular
organism or
natural colony)
Cells/mL
Units/mL
400 µm2
Units/mL
Making a total cell count is time-consuming and tedious, especially when colonies consist of
thousands of individual cells. The natural unit or clump is the most easily used system; however,
it is not necessarily accurate because sample handling and preserving may dislodge cells from
the colony. The unit method also may not be quantitatively accurate nor reflect abundance of
biomass or biovolume. Whatever method is chosen, identify it in reporting results.
If the distribution of organisms is random and the population fits a Poisson distribution, the
counting error may be estimated.2 For example, the approximate 95% confidence limits, as a
percentage of the number of units counted (N), equals:
Thus, if 100 units are counted, the 95% confidence limits approximate ± 20%. For a count of 400
units, the limits are about 10%.
2. Counting Procedures
To enumerate plankton use a counting cell or chamber that limits the volume and area for
ready calculation of population densities.
When counting with a Whipple grid, establish a convention for tallying organisms lying on
an outer boundary line. For example, in counting a ‘‘field’’ (entire Whipple square), designate
the top and left boundaries as ‘‘no-count’’ sides, and the bottom and right boundaries as
‘‘count’’ sides. Thus, tally every plankter touching a ‘‘count’’ side from the inside or outside but
ignore any touching a ‘‘no-count’’ side. If significant numbers of filamentous or other large
forms cross two or more boundaries of the grid, count them separately at a lower magnification
and include their number in the total count.
To identify organisms use standard bench references (see Section 10900).
Do not count dead cells or broken diatom frustules. Tally empty centric and pennate diatoms
separately as ‘‘dead centric diatoms’’ or ‘‘dead pennate diatoms’’ for use in converting the
diatom species proportional count to a count per milliliter.
Magnification is important in phytoplankton identification and enumeration. Although
magnifications of 100× to 200× are useful for counting large organisms or colonies, much higher
magnifications often are required. It is useful to categorize techniques for phytoplankton
counting according to the magnifications provided.
a. Low-magnification (up to 200×) methods: The Sedgwick-Rafter (S-R) cell is a device
commonly used for plankton counting because it is easily manipulated and provides reasonably
reproducible data when used with a calibrated microscope equipped with an eyepiece measuring
device such as the Whipple grid.
The greatest disadvantage associated with the cell is that objectives providing high
magnification cannot be used. As a result, the S-R cell is not appropriate for examining
nannoplankton. The S-R cell is approximately 50 mm long by 20 mm wide by 1 mm deep. The
total area of the bottom is approximately 1000 mm2 and the total volume is approximately 1000
mm3 or 1 mL. Carefully check the exact length and depth of the cell with a micrometer and
calipers before use.
1) Filling the cell—Before filling the S-R cell with sample, place the cover glass diagonally
across the cell and transfer sample with a large-bore pipet (Figure 10200:8). Placing cover slip in
this manner will help prevent formation of air bubbles in cell corners. The cover slip often will
rotate slowly and cover the inner portion of the S-R cell during filling. Do not overfill because
this would yield a depth greater than 1 mm and produce an invalid count. Do not permit large air
spaces caused by evaporation to develop in the chamber during a lengthy examination. To
prevent formation of air spaces, occasionally place a small drop of distilled water on edge of
cover glass.
Before counting let the S-R cell stand for at least 15 min to settle plankton. Count plankton
on the bottom of the S-R cell. Some phytoplankton, notably some blue-green algae or motile
flagellates in unpreserved samples, may not settle but rise to the underside of the cover slip.
When this occurs, count these organisms and add to total of those counted on the cell bottom to
derive total number of organisms. Count algae in strips or fields.
2) Strip counting—A ‘‘strip’’ the length of the cell constitutes a volume approximately 50
mm long, 1 mm deep, and the width of the total Whipple grid.
The number of strips to be counted is a function of the precision desired and the number of
units (cells, colonies, or filaments) per strip. Derive number of plankton in the S-R cell from the
following:
where:
C=
L=
D=
W=
S=
number of organisms counted,
length of each strip (S-R cell length), mm,
depth of a strip (S-R cell depth), mm,
width of a strip (Whipple grid image width), mm, and
number of strips counted.
Multiply or divide number of cells per milliliter by a correction factor to adjust for sample
dilution or concentration.
3) Field counting—On samples containing many plankton (10 or more plankters per field),
make field counts rather than strip counts. Count plankters in random fields each consisting of
one Whipple grid. The number of fields counted will depend on plankton density and statistical
accuracy desired (see Section 10200F.1). Calculate the number of plankton per milliliter as
follows:
where:
C=
A=
D=
F=
area of a field (Whipple grid image area), mm2,
depth of a field (S-R cell depth), mm, and
number of fields counted.
Multiply or divide the number of cells per milliliter by a correction factor to adjust for
sample dilution or concentration.
b. Intermediate magnification (low to 500×) methods: The Palmer-Maloney (P-M)
nannoplankton cell3 is designed specifically for nannoplankton enumeration. It has a circular
chamber with a 17.9-mm diam, 0.4-mm depth, and 0.1-mL volume. The shallow depth permits
use of 40 to 45× objectives with sufficient working distance. The principal disadvantage of the
P-M cell is that these magnifications (400 to 450×) often are insufficient for nannoplankton
identification and enumeration.
Because a relatively small sample portion is examined in the P-M cell do not use it unless
the sample contains a dense population (10 or more plankters per field). Such a small sample
portion from a less dense population causes serious underestimation of density.
Introduce sample with a pipet into one of the 2- by 5-mm channels on the side of the
chamber with the cover slip in place. After a 10-min settling period count the plankters in
random fields, with the number of fields depending on density and variety of plankton and the
statistical accuracy desired. Strips may be counted in this or any other circular cell by measuring
the effective diameter and counting two perpendicular strips that cross at the center. Calculate
the number per milliliter as follows:
where:
C=
A=
D=
F=
area of a field (Whipple grid image), mm2,
depth of a field (P-M cell depth), mm, and
number of fields counted.
Multiply or divide the number of cells per milliliter by a correction factor to adjust for
sample dilution or concentration.
Another readily available chamber is the standard medical hemacytometer used for
enumerating blood cells. It has a ruled grid machined into a counting plate and is fitted with a
ground-glass cover slip. The grid is divided into 1-mm2 divisions; the chamber is 0.1 mm deep.
Introduce sample by pipet and view under 450× magnification. Count all cells within the grid.
The chamber comes from the manufacturer with a detailed instruction sheet containing directions
on calculations and proper usage. A disadvantage to these counting cells is that the sample must
have a very high plankton density to yield statistically reliable data.
c. High-magnification methods: Examination of phytoplankton at high magnification
requires the use of oil immersion objectives. Suitable procedures include using inverted
microscope chambers, membrane filter mounts, sedimented slide mounts, the Lackey drop
method, and diatom mounts.
1) Inverted microscope counts—Prepare a sample for examination by filling the settling
chamber. After the desired settling time (see Section 10200C.1), transfer the chamber to the
microscope stage. Count perpendicular strips across the center of the bottom cover glass. Strip
counts may be made by using a Whipple grid or special counting oculars that have a pair of
adjustable parallel hairs and a single cross hair. Determine the width of the strip with a stage
micrometer and tally organisms as they pass the single cross hair that functions as a reference
point. Hold strip width constant for any series of samples. Alternatively examine random
nonoverlapping fields until at least 100 units of the dominant species are counted. For highest
accuracy, particularly because algae distribution may be nonuniform, count the entire chamber
floor. Alternatively, make a random field-minimum count to attain a precision level of at least
85%.4
where:
C=
At =
L=
W=
S=
V=
total area of bottom of settling chamber, mm2,
length of a strip, mm,
width of a strip (Whipple grid image width), mm,
number of strips counted, and
volume of sample settled, mL.
where:
A f = area of a field (Whipple grid image area), mm2,
F = number of fields counted,
and other terms are as defined above.
2) Membrane filter mounts—Concentrate sample as directed in Section 10200C.2 and
prepare membrane filter as directed in Section 10200D.2a.
Examine samples, concentrated on unlined membrane filters and mounted in oil as described
above. Count enough random fields to ensure desired level of statistical accuracy (see Section
10200F.1). Select magnification level and size of microscope field (quadrat) such that the most
abundant species appear in at least 70% but not more than 90% of microscopic fields examined
(80% is optimum). Adjust microscope field size by using part or all of the Whipple grid.
Examine 30 random microscope fields and record number of fields in which each species
occurred. Report results as organisms per milliliter, calculated as follows:
where:
N=
Q=
V=
D=
density (organisms/field) from Table 10200:II,
number of fields per filter,
milliliters filtered, and
dilution factor (0.96 for 4% formalin preservative).
3) Sedimented slide mounts—Examine mounts prepared as directed in Section 10200D.2b.
4) Lackey drop method—The Lackey drop (microtransect) method5 is a simple method of
obtaining counts of considerable accuracy with samples containing a dense plankton population.
It is similar to the S-R strip count.
Prepare slides as directed in Section 10200D.1. Oil immersion objectives can be used with
the semipermanent slides. Count organisms in enough strips to ensure desired level of statistical
accuracy (see Section 10200F.1). Calculate number of organisms per milliliter as follows:
where:
C = number of organisms counted,
At = area of cover slip, mm2,
As = area of one strip, mm2,
S = number of strips counted, and
V = volume of sample under the cover slip, mL.
5) Diatom mounts—Prepare samples as directed in Section 10200D.3.
For diatom species proportional count, examine diatom samples under oil immersion at a
magnification of at least 900×. Scan lateral strips the width of the Whipple grid until at least 250
cells are counted. Available time and accuracy required dictate the number of cells to be
counted. Determine percentage abundance of each species from tallied counts and calculate
counts per milliliter of each species by multiplying percent abundance by total live and dead
diatom count obtained from the plankton counting chamber. For greater accuracy distinguish
between living and dead diatoms at the species level.
6) Phytoplankton staining technique—Staining algae permits differentiation between ‘‘live’’
and ‘‘dead’’ diatoms.6 This permits enumerating total phytoplankton in a single sample without
sacrificing detailed diatom taxonomy. It also results in permanent reference slides. The
procedure is most useful when diatoms are major components of phytoplankton and it is
important to distinguish between living and dead diatoms.
Preferably preserve samples in Lugol’s solution or alternatively in formalin (see Section
10200B.3). For analysis thoroughly mix the sample and filter a portion through a 47-mm-diam
membrane filter (pore diam 0.45 or 0.65 µm). Use a vacuum of 16 to 20 kPa and never let
sample dry. Add 2 to 5 mL aqueous acid fuchsin solution (dissolve 1 g acid fuchsin in 100 mL
distilled water to which 2 mL glacial acetic acid has been added; filter) to the filter and let stand
for 20 min. After staining, filter sample, wash briefly with distilled water, and filter again.
Administer successive rinses of 50%, 90%, and 100% propanol to the sample while filtering.
Soak for 2 min in a second 100% propanol wash, filter, and add xylene. At least two washes are
required; let the final one soak 10 min before filtering. Trim the xylene-soaked filter and place
on a microscope slide on which there are several drops of mounting medium.†#(117) Apply
several more drops of medium to top of filter and install a cover glass. Carefully squeeze out
excess mounting medium. Make the final mount permanent by lacquering the edges of the cover
glass.
Count organisms using the most appropriate magnification. ‘‘Live’’ diatoms typically are red
while ‘‘dead’’ ones are unstained. Oil immersion is necessary for species identifications of
diatoms and many other algae. Count either strips or random fields and calculate plankton
densities per milliliter:
where:
C=
At =
Ac =
V=
total area of effective filter before trimming and mounting,
area counted (strips or fields), and
volume of sample filtered, mL.
3. References
1. INGRAM, W.M. & C.M. PALMER. 1952. Simplified procedures for collecting, examining,
and recording plankton in water. J. Amer. Water Works Assoc. 44:617.
3. PALMER, C.M. & T.E. MALONEY. 1954. A New Counting Slide for Nannoplankton.
Spec. Publ. No. 21, American Soc. Limnology & Oceanography.
4. SOURNIA, A., ed. 1978. Phytoplankton Manual. Monogr. Oceanogr. Methodol. No. 6.
United Nations Educational, Scientific & Cultural Org., Paris.
5. LACKEY, J.B. 1938. The manipulation and counting of river plankton and changes in
some organisms due to formalin preservation. Pub. Health Rep. 53:2080.
6. OWEN, B.B., JR., M. AFZAL & W.R. CODY. 1978. Staining preparations for phytoplankton
and periphyton. Brit. Phycol. J. 13:155.
10200 G.
Zooplankton Counting Techniques
1. Subsampling
Count entire samples having low zooplankton numbers (<200 zooplankters) without
subsampling. However, most zooplankton samples will contain more organisms than can be
enumerated practically; therefore, use a subsampling procedure. Before subsampling, remove
and enumerate all large uncommon organisms such as fish larvae in fresh water or coelenterates,
decapods, fish larvae, etc., in salt water. Subsample by the pipet or splitting method.
In the pipet method, adjust sample to a convenient volume in a graduated cylinder or Imhoff
cone. Concentrating the plankton by using a rubber bulb and clear acrylic plastic tube with fine
mesh netting fitted on the end is convenient and accurate (Figure 10200:9). For picoplankton and
the smaller microzooplankton, use sedimentation techniques described for concentrating
phytoplankton. Transfer sample to a beaker or other wide-mouth vessel for subsampling with a
Hensen-Stempel or similar wide-bore pipet. Gently stir sample completely and randomly with
the pipet and quickly withdraw 1 to 5 mL. Transfer to a suitable counting chamber.
Alternatively, subsample by splitting with any of a number of devices of which the Folsom
plankton splitter1 is best known (Figure 10200:10). Level splitter before using. Place sample in
the splitter and divide into subsplits. Rinse splitter into the subsamples. Repeat until a workable
number (200 to 500 individuals) is obtained in a subsample. Exercise care to provide unbiased
splits. Even when using the Folsom splitter unbiased subsamples cannot be unquestioningly
assumed;2 therefore, count animals in several subsamples from the same sample to verify that
the splitter is unbiased and to determine the sampling error introduced by using it.
Another method permits abundance estimates of more equivalent levels of precision among
taxa than obtained with either the Hensen-Stempel pipet or the Folsom splitter.3 Normal
counting procedures tally organisms on the basis of their abundance in a sample. Therefore, in a
sample with a dominant organism making up 50% of total numbers, the tally of the dominant
taxon will be large and have a small error. However, error about the subdominants will increase
as the tally of each taxon decreases. By accepting one level of precision, the technique3 has been
developed to obtain the same error about dominants and subdominants, permitting quantitative
comparisons between taxa over successive times or between stations.
2. Enumeration
Using a compound microscope and a magnification of 100×, enumerate small zooplankton
(protozoa, rotifers, and nauplii) in a 1- to 5-mL clear acrylic plastic counting cell fitted with a
glass cover slip. For larger, mature microcrustacea use a counting chamber holding 5 to 10 mL.
A Sedgwick-Rafter cell is not suitable because of size. An open counting chamber 80 by 50 mm
and 2 mm deep is desirable; however, an open chamber is difficult to move without jarring and
disrupting the count. A mild detergent solution placed on the chamber before counting reduces
organism movements or special counting trays with parallel or circular grooves or partitions4,5
can be used. Count microcrustacea with a binocular dissecting microscope at 20× to 40×
magnification. If identification is questionable, remove organisms with a microbiological
transfer loop and examine at a higher magnification under a compound microscope.
Report smaller zooplankton as number per liter and larger forms as number per cubic meter:
where:
C=
V′ =
V′′ =
V′′′ =
volume of the concentrated sample, mL,
volume counted, mL, and
volume of the grab sample, m3.
To obtain organisms per liter divide by 1000.
3. References
1. LONGHURST, A.R. & D.L.R. SEIBERT. 1967. Skill in the use of Folsom’s plankton sample
splitter. Limnol. Oceanogr. 12:334.
2. MCEWEN, G.F., M.W. JOHNSON & T.R. FOLSOM. 1954. A statistical analysis of the
Folsom sample splitter based upon test observations. Arch. Meteorol. Geophys.
Bioklimatol., Ser. A, 6:502.
3. ALDEN, R.W., III, R.C. DAHIYA & R.J. YOUNG, JR. 1982. A method for the enumeration of
zooplankton samples. J. Exp. Mar. Biol. Ecol. 59:185.
4. GANNON, J.E. 1971. Two counting cells for the enumeration of zooplankton
micro-crustacea. Trans. Amer. Microsc. Soc. 90:486.
5. DODSON, A.N. & W.H. THOMAS. 1964. Concentrating plankton in gentle fashion.
10200 H.
Chlorophyll
The concentration of photosynthetic pigments is used extensively to estimate phytoplankton
biomass.1,2 All green plants contain chlorophyll a, which constitutes approximately 1 to 2% of
the dry weight of planktonic algae. Other pigments that occur in phytoplankton include
chlorophylls b and c, xanthophylls, phycobilins, and carotenes. The important chlorophyll
degradation products found in the aquatic environment are the chlorophyllides, pheophorbides,
and pheophytins. The presence or absence of the various photosynthetic pigments is used, among
other features, to separate the major algal groups.
The three methods for determining chlorophyll a in phytoplankton are the
spectrophotometric,3-5 the fluorometric,6-8 and the high-performance liquid chromatographic
(HPLC) techniques.9 Fluorometry is more sensitive than spectrophotometry, requires less
sample, and can be used for in-vivo measurements.10 These optical methods can significantly
under- or overestimate chlorophyll a concentrations,11-18 in part because of the overlap of the
absorption and fluorescence bands of co-occurring accessory pigments and chlorophyll
degradation products.
Pheophorbide a and pheophytin a, two common degradation products of chlorophyll a, can
interfere with the determination of chlorophyll a because they absorb light and fluoresce in the
same region of the spectrum as does chlorophyll a. If these pheopigments are present, significant
errors in chlorophyll a values will result. Pheopigments can be measured either by
spectrophotometry or fluorometry, but in marine and freshwater environments the fluorometric
method is unreliable when chlorophyll b co-occurs. Upon acidification of chlorophyll b, the
resulting fluorescence emission of pheophytin b is coincident with that of pheophytin a, thus
producing underestimation and overestimation of chlorophyll a and pheopigments, respectively.
HPLC is a useful method for quantifying photosynthetic pigments9,13,15,16,19-21 including
chlorophyll a, accessory pigments (e.g., chlorophylls b and c), and chlorophyll degradation
products (chlorophyllides, pheophorbides, and pheophytins). Pigment distribution is useful for
quantitative assessment of phytoplankton community composition and zooplankton grazing
activity.22
1. Pigment Extraction
Conduct work with chlorophyll extracts in subdued light to avoid degradation. Use opaque
containers or wrap with aluminum foil. The pigments are extracted from the plankton
concentrate with aqueous acetone and the optical density (absorbance) of the extract is
determined with a spectrophotometer. The ease with which the chlorophylls are removed from
the cells varies considerably with different algae. To achieve consistent complete extraction of
the pigments, disrupt the cells mechanically with a tissue grinder.
Glass fiber filters are preferred for removing algae from water. The glass fibers assist in
breaking the cells during grinding, larger volumes of water can be filtered, and no precipitate
forms after acidification. Inert membrane filters such as polyester filters may be used where
these factors are irrelevant.
a. Equipment and reagents:
1) Tissue grinder:*#(118) Successfully macerating glass fiber filters in tissue grinders with
grinding tube and pestle of conical design may be difficult. Preferably use round-bottom
grinding tubes with a matching pestle having grooves in the TFE tip.
2) Clinical centrifuge.
3) Centrifuge tubes, 15-mL graduated, screw-cap.
4) Filtration equipment, filters, glass fiber†#(119) or membrane (0.45-µm porosity, 47-mm
diam); vacuum pump; solvent-resistant disposable filter assembly, 1.0-µm pore size;‡#(120)
10-mL solvent-resistant syringe.
5) Saturated magnesium carbonate solution: Add 1.0 g finely powdered MgCO3 to 100 mL
distilled water.
6) Aqueous acetone solution: Mix 90 parts acetone (reagent-grade BP 56°C) with 10 parts
saturated magnesium carbonate solution. For HPLC pigment analysis, mix 90 parts HPLC-grade
acetone with 10 parts distilled water.
b. Extraction procedure:
1) Concentrate sample by centrifuging or filtering as soon as possible after collection. If
processing must be delayed, hold samples on ice or at 4°C and protect from exposure to light.
Use opaque bottles because even brief exposure to light during storage will alter chlorophyll
values. Samples on filters taken from water having pH 7 or higher may be placed in airtight
plastic bags and stored frozen for 3 weeks. Process samples from acidic water promptly after
filtration to prevent possible chlorophyll degradation from residual acidic water on filter. Use
glassware and cuvettes that are clean and acid-free.
2) Place sample in a tissue grinder, cover with the 2 to 3 mL 90% aqueous acetone solution,
and macerate at 500 rpm for 1 min. Use TFE/glass grinder for a glass-fiber filter and glass/glass
grinder for a membrane filter.
3) Transfer sample to a screw-cap centrifuge tube, rinse grinder with a few milliliters 90%
aqueous acetone, and add the rinse to the extraction slurry. Adjust total volume to 10 mL, with
90% aqueous acetone. Use solvent sparingly and avoid excessive dilution of pigments. Steep
samples at least 2 h at 4°C in the dark. Glass fiber filters of 25- and 47-mm diam§#(121) have
dry displacement volumes of 0.03 and 0.10 mL, respectively, and introduce errors of about 0.3
and 1.0% if a 10-mL extraction volume is used.
4) Clarify by filtering through a solvent-resistant disposable filter (to minimize retention of
extract in filter and filter holder, force 1 to 2 mL air through the filter after the extract), or by
centrifuging in closed tubes for 20 min at 500 g. Decant clarified extract into a clean, calibrated,
15-mL, screw-cap centrifuge tube and measure total volume. Proceed as in 2, 3, 4, or 5 below.
2. Spectrophotometric Determination of Chlorophyll
1) Spectrophotometer, with a narrow band (pass) width (0.5 to 2.0 nm) because the
chlorophyll absorption peak is relatively narrow. At a spectral band width of 20 nm the
chlorophyll a concentration may be underestimated by as much as 40%.
2) Cuvettes, with 1-, 4-, and 10-cm path lengths.
3) Pipets, 0.1- and 5.0-mL.
4) Hydrochloric acid, HCl, 0.1N.
b. Determination of chlorophyll a in the presence of pheophytin a: Chlorophyll a may be
overestimated by including pheopigments that absorb near the same wavelength as chlorophyll
a. Addition of acid to chlorophyll a results in loss of the magnesium atom, converting it to
pheophytin a. Acidify carefully to a final molarity of not more than 3 × 10−3M to prevent certain
accessory pigments from changing to absorb at the same wavelength as pheophytin a.13 When a
solution of pure chlorophyll a is converted to pheophytin a by acidification, the
absorption-peak-ratio (OD664/OD665) of 1.70 is used in correcting the apparent chlorophyll a
concentration for pheophytin a.
Samples with an OD664 before/OD665 after acidification ratio (664b/665a) of 1.70 are
considered to contain no pheophytin a and to be in excellent physiological condition. Solutions
of pure pheophytin show no reduction in OD665 upon acidification and have a 664b/665a ratio
of 1.0. Thus, mixtures of chlorophyll a and pheophytin a have absorption peak ratios ranging
between 1.0 and 1.7. These ratios are based on the use of 90% acetone as solvent. Using 100%
acetone as solvent results in a chlorophyll a before-to-after acidification ratio of about 2.0.3
Spectrophotometric procedure—Transfer 3 mL clarified extract to a 1-cm cuvette and read
optical density (OD) at 750 and 664 nm. Acidify extract in the cuvette with 0.1 mL 0.1N HCl.
Gently agitate the acidified extract and read OD at 750 and at 665 nm, 90 s after acidification.
The volumes of extract and acid and the time after acidification are critical for accurate,
consistent results.
The OD664 before acidification should be between 0.1 and 1.0. For very dilute extracts use
cuvettes having a longer path length. If a larger cell is used, add a proportionately larger volume
of acid. Correct OD obtained with larger cuvettes to 1 cm before making calculations.
Subtract the 750-nm OD value from the readings before (OD 664 nm) and after acidification
(OD 665 nm).
Using the corrected values calculate chlorophyll a and pheophytin a per cubic meter as
follows:
where:
V1 = volume of extract, L,
V2 = volume of sample, m3,
L = light path length or width of cuvette, cm, and
664b, 665a = optical densities of 90% acetone extract before and after acidification,
respectively.
The value 26.7 is the absorbance correction and equals A × K
where:
A = absorbance coefficient for chlorophyll a at 664 nm = 11.0, and
K = ratio expressing correction for acidification.
c. Determination of chlorophyll a, b, and c (trichromatic method): Spectrophotometric
procedure—Transfer extract to a 1-cm cuvette and measure optical density (OD) at 750, 664,
647, and 630 nm. Choose a cell path length or dilution to give OD664 between 0.1 and 1.0.
Use the optical density readings at 664, 647, and 630 nm to determine chlorophyll a, b, and
c, respectively. The OD reading at 750 nm is a correction for turbidity. Subtract this reading
from each of the pigment OD values of the other wavelengths before using them in the equations
below. Because the OD of the extract at 750 nm is very sensitive to changes in the
acetone-to-water proportions, adhere closely to the 90 parts acetone:10 parts water (v/v) formula
for pigment extraction. Turbidity can be removed easily by filtration through a disposable,
solvent-resistant filter attached to a syringe or by centrifuging for 20 min at 500 g.
Calculate the concentrations of chlorophyll a, b, and c in the extract by inserting the
corrected optical densities in following equations:5
a) Ca = 11.85(OD664) − 1.54(OD647) − 0.08(OD630)
b) Cb = 21.03(OD647) − 5.43(OD664) − 2.66(OD630)
c) Cc = 24.52(OD630) − 7.60(OD647) − 1.67(OD664)
where:
Ca, Cb, and Cc = concentrations of chlorophyll a, b, and c, respectively, mg/L, and
OD664, OD647,
and OD630 = corrected optical densities (with a 1-cm light path) at the respective
wavelengths.
After determining the concentration of pigment in the extract, calculate the amount of
pigment per unit volume as follows:
3. Fluorometric Determination of Chlorophyll a
The fluorometric method for chlorophyll a is more sensitive than the spectrophotometric
method and thus smaller samples can be used. Calibrate the fluorometer spectrophotometrically
with a sample from the same source to achieve acceptable results. Optimum sensitivity for
chlorophyll a extract measurements is obtained at an excitation wavelength of 430 nm and an
emission wavelength of 663 nm. A method for continuous measurement of chlorophyll a in vivo
is available, but is reported to be less efficient than the in-vitro method given here, yielding
about one-tenth as much fluorescence per unit weight as the same amount in solution.
Pheophytin a also can be determined fluorometrically.24
a. Equipment and reagents: In addition to those listed under 1a and 2a above:
Fluorometer,i#(122) equipped with a high-intensity F4T.5 blue lamp, photomultiplier tube
R-446 (red-sensitive), sliding window orifices 1×, 3×, 10×, and 30×, and filters for light
emission (CS-2-64) and excitation (CS-5-60). A high-sensitivity door is preferable.
b. Extraction procedure: Prepare sample as directed in 1b above.
1) Calibrate fluorometer with a chlorophyll solution of known concentration as follows:
Prepare chlorophyll extract and analyze spectrophotometrically. Prepare serial dilutions of the
extract to provide concentrations of approximately 2, 6, 20, and 60 µg chlorophyll a/L. Make
fluorometric readings for each solution at each sensitivity setting (sliding window orifice): 1×,
3×, 10×, and 30×. Using the values obtained, derive calibration factors to convert fluorometric
readings in each sensitivity level to concentrations of chlorophyll a, as follows:
where:
Fs = calibration factor for sensitivity setting S,
Rs = fluorometer reading for sensitivity setting S, and,
C′a = concentration of chlorophyll a determined spectrophotometrically, µg/L.
2) Measure sample fluorescence at sensitivity settings that will provide a midscale reading.
(Avoid using the 1× window because of quenching effects.) Convert fluorescence readings to
concentrations of chlorophyll a by multiplying the readings by the appropriate calibration factor.
c. Determination of chlorophyll a in the presence of pheophytin a: This method normally is
not applicable to freshwater samples. See discussion under Section 10200H and ¶ 2b above.
1) Equipment and reagents—In addition to those listed under ¶ 1a and ¶ 2a above, pure
chlorophyll a##(123) (or a plankton chlorophyll extract with a spectrophotometric
before-and-after acidification ratio of 1.70 containing no chlorophyll b).
2) Fluorometric procedure—Calibrate fluorometer as directed in ¶ 3b1). Determine extract
fluorescence at each sensitivity setting before and after acifidication. Calculate calibration
factors (Fs) and before-and-after acidification fluorescence ratio by dividing fluorescence
reading obtained before acidification by the reading obtained after acidification. Avoid readings
on the 1× scale and those outside the range of 20 to 80 fluorometric units.
3) Calculations—Determine the ‘‘corrected’’ chlorophyll a and pheophytin a in sample
extracts with the following equations:8,24
where:
Fs = conversion factor for sensitivity setting S (see ¶ 2b, above),
Rb = fluorescence of extract before acidification,
Ra = fluorescence of extract after acidification,
r = Rb/Ra, as determined with pure chlorophyll a for the instrument (redetermine r
and Fs if filters or light source are changed),
Ve = volume of extract, and
Vs = volume of sample.
d. Extraction of whole water, nonfiltered samples: Alternatively, to prevent cell lysis during
filtration, extract whole water sample.
1) Equipment and reagents—Fluorometer equipped with a high-sensitivity R928
phototube**#(124) with output impedance of 36 ma/W at 675 nm and a high-sensitivity door.
Place neutral density filter (40–60N) in the rear light path,††#(125) selected to permit reagent
blanking on the highest sensitivity scale.
2) Extraction procedure—Decant 1.5 mL sample into screw-cap test tube and add 8.5 mL
100% acetone. Mix with vortex mixer and hold in the dark for 6 h at room temperature. Filter
through glass fiber filter‡‡#(126) or centrifuge. Measure fluorescence as described in Section
10200H.3 and estimate concentrations as in ¶ 3c. Because humic substances interfere, if they are
present filter a sample portion (see Section 10200H.1b) and process filtrate with sample.
Subtract filtrate (blank) fluorescence from that of sample.
4. High-Performance Liquid Chromatographic Determination of Algal Chlorophylls and
Their Degradation Products
a. Equipment and reagents: In addition to those listed for pigment extraction, ¶ 1a above:
1) High-pressure liquid chromatograph capable of a flow rate of 2.0 mL/m.
2) High-pressure injector valve equipped with a 100-µL sample loop.
3) Guard column (4.0 × 0.5 cm, C18 packing material, 3-µm particle size, or equivalent
protection system) for extending life of primary column.
4) Reverse-phase HPLC column.§§#(127)
5) Fluorescence detector capable of excitation at 430 ± 30 nm and measuring emission at
wavelengths greater than 600 nm.
6) Data recorder device: Strip chart recorder or, preferably, an electronic integrator.
7) Syringe, glass, 250-µL.
8) HPLC eluents: System A (80:15:5; methanol:reagent water: ion-pairing solution) and
System B (80:20; methanol:acetone). Use HPLC-grade solvents; measure volumes before
mixing. Filter eluents through a solvent-resistant 0.4-µm filter before use and degas with helium.
Prepare the ion-pairing (IP) solution from 15 g tetrabutylammonium acetatei i#(128) and 77 g
ammonium acetate###(129) made up to 1 L with reagent water.15
9) Calibration standards: Individually dissolve 1 mg each pure chlorophyll a and bi i#(130)
in 100 mL 90% acetone. Determine the exact concentrations spectrophotometrically (ε664 for
chlorophyll a in 90% acetone = 87.67 L g−1 cm−1; ∈647 for chlorophyll b in 90% acetone =
51.36 L g−1 cm−1).5 Prepare pheophytin a + a′ and b + b′ standards from the primary chlorophyll
a and b standards by acidification with hydrochloric acid; correct respective concentrations for
Mg2+ loss. Extract chlorophyll c with 90% acetone from diatoms, purify by thin-layer
chromatography (TLC)25 and calibrate spectrophotometrically (ε631 for a mixture containing
equal amounts of chlorophylls c1 and c2 in 90% acetone containing 1% pyridine = 42.6 L g−1
cm−1; the absence of this small amount of pyridine is presumed to cause only small differences
in the absorption properties of chlorophyll c.26 Alternatively, determine the chlorophyll c
content of a 90% acetone extract made from diatoms, spectrophotometrically (chlorophyll c1 +
c2, µg/mL = 24.36E630 − 3.73E664)5 and use as standard. Prepare chlorophyllide a from
diatoms,27 purify by TLC25 and calibrate spectrophotometrically in 90% acetone (ε664 for
chlorophyllide a = 128 L g−1 cm−1).28 Prepare pheophorbide a by acidification of chlorophyllide
a, purify by TLC,25 and calibrate spectrophotometrically in 90% acetone (∈665 for pheophorbide
a = 69.8 L g−1 cm−1).28 Standards stored under nitrogen in the dark at −20°C are stable for about
1 month.
b. Procedure:
1) Set up and equilibrate the HPLC system with solvent System A at a flow rate of 2
mL/min. Adjust fluorometer sensitivity to provide full-scale reading with the most concentrated
chlorophyll a standard.
2) Calibrate HPLC system by preparing working standards from the primary standards (on
day of use). Once retention times of the standards are determined for a particular system,
simplify standardization by preparing serial dilutions from mixed standards. Prepare separately
mixed standards for the chlorophylls and chlorophyllide a and for the pheophytins and
pheophorbide a. Mix 1-mL portions of standards with 300 µL ion-pairing solutions and
equilibrate for 5 min before injection (use of ion-pairing agents greatly enhances separation of
dephytolated pigments, chlorophyllide a, chlorophyll c, and pheophorbide a). Prepare blanks by
mixing 1 mL 90% acetone with 300 µL IP solution. Rinse syringe twice with 150 µL standard
and draw about 250 µL standard into syringe for injection. Place syringe in injector valve,
overfilling the 100-µL sample loop. Construct calibration curves by plotting fluorescence peak
areas (or heights) against standard pigment concentrations.
3) Prepare samples for injection by mixing a 1-mL portion of the 90% acetone pigment
extract with 300 µL IP solution.
4) Use a two-step solvent program to optimize separation of the chorophylls from their
degradation products.15 After injection, change from solvent System A to System B over 5 min
and follow with System B for 15 min at a flow rate of 2 mL/min. Re-equilibrate the column with
System A for 5 min before the next injection for a total analysis time of approximately 25 min.
Degas the solvent systems with helium during analysis. Increase lifetime of HPLC column by
storing it in 100% methanol between runs. Periodically flush the HPLC system with reagent
water to avoid buildup of ion pairing agents.
5) Calculate individual pigment concentrations using the following formula:
where:
Ci =
As =
Fi =
peak area).
VI =
VE =
VS =
individual pigment concentration, mg/L,
area of individual pigment peak from sample injection,
standard response factor (mg pigment/0.1 mL standard divided by corresponding
injection volume (0.1 mL),
extraction volume, mL, and
sample volume, L.
6) This method is designed only for quantification of chlorophylls and their degradation
products. Detect carotenoid pigments, which also are present in 90% acetone extracts but do not
fluoresce, by absorbance spectroscopy (at about 440 nm).21
7) The elution order and approximate retention times for the major chlorophyll pigments and
their degradation products are shown in Figure 10200:11. The detection limits (s/n = 2) vary
with fluorometer configuration and flow rate; however, they range from 10 to 100 pg per
injection for most chlorophylls and their degradation products.15,21,29 The accuracy of the HPLC
method depends primarily on purity of pigment standards. Preferably measure absorption spectra
(350 to 750 nm) of the standards and compare with published data. Pigment purity also can be
assessed by HPLC analysis, providing there are no co-eluting contaminants with absorption and
fluorescence bands overlapping those of the standards. HPLC and spectrophotometrically
derived pigment concentrations for available EPA standards agree reasonably well (± 20%) if
spectrophotometric results are corrected for the presence of pheopigments and the HPLC results
are expressed as pigment equivalents (e.g., chlorophyll a equivalents = chlorophyllide a +
chlorophyll a + chlorophyll a′, provided that the proper molecular weight corrections are
applied).30 Thus, if significant amounts of chlorophyll derivatives are present, pigment
concentrations determined spectrophotometrically will be overestimated. The agreement
between HPLC and fluorometrically derived results depends on the presence of accessory
chlorophylls b, c, and their derivatives. Triplicate injections of a fivefold dilution of an EPA
sample gave coefficients of variation of 7.5% (chlorophyllide a), 9.1% (chlorophyll c), 13.4%
(pheophorbide a), 9.6% (chlorophyll b), 0.5% (chlorophyll a), 6.2% (pheophytin a), and 22.9%
(pheophytin a′), with an average value of 10% for the seven pigments analyzed.
5. High-Performance Liquid Chromatographic Determination of Algal Chlorophyll and
Carotenoid Pigments (PROPOSED)
a. Equipment and reagents: In addition to those listed for pigment extraction, ¶ 1a above:
1) High-performance liquid chromatographic pump capable of gradient delivery of three
different solvents at a flow rate of 1 mL/min.
2) High-pressure injector valve equipped with a 200-µL sample loop.
3) Guard column (50 × 4.6 mm, C18 packing material,***#(131) 5-µm particle size) for
extending life of primary column.
4) Reverse-phase HPLC column with endcapping (250 × 4.6 mm, 5-µm particle size, C18
column***#(132)).
5) Variable wavelength or filter absorbance detector with low-volume flowthrough cell.
Detection wavelength is 436 nm.
6) Data recording device: Strip chart recorder or, preferably, an electronic integrator or
computer equipped with hardware and software for chromatographic data analysis.
7) Syringe, glass, 500-µL.
8) HPLC eluents: Eluent A (80:20, v:v; methanol:0.5M ammonium acetate, pH 7.2); Eluent
B (90:10, v:v; acetonitrile:water), and Eluent C, ethyl acetate. Use HPLC-grade solvents.
Measure volumes before mixing. Filter eluents through a solvent-resistant 0.4-µm filter before
use and degas with helium.
9) Calibration standards: Chlorophylls a and b, and β,β-carotene can be
purchased†††#(133) as can zeaxanthin and lutein.‡‡‡#(134) Other pigment standards can be
purified from plant extracts by thin-layer chromatography (TLC)25 or preparative-scale HPLC.
Determine concentration of all standards using a monochromator-based spectrophotometer in the
appropriate solvents before calibration of the HPLC system. The recommended extinction
coefficients for most common algal pigments found in freshwater systems are given in Table
10200:III. Measure absorbance in a 1-cm cuvette at the appropriate wavelength (usually at λmax)
and 750 nm (to correct for light scattering). Calculate concentrations of standards as follows:
where:
Ci = individual pigment concentration, mg/L,
A = absorbance at specific wavelength,
E1cm = weight-specific absorption coefficient, L g−1 cm−1,
b = pathlength of cuvette, cm, and
1000 = conversion factor, g to mg.
Standards stored under nitrogen in the dark at −20°C are stable for about 1 month.
b. Procedure:
1) Set up and equilibrate the HPLC system with Eluent A at a flow rate of 1 mL/min.
2) Calibrate the HPLC using working standards (about 0 to 1000 ng/mL) prepared from
primary standards on day of use. Mix 1 mL standard with 300 µL distilled water, shake, and
equilibrate for 5 min before injection (diluting standards and sample extracts with water
increases affinity of pigments for the column in the loading step, resulting in an improved
separation of more polar pigments). Rinse syringe twice with 300 µL standard and draw 500 µL
standard into syringe for injection. Place syringe in injector valve, overfilling the 200 µL sample
loop 2.5-fold. To check for possible interferences in the extraction solvent and/or filter, prepare a
blank by extracting a glass fiber filter in 90% acetone; mixing 1 mL 90% acetone filter extract
and 300 µL distilled water; and injecting into the HPLC system. Plot absorbance peak areas (or
heights) against standard pigment concentrations. Calculate response factors as the slope of the
regression between the weights of the injected standards (ng) and the areas of the parent pigment
(plus areas of structurally-related isomers if present). These isomers contribute to the absorption
signal of the standards; disregarding them results in over-estimation of pigments in sample
extracts.31
3) Prepare samples for injection by mixing a 1 mL portion of the 90% acetone pigment
extract and 300 µL distilled water, shake, and equilibrate for 5 min before injection.
4) Following sample injection, use a gradient program to optimize separation of chlorophyll
and carotenoid pigments. The system described in Table 10200:IV has been developed from the
original method32 to insure elution of most hydrophobic pigments. Degas solvent system with
helium during analysis. Periodically flush HPLC system with distilled water to avoid
accumulation of ion-pairing reagents.
5) Routinely determine peak identities by comparing retention times of sample peaks with
those of pure standards. Confirm peak identities spectrophotometrically by collecting eluting
peaks from the column outlet (or directly with an on-line diode array spectrophotometer).
Absorption maxima for most common pigments found in freshwater systems are given in Table
10200:III.
6) Calculate individual pigment concentrations using the formula given in ¶ 4b5) preceding.
7) This method is designed for separation of chlorophyll and carotenoid pigments (Figure
10200:12), however, it also separates major chlorophyll breakdown products.
8) Method precision was assessed by making triplicate injections of a mixture of
phytoplankton and plant extracts. Coefficients of variation ranged from 0.6 to 6.0% (Table
10200:III). Using an appropriate internal standard increases precision.
6. References
1. ROTT, E. Spectrophotometric and chromatographic chlorophyll analysis: comparison of
results and discussion of the trichromatic method. Ergebn. Limnol. (Suppl. to Arch.
Hydrobiol.) 14:37.
2. MARKER, A.F.H., E.A. NUSCH, H. RAI & B. RIEMANN. 1980. The measurement of
photosynthetc pigments in freshwaters and standardization of methods: Conclusions
and recommendations. Ergebn. Limnol. (Suppl. to Arch. Hydrobiol.) 14:91.
3. LORENZEN, C.J. 1967. Determination of chlorophyll and pheo-pigments:
spectrophotometric equations. Limnol. Oceanogr. 12:343.
4. FITZGERALD, G.P. & S.L. FAUST. 1967. A spectrophotometric method for the estimation
of percentage degradation of chlorophylls to pheopigments in extracts of algae. Limnol.
Oceanogr. 12:335.
5. JEFFREY, S.W. & G.F. HUMPHREY. 1975. New spectrophotometric equations for
determining chlorphylls a, b, and c, in higher plants, algae and natural phytoplankton.
Biochem. Physiol. Pflanzen 167: 191.
6. YENTSCH, C.S. & D.W. MENZEL. 1963. A method for the determination of phytoplankton
chlorophyll and phaeophytin by fluorescence. Deep Sea Res. 10:221.
7. LOFTUS, M.E. & J.H. CARPENTER. 1971. A fluorometric method for determining
chlorophylls a, b, and c. J. Mar. Res. 29:319.
8. HOLM-HANSEN, O., C.J. LORENZEN, R.W. HOLMES & J.D.H. STRICKLAND. 1965.
Fluorometric determination of chlorophyll. J. Cons. Cons. Perma. Int. Explor. Mer
30:3.
9. ABAYCHI, J.K. & J.P. RILEY. 1979. The determination of phytoplankton pigments by
high-performance liquid chromatography. Anal. Chim. Acta 107:1.
10. LORENZEN, C.J. 1966. A method for the continous measurement of in vivo chlorophyll
concentration. Deep Sea Res. 13:223.
11. JACOBSEN, T.R. 1978. A quantitative method for the separation of chlorophylls a and b
from phytoplankton pigments by high-pressure liquid chromatography. Mar. Sci.
Comm. 4:33.
12. BROWN, L.M., B.T. HARGRAVE & M.D. MACKINNON. 1981. Analysis of chlorophyll a in
sediments by high-performance liquid chromatography. Can. J. Fish. Aquat. Sci.
38:205.
13. GIESKES, W.W. & G.W. KRAAY. 1983. Unknown chlorophyll a derivatives in the North
Sea and the tropical Atlantic Ocean revealed by HPLC analysis. Limnol. Oceanogr.
28:757.
14. GOWEN, R.J., P. TETT & B.J.B. WOOD. 1983. Changes in the major dihydroporphyrin
plankton pigments during the spring bloom of phytoplankton in two Scoottish
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
sea-lochs. J. Mar. Biol. Assoc. U.K. 63:27.
MANTOURA, R.F.C. & C.A. LLWEWLLYN. 1983. The rapid determination of algal
chlorophyll and caroteniod pigments and their breakdown products in natural waters by
reverse-phase high-performance liquid chromatography. Anal. Chim. Acta 151:297.
GIESKES, W.W.C. & G.W. KRAAY. 1984. Phytoplankton, its pigments, and primary
production at a central North Sea station in May, July and September 1981. Neth. J.
Sea Res. 18.71.
HALLEGRAEFF, G.M. & S.E. JEFFREY. 1985. Description of new chlorophyll a alteration
products in marine phytoplankton. Deep Sea Res. 32:697.
TREES, C.C., M.C. KENNICUTT II & J.M. BROOKS. 1985. Errors associated with the
standard fluorometric determination of chlorophylls and phaeopigments. Mar. Chem.
17:1.
ESKINS, K., C.R. SCHOFIELD & H.J. DUTTON. 1977. High-performance liquid
chromatography of plant pigments. J. Chromatogr. 135:217.
WRIGHT, S.W. & J.D. SHEARER. 1984. Rapid extraction and high performance liquid
chromatography of chlorophylls and carotenoids from marine phytoplankton. J.
Chromatogr. 294:281.
BIDIGARE, R.R., M.C. KENNICUTT II & J.M. BROOKS. 1985. Rapid determination of
chlorophylls and their degradation products by high-performance liquid
chromatography. Limnol. Oceanogr. 30:432.
JEFFRY, S.W. 1974. Profiles of photosynthetic pigments in the ocean using thin-layer
chromatography. Mar. Biol. 26:101.
PHINNEY, D.A. & C.S. YENTSCH. 1985. A novel phytoplankton chlorophyll technique:
toward automated analysis. J. Plankton Res. 7: 633.
STRICKLAND, J.D.H. & T.R. PARSONS. 1968. A Practical Manual of Sea Water Analysis.
JEFFREY, S.W. 1981. An improved thin-layer chromatographic technique for marine
phytoplankton pigments. Limnol. Oceanogr. 26:191.
JEFFREY, S.W. 1972. Preparation and some properties of crystalline chlorophyll c1 and
c2 from marine algae. Biochim. Biophys. Acta 279: 15.
27. BARRETT, J. & S.W. JEFFREY. 1971. A note on the occurrence of chlorophyllase in
marine algae. J. Exp. Mar. Biol. Ecol. 7:255.
28. LORENZEN, C.J. & J. NEWTON DOWNS. 1986. Specific absorption coefficients of
chlorophyllide a and pheophorbide a in 90 percent acetone, and comments on the
fluorometric determination of chlorophyll and pheopigments. Limnol. Oceanogr.
31:449.
29. SARTORY, D.P. 1985. The determination of algal chlorophyllous pigments by high
performance liquid chromatography and spectrophotometry. Water Res. 19:605.
30. MURRAY, A.P., C.F. GIBBS & A.R. LONGMORE. 1986. Determination of chlorophyll in
marine waters: Intercomparison of a rapid HPLC method with full HPLC,
spectrophotometric and fluorometric methods. Mar. Chem. 19:211.
31. BIDIGARE, R.R. 1991. Analysis of algal chlorophylls and carotenoids. In D.C. Hurd &
D.W. Spencer, eds., Marine Particles: Analysis and Characterization. America
Geophysical Union, Washington, D.C.
32. WRIGHT, S.W., S.W. JEFFREY, R.F.C. MANTOURA, C.A. LLEWELLYN, T. BJORNLAND, D.
REPETA & N. WELSCHMEYER. 1991. Improved HPLC method for the analysis of
chlorophylls and carotenoids from marine phytoplankton. Mar. Ecol. Prog. Ser.
77:183.
33. JEFFREY, S.W. & F.T. HAXO. 1968. Photosynthetic pigments of symbiotic dinoflagellates
(zooxanthallae) from corals and clams. Biol. Bull. 135:149.
34. JENSEN, A. 1978. Chlorophylls and carotenoids. In J.A. Helleburst & J.S. Craige, eds.,
Handbook of Phycological Methods: Physiological and Biochemical Methods.
Cambridge University Press, Cambridge, England.
35. DAVIS, B.H. 1976. Carotenoids. In T.W. Goodwin, ed., Chemistry and Biochemistry of
Plant Pigments. Academic Press, New York, N.Y.
36. JOHANSEN, J.E., W.A. SVEC & S. LIAAEN-JENSEN. 1974. Carotenoids of the Dinophyceae.
Phytochem. 13:2261.
10200 I.
Determination of Biomass (Standing Crop)
Biomass is a quantitative estimate of the total mass of living organisms within a given area
or volume. It may include the mass of a population (species biomass) or of a community
(community biomass) but gives no information on community structure or function. The most
accurate methods for estimation of biomass are dry weight, ash-free dry weight, and volume of
living organisms. Indirect methods include estimates of total carbon, caloric content, nitrogen,
lipids, carbohydrates, silica (diatoms), and chlorophyll (algae). Adenosine triphosphate1 (ATP)
and deoxyribonucleic acid2,3 (DNA) also have been used as indirect estimates. All estimates of
biomass can be affected by the presence of organic and inorganic detritus; ATP and DNA
analyses include contributions from the bacterial flora.4
1. Chlorophyll a
Chlorophyll a is used as an algal biomass indicator.5 Assuming that chlorophyll a
constitutes, on the average, 1.5% of the dry weight of organic matter (ash-free weight) of algae,
estimate the algal biomass by multiplying the chlorophyll a content by a factor of 67.
2. Biovolume (Cell Volume)
Plankton data derived on a volume-per-volume basis often are more useful than numbers per
milliliter.6 Determine cell volume by using the simplest geometric configuration that best fits the
shape of the cell being measured (such as sphere, cone, cylinder).7 Cell sizes of an organism can
differ substantially in different waters and from the same waters at different times during the
year; therefore, average measurements from 20 individuals of each species for each sampling
period. Calculate the total biovolume of any species by multiplying the average cell volume in
cubic micrometers by the number per milliliter.
Compute total wet algal volume as:
where:
Vt = total plankton cell volume, mm3/L,
Ni = number of organisms of the ith species/L, and
Vi = average volume of cells of ith species, µm3.
3. Cell Surface Area
An estimation of cell surface area is valuable in analyzing interactions between the cell and
surrounding waters. Compute average surface area in square micrometers and multiply by the
number per milliliter of the species being considered.
4. Displacement Volume
This method8 measures an equivalent volume of liquid that is displaced by the sample.
Displacement volume may be determined by several methods; for simple, direct measurement
proceed as follows: Place sample in sieve of mesh size equal to or smaller than net used in
capture; let sample drain and transfer to a measured volume of water in a graduated cylinder;
measure the new volume containing sample plus known volume. The displacement volume
equals the new volume minus original measured volume of water.
5. Gravimetric Methods
The biomass of the plankton community can be estimated from gravimetric determinations,
although silt and organic detritus interfere. Determine dry weight by placing 100 mg wet
concentrated sample in a clean, ignited, and tared porcelain crucible and dry at 105°C for 24 h.
Alternatively, filter a known volume of sample through 0.45-µm-pore-diam membrane or a
prerinsed, dried, and preweighed glass-fiber filter. (Note that the small sample used in direct
filtration may lead to error if not handled properly.) Cool sample in a desiccator and weigh.
Obtain ash weight by igniting the dried sample at 500°C for 1 h. Cool, rewet ash with distilled
water, and bring to constant weight at 105°C. The ash is rewetted to restore water of hydration of
clays and other minerals; this may amount to as much as 10% of weight lost during
incineration.9 The ash-free dry weight is the difference between the dry weight and the weight of
the ash residue after ashing. The ash-free dry weight is preferred to dry weight to compare mixed
assemblages. The ash content may constitute 50% or more of the dry weight in phytoplankton
having inorganic structures, such as the diatoms. In other forms the ash content is only about 5%
of dry weight.
6. Adenosine Triphosphate (ATP)
Methods of measuring adenosine triphosphate (ATP) in plankton provide the only means of
determining the total viable plankton biomass. ATP occurs in all plants and animals, but only in
living cells; it is not associated with nonliving particulate material. The ratio of ATP to biomass
varies from species to species, but appears to be constant enough to permit reliable estimates of
biomass from ATP measurements.10 The method is simple and relatively inexpensive and the
instrumentation is stable and reliable. The method also has many potential applications in
entrainment and bioassay work, especially plankton mortality studies.
1) Glassware: clean, sterile, dry borosilicate glass flasks, beakers, and pipets.
2) Filters: 47-mm-diam, 0.45-µm-porosity membrane filters.
3) Filtration equipment.
4) Freezer (−20°C).
5) Boiling water bath.
6) Detection instruments designed specifically for measuring ATP.*#(135)
7) Microsyringes: 10-, 25-, 50-, 100-, and 250-µL.
8) Reaction cuvettes and vials.
9) Tris buffer (0.02M, pH 7.75): Dissolve 7.5 g trishydroxymethylaminomethane in 3 L
distilled water and adjust to pH 7.75 with 20% HCl. Autoclave 150-mL portions at 115°C for 15
min.
10) Luciferin-luciferase enzyme preparation:†#(136) Rehydrate frozen (−20°C) lyophilized
extracts of firefly lanterns with Tris buffer as directed by the supplier; let stand at room
temperature 2 to 3 h, then centrifuge at 300 × g for 1 min and decant the supernatant into a clean,
dry test tube; let stand at room temperature for 1 h.
11) Purified ATP standard: Dissolve 12.3 mg disodium ATP in 1 L distilled water and dilute
1.0 mL to 100 mL with Tris buffer; 0.2 mL = 20 ng ATP.
b. Procedure:
1) Calibration—To determine the calibration factor (F), prepare a series of dilutions of
purified ATP standard and record the light emission from several portions of each concentration
of standard. Correct mean area of standards by subtracting peak reading or mean area of several
blanks using 0.2 mL Tris buffer. Calculate calibration factor FS as:
where:
FS = calibration factor at sensitivity S,
AS = peak reading or mean area under standard ATP curve corrected for blank, and
C = concentration of ATP in standard solution, ng/mL.
2) Sample analysis—Collect a 1- to 2-L sample in a clean, sterile sampler. Pass through a
250-µm net to remove large zooplankton10 and filter through a 47-mm 0.45-µm-porosity filter
by applying a vacuum of about 30 kPa. (IMPORTANT: Break the suction before the last film of
water is pulled through the filter.) Quickly place filter in a small beaker. Immediately cover filter
with 3 mL boiling Tris buffer, using an automatic pipet. Place beaker in boiling water bath for 5
min and, with a Pasteur pipet, transfer extract to a clean, dry, calibrated test tube. Rinse filter and
beaker with 2 mL boiling Tris buffer; combine extracts, record volume, bring volume up to 5 mL
with Tris buffer, cover tubes with parafilm and, if samples cannot be analyzed immediately,
freeze at −25°C. Extracts may be stored for many months in a freezer. Prepare at least triplicate
extracts of each sample.
The analytical procedure depends on detection equipment used. If a scintillation counter is
used, pipet 0.2 mL enzyme preparation into a glass vial. Measure the light emission of the
enzyme preparation (blank) for 2 to 3 min at sensitivity settings near that anticipated for the
sample. Add 0.2 mL sample extract to the vial, record the time, and swirl. Start recording light
output 10 s after combining ATP extract and enzyme preparation; record output for 2 to 3 min,
using the same time period for all samples. Determine the mean of areas under the curves
obtained and correct by subtracting mean of areas under the curves obtained from blanks
prepared as directed in Strickland and Parsons.11
c. Calculations: Calculate concentration of ATP as:
Ac =
Ve =
Vs =
Fs =
mean corrected area under extract curves,
extract volume, mL,
volume of sample, L, and
calibration factor.
If an ATP content of 2.4 µg ATP/mg dry weight organic matter is assumed,12 total living
plankton biomass (B), as dry weight organic matter, is given as:
7. References
1. HOLM-HANSEN, O. & C.R. BOOTH. 1966. The measurement of adenosine triphosphate in
the ocean and its ecological significance. Limnol. Oceanogr. 11:510.
2. HOLM-HANSEN, O., W.H. SUTCLIFFE, JR. & J. SHARP. 1968. Measurement of
deoxyribonucleic acid in the ocean and its ecological significance. Limnol. Oceanogr.
13:507.
3. HOLM-HANSEN, O. 1969. Determination of microbial biomass in ocean profiles. Limnol.
Oceanogr. 14:740.
4. PAERL, H.W., M.M. TILZER & C.R. GOLDMAN. 1976. Chlorophyll a vs. ATP as algal
biomass indicators in lakes. J. Phycol. 12:242.
5. CREITZ, G.I. & F.A. Richards. 1955. The estimation and characterization of plankton
populations by pigment analysis. J. Mar. Res. 14: 211.
6. KUTKUHN, J.H. 1958. Notes on the precision of numerical and volumetric plankton
estimates from small sample concentrations. Limnol. Oceanogr. 3:69.
7. VOLLENWEIDER, R.A. 1969. A Manual on Methods for Measuring Primary Production
in Aquatic Environments. IBP Handbook No. 12. Blackwell Scientific Publ., Oxford,
England.
8. JACOBS, F. & G.C. GRANT. 1978. Guidelines for zooplankton sampling in quantitative
baseline and monitoring programs. EPA-600/3-78-026, U.S. Environmental Protection
Agency.
9. NELSON, D.J. & D.C. SCOTT. 1962. Role of detritus in the productivity of a rock-outcrop
community in a Piedmont stream. Limnol. Oceanogr. 7:396.
10. RUDD, J.W.M. & R.D. HAMILTON. 1973. Measurement of adenosine triphosphate (ATP)
in two precambrian shield lakes of northwestern Ontario. J. Fish. Res. Board Can.
30:1537.
11. STRICKLAND, J.D.H. & T.R. PARSONS. 1968. A Practical Manual of Sea Water Analysts.
12. WEBER, C.I. 1973. Recent developments in the measurement of the response of
plankton and periphyton to changes in their environment. In G. Glass, ed. Bioassay
Techniques and Environmental Chemistry. Ann Arbor Science Publ., Ann Arbor,
Mich.
10200 J.
Metabolic Rate Measurements
The physiological condition and the spectrum of biological interactions of the aquatic
community must be considered for evaluation of the state of natural waters. Earlier, numbers,
species composition, and biomass were the prime considerations. Recognition of the limitations
of this approach led to the measurement of rates of metabolic processes such as photosynthesis
(productivity), nitrogen fixation, respiration, and electron transport. These provide a better
understanding of the complex nature of the aquatic ecosystem. An indication of photosynthetic
efficiency can be determined by the productivity index (mg C fixed/unit chlorophyll a).1
1. Nitrogen Fixation
The ability of an organism to fix nitrogen is a great competitive advantage and plays a major
role in population dynamics. Two reliable methods for estimating nitrogen fixation rates in the
laboratory are the 15N isotope tracer method2,3 and the acetylene reduction method.4 Because
the rate of nitrogen fixation varies greatly with different organisms and with the concentration of
combined nitrogen, nitrogen fixation rates cannot be used to estimate biomass of nitrogen-fixing
organisms. However, the acetylene reduction method is useful in measuring nitrogen budgets
and in algal assay work.5
2. Productivity, Oxygen Method
Productivity is defined as the rate at which inorganic carbon is converted to an organic form.
Chlorophyll-bearing organisms (phytoplankton, periphyton, macrophytes) serve as primary
producers in the aquatic food chain. Photosynthesis ultimately results in the formation of a wide
range of organic compounds, release of oxygen, and reduction of carbon dioxide (CO2) in the
surrounding waters. Primary productivity6 can be determined by measuring the changes in
oxygen and CO2 concentrations.7 In poorly buffered waters, pH can be a sensitive property for
detecting variations in the system. As CO2 is removed during photosynthesis, the pH rises. This
shift can be used to estimate both photosynthesis and respiration.8 The sea and many fresh
waters are too highly buffered to make this useful, but it has been applied successfully to
productivity studies in some lake waters.
Two methods of measuring the rate of carbon uptake and net photosynthesis in situ are the
oxygen method9 and the carbon 14 method.10 In both methods, clear (light) and darkened (dark)
bottles are filled with water samples and suspended at regular depth intervals for an incubation
period of several hours or samples are incubated under controlled conditions in environmental
growth chambers in the laboratory.
The basic reactions in algal photosynthesis involve uptake of inorganic carbon and release of
oxygen, summarized by the relationship:
CO2 + H2O → (CH2O) + O2
The chief advantages of the oxygen method are that it provides estimates of gross and net
productivity and respiration and that analyses can be performed with inexpensive laboratory
equipment and common reagents. The dissolved oxygen (DO) concentration is determined at the
beginning and end of the incubation period. Productivity is calculated on the assumption that one
atom of carbon is assimilated for each molecule of oxygen released.
a. Equipment:
1) BOD bottles, numbered, 300-mL, clear and opaque borosilicate glass, with ground glass
stopper and flared mouth, for sample incubation. Acid-clean the bottles, rinse thoroughly with
distilled water, and just before use, rinse with water being tested. Do not use
phosphorus-containing detergents.
If suitable opaque bottles are not available, make clear BOD bottles opaque by painting them
black and wrapping with black waterproof tape. As a further precaution, wrap entire bottle in
aluminum foil or place in light-excluding container during incubation.
2) Supporting line or rack that does not shade suspended bottles.
3) Nonmetallic opaque acrylic Van Dorn sampler or equivalent, of 3- to 5-L capacity.
4) Equipment and reagents for dissolved oxygen determinations: See Section 4500-O.
5) Pyrheliometer.
6) Submarine photometer.
7) Thermometer.
b. Procedure:
1) Obtain a profile of the input of solar radiation for the photoperiod with a pyrheliometer.
2) Determine depth of euphotic zone (the region that receives 1% or more of surface
illumination) with a submarine photometer. Select depth intervals for bottle placement. The
photosynthesis-depth curve will be approximated closely by placing samples at intervals equal to
one-tenth the depth of the euphotic zone. Estimate productivity in relatively shallow water with
fewer depth intervals.
3) Measure oxygen concentration with probe or by titration and temperature and salinity to
determine whether water is supersaturated with respect to oxygen (see Table 4500-O:I). If water
is supersaturated, bubble nitrogen gas through sample to lower initial oxygen concentration to
less than 80% saturation.
4) Keep samples out of direct sunlight during handling. Introduce samples taken from each
preselected depth into duplicate clear, darkened, and initial-analysis bottles. Insert delivery tube
of sampler to bottom of sample bottle and fill so that three volumes of water are allowed to
overflow. Remove tube slowly and close bottle. Use water from the same grab sample to fill a
‘‘set’’ (one light, one dark, and one initial bottle).
5) Immediately treat (fix) samples taken for the chemical determination of initial dissolved
oxygen (see Section 4500-O) with manganous sulfate (MnSO4), alkaline iodide, and sulfuric
acid (H2SO4) or check with an oxygen probe. Analyses may be delayed several hours if
necessary, if samples are fixed or iced and stored in the dark.
6) Suspend duplicate paired clear and darkened bottles at the depth from which the samples
were taken and incubate for at least 2 h, but never longer than it takes for oxygen-gas bubbles to
form in the clear bottles or DO to be depleted in the dark bottles.
7) At the end of the exposure period, immediately determine DO as described above.
c. Calculations: The increase in oxygen concentration in the light bottle during incubation is
a measure of net production which, because of the concurrent use of oxygen in respiration, is
somewhat less than the total (or gross) production. The loss of oxygen in the dark bottle is used
as an estimate of total plankton respiration. Thus:
Net photosynthesis = light bottle DO − initial DO
Respiration = initial DO − dark bottle DO
Gross photosynthesis = light bottle DO − dark bottle DO
Average results from duplicates.
1) Calculate the gross or net production for each incubation depth and plot:
mg carbon fixed/m3 = mg oxygen released/L × 12/32 × 1000 L/m3 × K
where K is the photosynthetic quotient (PQ), ranging from 1 to 2, depending on the nitrogen
supply.11,12
Use the factor 12/32 to convert oxygen to carbon; under ideal conditions 1 mole of O2 (32 g)
is released for each mole of carbon (12 g) fixed.
2) Productivity is defined as the rate of production and generally is reported in grams carbon
fixed per square meter per day. Determine the productivity of a vertical column of water 1 m
square by plotting productivity for each exposure depth and graphically integrating the area
under the curve.
3) Using the solar radiation profile and photosynthesis rate during incubation adjust the data
to represent phytoplankton productivity for the entire photoperiod. Because photosynthetic rates
vary widely during the daily cycle,13,14 do not attempt to convert data to other test
circumstances.
3. Productivity, Carbon 14 Method
A solution of radioactive carbonate (14CO32–) is added to light and dark bottles that have
been filled with sample as described for the oxygen method. After incubation in situ, collect the
plankton on a membrane filter, treat with hydrochloric acid (HCl) fumes to remove inorganic
carbon 14, and assay for radioactivity. The quantity of carbon fixed is proportional to the
fraction of radioactive carbon assimilated.
This procedure differs from the oxygen method in that it affords a direct measurement of
carbon uptake and measures only net photosynthesis.15 It is basically more sensitive than the
oxygen method, but fails to account for organic materials that leach from cells16,17 during
incubation.
1) Pyrheliometer.
2) Submarine photometer.
3) BOD bottles and supporting apparatus: See ¶ 2a1) and 2), above.
4) Membrane-filtering device and 25-mm filters with pore diameters of 0.22, 0.30, 0.45, 0.80,
and 1.2 µm.
5) Counting equipment for measuring radioactivity: Scaler with end-window tube, gas flow
meter, or liquid scintillation counter (see Section 7030B). The thin-window tube is the least
expensive detector and, when used with a small scaler, provides acceptable data at modest cost.
6) Fuming chamber: Use a glass desiccator with a depth of about 1.4 cm conc HCl in
desiccant chamber. The fuming chamber is recommended for filter decontamination.18,19
7) Syringe or pipet, nonmetallic.
8) Chemical reagents: See Section 4500-CO2 (Carbon Dioxide) and Section 2320
(Alkalinity).
9) Radioactive carbonate solutions:
a) Sodium chloride dilution solution, 5% NaCl (w/v): Add 0.3 g sodium carbonate (Na2CO3)
and one pellet sodium hydroxide (NaOH) per liter. Use for marine studies only.
b) Carrier-free radioactive carbonate solution, commercially available in sealed vials having
approximately 5 µCi 14C/mL. Confirm absence of suspended and dissolved toxic metals20 or
filter and pass through an ion-exchange column.*#(137)
c) Working solutions with activities of 1, 5, and 25 µCi 14C/2 mL. For studies of fresh water
use carrier-free radioactive carbonate and for studies of marine water prepare by diluting
carrier-free radioactive carbonate solution with NaCl dilution solution.
d) Stock ampules: Prepare ampules containing 2 mL of required working solution. Fill
ampules and autoclave sealed ampules at 121°C for 20 min.21
b. Procedure:
1) Obtain a record of incident solar radiation for the photoperiod with a pyrheliometer.
2) Determine depth intervals for sampling and incubation as described above.
3) Use duplicate light and dark bottles at each depth. Also use dark bottles or bottles
harvested at time zero. Fill bottles with sample, add 2 mL radioactive carbonate solution (using a
nonmetallic pipet) to the bottom of each bottle, and mix thoroughly by repeated inversion. The
concentration of carbon 14 should be approximately 10 µCi/L in relatively productive waters, to
100 µCi/L, or higher, in oliogotrophic (open ocean) waters. To obtain statistical significance,
have at least 1000 cpm in the filtered sample. Take duplicate samples at each depth to determine
initial concentration of inorganic carbon (CO2, HCO3–, and CO32–) available for photosynthesis
(see Section 4500-CO2). For estuarine and marine samples, estimate total inorganic carbon
concentrations with a simple titration procedure22 and make initial temperature, salinity, and pH
measurements.
4) Incubate samples for up to 4 h. If measurements are required for the entire photoperiod,
overlap 4-h periods from dawn until dusk. A 4-h incubation period may be sufficient provided
energy input is used as the basis for integrating incubation period to entire photoperiod. For
incubation procedure, see ¶ 2b6) above.
5) After incubating remove sample bottles and immediately place in the dark. Filter
unpreserved samples without delay. Avoid sample preservation to avoid lysing cells or
determine extracellular products.
6) Filter two portions of each sample through a membrane filter, taking care that the largest
pore size is consistent with quantitative retention of plankton. Although the 0.45-µm pore filter
usually is adequate, determine the efficiency of sample retention immediately before analysis,
with a wide range of pore sizes.23,24 Apply approximately 30 kPa of vacuum during filtration.
Excess vacuum may cause extensive cell rupture and loss of radioactivity through the
membrane.25 Use maximum sample volume consistent with rapid filtration (1 to 2 min), but do
not clog filter.
7) Place membranes in HCl fumes for 20 min. Count filters as soon as possible, although
extended storage in a desiccator is acceptable.
8) Determine radioactivity by counting with an end-window tube, windowless gas flow
detector, or liquid scintillation counter.
9) Determine counting geometry of thin-window and windowless gas flow detectors.26 Using
three ampules of carbon 14, prepare a series of barium carbonate (BaCO3) precipitates on tared
0.45-µm membrane filters as directed below. The precipitates will contain the same amount of
carbon 14 activity but will have different thicknesses ranging from 0.5 to 6.0 mg/cm2. Dilute
each ampule to 500 mL with a solution of 1.36 g Na2CO3/L CO2-free distilled water. Pipet
0.5-mL portions into each of seven conical flasks containing 0, 0.5, 1.5, 2.5, 3.5, 4.5, and 5.5
mL, respectively, of a solution of 1.36 g Na2CO3 /L CO2-free distilled water. Add, respectively,
0.3, 0.6, 1.2, 1.8, 2.4, 3.0, and 3.6 mL 1.04% barium chloride (BaCl2) solution. Let BaCO3
precipitate stand 2 h with gentle swirling every half hour. Collect each precipitate on a filter
(using an apparatus with a filtration area comparable to that of the samples). With suction, dry
filters without washing; place in a desiccator for 24 h, weigh, and count. The counting rate
increases exponentially with decreasing precipitate thickness. Extrapolate graphically (or
mathematically) to zero precipitate thickness and multiply the zero-thickness counting rate by
1000 to correct for ampule dilution. This represents the amount of activity added to each sample
bottle used to determine fraction of carbon 14 taken up in light and dark bottles.
c. Calculations:
1) Subtract the mean dark-bottle or time-zero sample count from the mean light-bottle counts
for each replicate pair.
2) Determine the total dissolved inorganic carbon available for photosynthesis (carbonate,
bicarbonate, and free CO2) from pH and alkalinity measurements; make direct measurement of
total CO2 according to Section 4500-CO2 or the methods described in the literature.27-30
3) Determine quantity of carbon fixed by using the following relationship:
4) Integrate productivity for the entire depth of euphotic zone and express as grams carbon
fixed per square meter per day [see ¶ 2c2) above].
5) Using the solar radiation records and photosynthesis rates during incubation, adjust data to
represent phytoplankton productivity for the entire photoperiod. If samples were incubated for
less than the full photoperiod, apply a correction factor.
4. References
1. GUNDERSEN, K. 1973. In-situ determination of primary production by means of the new
incubator, ISIS. Helgolander wiss. Meeresunters. 24:465.
2. BURRIS, R.H., F.J. EPPLING, H.B. WAHLIN & P.W. WILSON. 1942. Studies of biological
nitrogen fixation with isotopic nitrogen. Proc. Soil Sci. Soc. Amer. 7:258.
3. NEESS, J.C., R.C. DUGDALE, V.A. DUGDALE & J.J. GOERING. 1962. Nitrogen metabolism
in lakes. I. Measurement of nitrogen fixation with N15. Limnol. Oceanogr. 7:163.
4. STEWART, W.D.P., G.P. FITZGERALD & R.H. BURRIS. 1967. In situ studies on N2 fixation
using the acetylene reduction technique. Proc. Nat. Acad. Sci. 58:2071.
5. STEWART, W.D.P., G.P. FITZGERALD & R.H. BURRIS. 1970. Acetylene reduction assay for
determination of phosphorus availability in Wisconsin lakes. Proc. Nat. Acad. Sci.
66:1104.
6. GOLDMAN, C.R. 1968. Aquatic primary production. Amer. Zoologist 8:31.
7. ODUM, H.T. 1957. Primary production measurements in eleven Florida springs and a
marine turtle-grass community. Limnol. Oceanogr. 2:85.
8. BEYERS, R.J. & H.T. ODUM. 1959. The use of carbon dioxide to construct pH curves for
the measurements of productivity. Limnol. Oceanogr. 4:499.
9. GAARDER, T. & H.H. GRAN. 1927. Investigations of the production of plankton in Oslo
Fjord. Rapp. Proces-Verbaux. Reunions Cons. Perma. Int. Explor. Mer 42:1.
10. STEEMAN-NEILSEN, E. 1952. The use of radioactive carbon (C-14) for measuring
organic production in the sea. J. Cons. Perma. Int. Explor. Mer 18:117.
11. WILLIAMS, P.J. LEB., R.C.T. RAINE & J.R. BRYAN. 1979. Agreement between the 14C and
oxygen methods of measuring phytoplankton production: Reassessment of the
photosynthetic quotient. Oceanol. Acta 2:411.
12. DAVIES, J.M. & P.J. LEB. WILLIAMS. 1984. Verification of 14C and O2 derived primary
organic production using an enclosed system. J. Plankton Res. 6:457.
13. RYTHER, J.H. 1956. Photosynthesis in the ocean as a function of light intensity. Limnol.
Oceanogr. 1:61.
14. FEE, E.J. 1969. A numerical model for the estimation of photosynthetic production,
integrated over time and depth, in natural waters. Limnol. Oceanogr. 14:906.
15. STEEMAN-NEILSEN, E. 1964. Recent advances in measuring and understanding marine
primary production. J. Ecol. 52(Suppl.):119.
16. ALLEN, M.B. 1956. Excretion of organic compounds by Chlamydomonas. Arch.
Mikrobiol. 24:163.
17. FOGG, G.E. & W.D. WATT. 1965. The kinetics of release of extracellular products of
photosynthesis by phytoplankton. In C.R. Goldman, ed. Primary Productivity in
Aquatic Environments. Suppl. 18, Univ. California Press, Berkeley.
18. WETZEL, R.G. 1965. Necessity for decontamination of filters in C14 measured rates of
photosynthesis in fresh waters. Ecology 46:540.
19. MCALLISTER, C.D. 1961. Decontamination of filters in the C14 method of measuring
marine photosynthesis. Limnol. Oceanogr. 6:447.
20. CARPENTER, E.J. & J.S. LIVELY. 1980. Review of estimates of algal growth using 14C
tracer techniques. In P.G. Falkowski, ed. Primary Productivity in the Sea. Brookhaven
Symp. Biol. No. 31. Plenum Press, New York, N.Y.
22. PARSONS, T.R., Y. MAITA & C.M. LALLI. 1984. A Manual of Chemical and Biological
Methods for Seawater Analysis. Pergamon Press, New York, N.Y.
23. LASKER, R. & R.W. HOLMES. 1957. Variability in retention of marine phytoplankton by
membrane filters. Nature 180:1295.
24. HOLMES, R.W. & C.G. ANDERSON. 1963. Size fractionation of C14-labelled natural
phytoplankton communities. In C.H. Oppenheimer, ed. Symposium on Marine
Microbiology. Charles C. Thomas, Springfield, Ill.
25. ARTHUR, C.R. & F.H. RIGLER. 1967. A possible source of error in the C14 method of
measuring primary productivity. Limnol. Oceanogr. 12:121.
26. JITTS, H.R. & B.D. SCOTT. 1961. The determination of zero-thickness activity in Geiger
counting of C14 solutions used in marine productivity studies. Limnol. Oceanogr.
6:116.
27. SAUNDERS, G.W., F.B. TRAMA & R.W. BACHMANN. 1962. Publ. No. 8, Great Lakes
Research Div., Univ. Michigan, Ann Arbor.
28. DYE, J.F. 1944. The calculation of alkalinities and free carbon dioxide in water by use
of nomographs. J. Amer. Water Works Assoc. 36:859.
29. MOORE, E.W. 1939. Graphic determination of carbon dioxide and the three forms of
alkalinity. J. Amer. Water Works Assoc. 31:51.
30. PARK, K., D.W. HOOD & H.T. ODUM. 1958. Diurnal pH variation in Texas bays and its
application to primary production estimations. Publ. Inst. Mar. Sci. Univ. Tex. 5:47.
10300
PERIPHYTON*#(138)
10300 A.
Introduction
1. Definition and Significance
Microorganisms growing on stones, sticks, aquatic macrophytes, and other submerged
surfaces are useful in assessing the effects of pollutants on lakes, streams, and estuaries. Included
in this group of organisms, here designated periphyton,1,2 are the zoogleal and filamentous
bacteria, attached protozoa, rotifers, and algae, and the free-living microorganisms that swim,
creep, or lodge among the attached forms.
Unlike the plankton, which often do not respond fully to the influence of pollution in rivers
for a considerable distance downstream, the periphyton show marked responses immediately
below pollution sources. Examples are the beds of Sphaerotilus and other ‘‘slime organisms’’
commonly observed in streams below discharges of organic wastes. Because the abundance and
composition of the periphyton at a given location are governed by the water quality at that point,
observations of their condition generally are useful in evaluating conditions in bodies of water.
The use of periphyton in assessing water quality often is hindered by the lack of suitable
natural substrates at the desired sampling station. Furthermore, it often is difficult to collect
quantitative samples from these surfaces. To circumvent these problems artificial substrates have
been used to provide a uniform surface type, area, and orientation.3
2. References
1. ROLL, H. 1939. Zur Terminologie des Periphytons. Arch. Hydrobiol. 35:39.
2. YOUNG, O.W. 1945. A limnological investigation of periphyton in Douglas Lake,
Michigan. Trans. Amer. Microsc. Soc. 64:1.
3. SLADECKOVA, A. 1962. Limnological investigation methods for the periphyton
(‘‘Aufwuchs’’) community. Bot. Rev. 28:286.
10300 B.
Sample Collection
1. Station Selection
In rivers, locate stations a short distance upstream and at one or more points downstream
from the suspected pollution source or intended study area in the areas of central mixing. In large
rivers, sample both sides of the stream in main flow areas. Because the effects of a pollutant
depend on the assimilative capacity of the stream and on the nature of the pollutant, progressive
changes in water quality downstream from the pollution source may be caused entirely by
dilution and cooling—as in the case of nutrients, toxic industrial wastes, and thermal
pollution—or by gradual mineralization of degradable organic compounds. Cursory examination
of shoreline and bottom periphyton growths on natural substrates downstream from an outfall
may indicate conspicuous zones of biological response to water quality that will be useful in
determining appropriate sites for sampling stations. When an intensive sampling program is not
feasible, a minimum of three sampling stations, one in a reference area upstream from a pollution
source and the others in the community downstream from the source, where complete mixing
with the receiving water has occurred, will provide minimal data on the periphyton community.
In lentic waters (e.g., lakes, reservoirs, ponds) and other standing-water bodies where zones
of pollution may be arranged concentrically, locate stations in areas adjacent to a waste outfall
and in unaffected areas. Use control stations similar to the affected ones (e.g., similar in water
depth and distance from shore).
a. Natural substrates: Collect qualitative samples by scraping submerged stones, sticks,
pilings, and other available substrates. Many devices have been developed to collect quantitative
samples from irregular surfaces. Appropriate techniques for the removal of periphyton from both
living and nonliving surfaces have been described.1-4
b. Artificial substrates: The most widely used artificial substrate is the standard, plain, 25- by
75-mm glass microscope slide, but other materials such as clear vinyl plastic also are suitable.
Do not change substrate type during a study because colonization varies with substrate. In small,
shallow streams and in the littoral regions of lakes and reservoirs where light penetrates to the
bottom, place slides or other substrates vertically in frames anchored to the bottom. In large,
deep streams or standing-water bodies where turbidity varies widely, place slides vertically with
the slide face at right angles to the prevailing current. A floating rack, as shown in Figure
10300:1,*#(139) is suitable. Expose several slides (minimally five; three for biomass, one for
species, and one backup for each time interval) for each type of analysis to assure collecting
sufficient material and to determine variability in results caused by normal differences in
colonization of individual slides. In addition to effects of pollutants, length of substrate exposure
and seasonal changes in temperature and other natural environmental conditions may have a
profound effect on sample composition. No community on an artificial substrate is representative
of the natural community.
Place, expose, and handle all artificial substrate samplers in conditions as nearly identical as
possible, whether they are replicate samplers at a particular sampling location or samplers at
different locations. Sampler type and/or construction cause changes in surrounding physical
conditions that in turn affect periphyton growth. Variations of 10 to 25% between sample
replicates are common. Therefore, to reduce sampling error and increase interpretive power,
reduce the magnitude of all possible test variables and use sufficient replication.
c. Exposure period: Colonization on clean slides proceeds at an exponential rate for the first
1 or 2 weeks and then slows. Because exposures of less than 2 weeks may result in very sparse
collections, and exposures of more than 2 weeks may result in loss of material due to sloughing,
sample for 2 weeks during the summer. This exposure period precludes collecting sexually
mature thalli of larger, slow-growing filamentous algae such as Cladophora and Stigeoclonium.
To obtain optimum growth during the winter, use a longer exposure period. For the most
exacting work, determine the optimum exposure period by testing colonization rates over a
period of about 6 weeks.
Secondary problems associated with macroinvertebrate infestation and grazing may occur,
often within 7 to 14 d. To reduce the confounding influence of grazing, increase substrate
sampling area and expose for 7 to 10 d.
3. Sample Preservation
Preserve samples that are taken for counting and identification in 5% neutralized formalin,
Lugol’s iodine, or merthiolate (see Section 10200B.2).
Preserve slides intact in bottles of suitable size or scrape into containers in the field. Air-dry
slides for dry and ash-free dry weight in the field and store in a 3.0- × 7.7-cm glass bottle. Place
slides for chlorophyll analyses in acetone or methanol in the field or collect and freeze with
trichlorotrifluoroethane†#(140) (or alternative) or CO2 and hold on dry ice until returned to the
laboratory. Store all samples in the dark.
4. References
1. SLADECKOVA, A. 1962. Limnological investigation methods for the periphyton
(‘‘Aufwuchs’’) community. Bot. Rev. 28:286.
2. GOUGH, S.B. & W.J. WOELKERLING. 1976. On the removal and quantification of algal
aufwuchs from macrophyte hosts. Hydrobiologia 48: 203.
3. BOOTH, W.E. 1981. A method for removal of some epiphytic diatoms. Botanica Marina
24:603.
4. DELBECQUE, E.J.P. 1985. Periphyton on nymphaeids: An evaluation of methods and
separation techniques. Hydrobiologia 124:85.
5. Bibliography
COOKE, W.B. 1956. Colonization of artificial bare areas by microorganisms. Bot. Rev. 22:613.
HOHN, M.H. 1966. Artificial substrate for benthic diatoms—collection, analysis, and
interpretation. In K.W. Cummings, C.A. Tryon, Jr., & R.T. Hartman, eds.
Organism-Substrate Relationships in Streams. Spec. Publ. No. 4, p. 87. Pymatuning Lab.
Ecology, Univ. Pittsburgh, Pittsburgh, Pa.
KEVERN, N.R., J.L. WILHM & G.M. VAN DYNE. 1966. Use of artificial substrata to estimate the
productivity of periphyton communities. Limnol. Oceanogr. 11:499.
ARTHUR, J.W. & W.B. HORNING. 1969. The use of artificial substrates in pollution surveys. Amer.
Midland Natur. 82:83.
TIPPETT, R. 1970. Artificial surfaces as a method of studying populations of benthic micro-algae
in fresh water. Brit. Phycol. J. 5:187.
ERTL, M. 1971. A quantitative method of sampling periphyton from rough substrates. Limnol.
Oceanogr. 16:576.
ANDERSON, M.A. & S.L. PAULSON. 1972. A simple and inexpensive woodfloat periphyton
sampler. Progr. Fish-Cult. 34:225.
NORTH AMERICAN ENTHOLOGICAL SOCIETY. 1974–1991. (Annual) Current and Select
Bibliographies on Benthic Biology. Springfield, Ill.
MARKER, A.F.H., C.A. CROWTHER & R.J.M. GUNN. 1980. Methanol and acetone as solvents for
estimating chlorophyll a and phaeopigments by spectrophotometry. Arch. Hydrobiol. Ergebn.
Limnol. 14:52.
NEROZZI, A. & P. SILVER. 1983. Periphytic community analysis in a small oligotropic lake. Proc.
Penn. Acad. Sci. 57:138.
WETZEL, R., ed. 1983. Periphyton of Freshwater Ecosystems. Developments in Hydrobiology
17. Dr. W. Junk BV Publishers, The Hague, The Netherlands.
HAMILTON, P.B. & H.C. DUTHIE. 1984. Periphyton colonization of rock surfaces in a boreal forest
stream studied by scanning electron microscopy and track autoradiography. J. Phycol.
20:525.
NIELSEN, T.S., W.H. FUNK, H.L. GIBBONS & R.M. DUFFNER. 1984. A comparison of periphyton
growth on artificial and natural substrates in the Upper Spokane River, Washington, USA.
Northwest Sci. 58: 243.
PIP, E. & G.G.C. ROBINSON. 1984. A comparison of algal periphyton composition on 11 species of
submerged macrophytes. Hydrobiol. Bull. 18:109.
POULIN, M., L. BERARD-THERRIAULT & A. CARDINAL. 1984. Benthic diatoms from hard
substrates of marine and brackish waters of Quebec Canada 3. Fragilarioideae, Fragilariales,
Fragilariaceae. Nat. Can. (Que). 111:349.
STEVENSON, R.J. 1984. How currents on different sides of substrates in streams affect
mechanisms of benthic algal accumulation. Int. Rev. ges. Hydrobiol. 69:241.
VYMAZAL, J. 1984. Short-term uptake of heavy metals by periphytic algae. Hydrobiologia
119:171.
AUSTIN, A. & J. DENISEGER. 1985. Periphyton community changes along a heavy metals gradient
in a long narrow lake. Environ. Exper. Bot. 25:41.
FLOWER, R.J. 1985. An improved epilithon sampler and its evaluation in two acid lakes. Brit.
Phycol. J. 20:109.
LAMBERTI, G.A. & V.H. RESH. 1985. Comparability of introduced tiles and natural substrates for
sampling lotic bacteria, algae, and macroinvertebrates. Freshwater Biol. 15:21.
PIEKARCZYK, R. & E. MCARDLE. 1985. Pioneer colonization and interaction of photosynthetic
and heterotrophic microorganisms on an artificial substrate of polyurethane foam in E.J.
Beck Lake, Illinois, USA. Trans. Ill. State Acad. Sci. 78:81.
10300 C.
Sample Analysis
1. Sedgwick-Rafter Counts
Remove periphyton from slides with a razor blade and rubber policeman. Disperse
scrapings in 100 mL or other suitable volume of preservative with vigorous shaking, or use a
blender. Transfer a 1-mL portion to a Sedgwick-Rafter cell, and make a strip count as described
in Section 10200F.2a. If material in the Sedgwick-Rafter cell is too dense to count directly,
discard and replace with a diluted sample.
Sedgwick-Rafter cells do not permit examination at magnifications higher than 200×. The
Palmer cell1, a thinner version of the S-R cell, permits examination at 400 to 500× with a
standard compound microscope.
Express counts as cells or filaments per square millimeter of substrate area, calculated as in ¶
C.2.
2. Inverted Microscope Method Counts
Using an inverted microscope for periphyton counts permits magnifications higher than
those possible with the Sedgwick-Rafter cell. Remove periphyton quantitatively from slides
with a razor blade and policeman. Transfer a measured portion, after serial dilution if necessary,
into a standardized plankton sedimentation chamber. After a suitable period of settling (see
Section 10200C.1), count organisms in the settling chamber by counting all organisms within a
known number of strips or random fields. Calculate algal density per unit area of substrate as
follows:
where:
N = number of organisms counted,
At = total area of chamber bottom, mm2,
Vt = total volume of original sample suspension, mL,
Ac = area counted (strips or fields), mm2,
Vs = sample volume used in chamber, mL, and
As = surface area of slide or substrate, mm2.
Separation of periphyton from silt and detritus may be enhanced by adding a drop or less of a
saturated iodine solution to the counting chamber just before counting. This method is especially
useful when Chlorophyta are the predominant organisms because iodine stains starch food
reserves blue. Iodine can be added even to preserved samples.
3. Diatom Species Counts
Preparation of permanent diatom mounts from periphyton samples differs from preparation
of mounts from plankton samples because of the need to remove extracellular organic matter
(such as gelatinous materials). If this organic matter is not removed it will produce a thick brown
or black carbonaceous deposit on the cover glass when the sample is incinerated. Clear organic
matter by incineration by placing a small, known volume of sample (< 1 mL) directly on a cover
slip. Let water evaporate and ash at 525°C (not more) for 6 to 10 min. Mount cover slip for
direct examination of diatom frustules. Alternatively, decompose organic substances by
oxidation with ammonium persulfate or with HNO3 or 30% H2O2 and K2Cr2O7 (see Section
10200D.3) before mounting sample. To oxidize with persulfate place a measured sample of
approximately 5 mL in a disposable 10-mL vial. Let stand 24 h, withdraw supernatant liquid by
aspiration, replace with a 5% solution of (NH4)2S2O8, and mix thoroughly. Do not exceed a total
volume of 8 mL. Heat vial to approximately 90°C for 30 min. Let stand 24 h, withdraw
supernatant liquid, and replace with reagent-grade water. After three changes of reagent-grade
water, with a disposable pipet transfer a drop of the diatom suspension to a cover glass,
evaporate to dryness, and prepare and count a mount as described for plankton (Section 10200).
Count as least 500 frustules and express results as relative numbers or percentage of each species
per unit area.
4. Stained Sample Preparation and Counting
Staining periphyton samples permits distinguishing algae from detritus and ‘‘live’’ from
‘‘dead’’ diatoms. This distinction is especially important because periphyton often contains
many dead diatoms of planktonic as well as periphytic origin.
In the first method, cells are exposed to a vital stain to evaluate the percentages of live,
senescent, and dead algae, particularly diatoms, by estimating relative metabolic activities. The
colorless tetrazolium violet is reduced in the cytochrome system of metabolically active cells to
form violet-colored triphenylformazan. When cells are senescent or dead, the reaction fails.
Make tetrazolium violet solution by adding 2.0 g tetrazolium violet to 1.0 L water. The
solution may be buffered to a pH of 7.5 to 7.7 with tris-hydroxymethyl amine. Add 1 mL
tetrazolium violet solution to 9 mL sample and incubate 2 to 4 h at room temperature. Count
diatom frustules and other cells (at least 300/sample) and place into the following categories: a)
active: violet precipitate observed within the cell or mitochondria; b) senescent: chlorophyll
present, but no violet precipitate; c) dead: no chlorophyll or violet precipitate present.
In the second method, all algal components of periphyton may be studied in one preparation,
without sacrificing detailed diatom taxonomy.2 This method yields permanent slides for
reference collections.
Thoroughly mix preserved samples in the preservative solution. Prepare acid fuchsin stain by
dissolving 1 g acid fuchsin in 100 mL reagent-grade water, adding 2 mL glacial acetic acid, and
filtering. Place a measured sample in a centrifuge tube with 10 to 15 mL acid fuchsin stain. Mix
sample and stain several times during a 20-min staining period; centrifuge at 1000 g for 20 min.
Decant stain, being careful not to disturb sediment, or siphon off supernatant. Add 10 to 15
mL 90% propanol, mix, centrifuge for 20 min, and decant supernatant. Repeat using two washes
of 100% propanol and one wash of xylene. Centrifuge, decant xylene, and add fresh xylene. At
this stage, store sample in well-sealed vials or prepare slides.
Slides for periphyton examinations require random dispersion of a known amount of xylene
suspension. Use a microstirrer to break up clumps of algae before removing sample portion from
xylene suspension. Count a number of drops of suspended sample into a thin ring of mounting
medium*#(141) on a slide. Mix the xylene suspension and medium with a spatula until the
xylene has evaporated. Warm the slide on a hot plate at 45°C and cover sample with a cover slip.
Count diatoms on the prepared slides using the magnification most appropriate to the desired
level of taxonomic identification. Count strips or random fields. Calculate diatom density per
unit area of substrate:
where the terms are as defined in Section 10300C.2.
5. Dry and Ash-Free Weight
Collect at least three replicate slides for weight determinations.3 Slides air-dried in the field
can be stored indefinitely if protected from abrasion, moisture, and dust. Use slides expressly
designated for dry and ash-free weight analysis.
a. Equipment:
1) Analytical balance, with a sensitivity of 0.1 mg.
2) Drying oven, double-wall, thermostatically controlled to within ±1°C.
3) Electric muffle furnace with automatic temperature control.
4) Crucibles, porcelain, 30-mL capacity.
5) Single-edge razor blades or rubber policeman.
b. Procedure:
1) Dry slides to constant weight at 105°C, and ignite for 1 h at 500°C. If weights are to be
obtained from field-dried material, re-wet dried material with reagent-grade water and remove
from slides with a razor blade or rubber policeman. Place scrapings from each slide in a separate
prewashed, prefired, tared crucible; dry to constant weight at 105°C; cool in a desiccator and
weigh; and ignite for 1 h at 500°C.
2) Re-wet ash with reagent-grade water and dry to constant weight at 105°C. This
reintroduces water of hydration of clay and other minerals, which is not driven off at 105°C but
is lost during ashing. If not corrected for, this water loss will be recorded as volatile organic
matter.4
c. Calculations: Calculate mean weight from slides and report as dry weight [(crucible +
sample weight at 105° C) minus (tare weight of crucible)] per square meter of exposed surface. If
25-by 75-mm slides are used, then
Calculate ash weight for sample [(crucible + sample weight at 500°C) minus (tare weight of
crucible)]. Subtract ash weight from dry weight to obtain ash-free weight, and report as ash-free
weight per square meter of exposed surface.
6. Chlorophyll and Pheophytin
The chlorophyll content of attached algal communities is a useful index of the
phytoperiphyton biomass. Quantitative chlorophyll determinations require the collection of
periphyton from a known surface area. Extract the pigments with aqueous acetone or methanol
(see Section 10200G) and use a spectrophotometer or fluorometer for analysis. If immediate
pigment extraction is not possible, samples may be stored frozen for as long as 30 d if kept in the
dark.5 The ease with which chlorophylls are removed from cells varies considerably with
different algae; to achieve complete pigment extraction disrupt the cells mechanically with a
grinder, blender, or sonic disintegrator, or freeze them. Grinding is the most rigorous and
effective of these methods.
The Autotrophic Index (AI) is a means of determining the trophic nature of the periphyton
community (see Section 10200G). It is calculated as follows:
Normal AI values range from 50 to 200; larger values indicate heterotrophic associations or
poor water quality. Nonviable organic material affects this index. Depending on the community,
its location and growth habit, and method of sample collection, there may be large amounts of
nonliving organic material that may inflate the numerator and produce disproportionately high
AI values. Nonetheless, the AI is an approximate means of describing changes in periphyton
communities between sampling locations.
a. Equipment and reagents: See Section 10200G.
b. Procedure: In the field, place individual glass microscope slides used as substrates directly
into 100 mL of a mixture of 90% acetone (water with 10% saturated MgCO3 solution).
Immediately store on dry ice in the dark. (NOTE: Vinyl plastic is soluble in acetone. If vinyl
plastic is used as the substrate, scrape periphyton from it before solvent extraction.) If extraction
cannot be carried out immediately, freeze samples in the field and keep frozen until processed.
Rupture cells by grinding in a tissue homogenizer and steep in acetone for 24 h in the dark at
or near 4°C.
To determine pigment concentration, follow the procedures given in Section 10200G.
c. Calculation: After determining pigment concentration in the extract, calculate amount of
pigment per unit surface area of sample as follows:
where:
Ca is as defined in Section 10200G.
7. References
1. WETZEL, R.G. & G.E. LIKENS. 1991. Limnological Analyses, 2nd ed. Springer-Verlag,
New York, N.Y.
2. OWEN, B.B., JR. 1977. The effect of increased temperatures on algal communities of
artificial stream channels. Ph.D. dissertation, Univ. Alberta, Edmonton.
3. NEWCOMBE, C.L. 1950. A quantitative study of attachment materials in Sodon Lake,
Michigan. Ecology 31:204.
4. NELSON, D.J. & D.C. SCOTT. 1962. Role of detritus in the productivity of a rock outcrop
community in a piedmont stream. Limnol. Oceanogr. 7:396.
5. GRZENDA, A.R. & M.L. BREHMER. 1960. A quantitative method for the collection and
measurement of stream periphyton. Limnol. Oceanogr. 5:190.
8. Bibliography
EATON, J.W. & B. MOSS. 1966. The estimation of numbers and pigment content in epipelic algal
populations. Limnol. Oceanogr. 11:584.
MOSS, B. 1968. The chlorophyll a content of some benthic algal communities. Arch. Hydrobiol.
65:51.
CRIPPEN, R.R. & J.L. PERRIER. 1974. The use of neutral red and Evans blue for live-dead
determinations of marine plankton. Stain Technol. 49:97.
OWEN, B.B., M. AFZAL & W.R. CODY. 1978. Staining preparations for phytoplankton and
periphyton. Brit. Phycol. J. 13:155.
OWEN, B.B., M. AFZAL & W.R. CODY. 1979. Distinguishing between live and dead diatoms in
periphyton communities. In R.L. Weitzel, ed. Methods and Measurements of Periphyton
Communities: A Review. STP 690, American Soc. Testing & Materials, Philadelphia, Pa.
WETZEL, R.G., ed. 1983. Periphyton of Freshwater Ecosystems. Developments in Hydrobiology
17. Dr. W. Junk BV Publishers, The Hague, The Netherlands.
DELBECQUE, E.J.P. 1985. Periphyton on Nymphaeids: An evaluation of methods and separation
techniques. Hydrobiologia 124:85.
TREES, C.C., M.C. KENNICUTT & J. M. BROOKS. 1985. Errors associated with the standard
fluorometric determination of chlorophylls and phaeopigments. Mar. Chem. 17:1.
10300 D.
Primary Productivity
The productivity of periphyton communities is a function of water quality, substrate, and
seasonal patterns in temperature and solar illumination. Productivity may be estimated from
temporal changes in biomass (standing crop) or from the rate of oxygen evolution or carbon
uptake.1
1. Biomass Accumulation
a. Ash-free dry weight: The accumulation rate of organic matter on artificial substrates by
attachment, growth, and reproduction of colonizing organisms has been used widely to estimate
the productivity of streams and reservoirs.2,3 To use this method, expose several replicate clean
substrates for a predetermined period, scrape the accumulated material from the slides, and ash
as described previously.
where:
P = net productivity, mg ash-free weight/m2/d,
t = exposure time, d, and
A = area of a slide, m2.
Obtain estimates of seasonal changes in biomass of established communities by placing
many replicate substrates at a sampling point and then retrieving a few at a time at regular
intervals. Replace removed slides with new clean slides. The recommended collection interval
ranges from 2 to 4 weeks for a year or longer.2 Gain in ash-free weight per unit area from one
collection period to the next is a measure of net production.
b. ATP estimates: Measurement of adenosine triphosphate (ATP) has been used in recent
years to estimate microbial biomass in water. This technique is applicable to periphyton.4 It
provides an additional tool for assessing the magnitude and rate of biomass accumulation on
substrates in natural waters. At present, the procedure should be limited to communities
colonizing artificial substrates.
1) Equipment and reagents—See Section 10200I.6a.
2) Procedure—Either scrape periphyton from an exposed artificial substrate or, if standard
glass microscope slides are used, place them in polyethylene slide mailers containing preheated
(99°C) Tris buffer. Immerse in a boiling water bath for 10 min to extract ATP. If samples are not
assayed immediately, freeze at −25°C; they may be stored in a freezer for up to several months.
Complete analysis as directed in Section 10200I.6b. Slides exposed in waters containing high
turbidity may collect substantial amounts of particulates including clays. ATP sorbs to these
materials; the sorption results in a quenching effect.
3) Calculations—See Section 10200I.6c.
2. Standing Water Productivity Measured by Oxygen Method
Hourly and daily rates of oxygen evolution and carbon uptake by periphyton growing in
standing water can be studied by confining this community briefly in bottles, bell jars, or other
chambers. In contrast, the metabolism of organisms in flowing water is highly dependent on
current velocity and cannot be determined with precision under static conditions. Productivity
estimates for flowing waters and those for standing waters present different problems; therefore,
separate procedures are given.
Productivity and respiration of epilithic and epipelic periphyton in littoral regions of lakes
and ponds can be determined by inserting transparent and opaque bell jars or open-ended plastic
chambers into substrata along transects perpendicular to the shoreline.5,6 Chambers are left in
place for one-half the daily photoperiod. The DO concentration in a chamber is determined at the
beginning and end of the exposure period. Gross productivity is the sum of the net gain in DO in
the transparent chamber and the oxygen used in respiration. Values obtained are doubled to
estimate productivity for the entire photoperiod. Alternatively, determine the proportion of the
incubation period of the total insolation during the photoperiod more accurately by measuring
the insolation of the incubation period as a percentage of the total daily insolation. Both these
methods assume that photosynthesis is proportional to irradiance (i.e., not light saturated and no
photoinhibition).
Failure to account for changes in DO in chambers caused by phytoplankton photosynthesis
and respiration may cause serious errors in the estimates of periphyton metabolism. It is essential
that these values be obtained at the time the periphyton is studied by using the light- and
dark-bottle method (see Section 10200I).
1) Clear and darkened glass or plastic*#(142) chambers, approximately 20 cm in diameter
and 30 cm high, with a median lateral port, sealed with a serum bottle stopper for removal of
small water samples for DO analyses or for the insertion of an oxygen probe. Fit the chamber
with a small, manually operated, propeller-shaped stirring paddle.
2) Dissolved oxygen probe, or equipment and reagents required for Winkler dissolved
oxygen determinations: See Section 4500-O.
b. Procedure: At each station place both a transparent and an opaque chamber over the
substrate at sunrise or mid-daylight and leave in place for one-half the daily photoperiod. In
extremely productive environments or to define the hourly primary productivity changes
throughout the day, use incubation periods shorter than one-half the photoperiod. The minimum
incubation period giving reliable results is 2 h. Determine DO concentration at the beginning of
the incubation period.
Include a set of Gaarder-Gran light- and dark-bottle productivity and respiration
measurements with each set of chambers to obtain a correction for phytoplankton metabolism.
Incubate for the same time period as the chambers. See Section 10200I.
At end of exposure period, carefully mix the water in the chambers and determine DO
concentration.
c. Calculations: When the exposure period is one-half of the photoperiod, calculate gross
primary productivity of the periphyton community as:
where:
PG = gross production, mg O2/m2/d12,
Vc = volume of clear chamber, L,
C′fc and C′ic = final and initial concentrations, respectively, of DO in the clear chamber,
mg/L, corrected for phytoplankton metabolism,
Vo = volume of opaque chamber, L,
C′io and C′fo = initial and final concentrations, respectively, of DO in the opaque
chamber, mg/L, corrected for phytoplankton metabolism, and
A = substrate area, m2.
Correct for the effects of phytoplankton metabolism in the overall oxygen change in the clear
chamber by the following equations:
C′fc = Cfc − Cf lb
C′ic = Cic − Cilb
C′fo = Cfo − Cfdb
C′io = Cio − Cidb
where:
Cfc =
Cfl b =
Cic =
Cilb =
Cfo =
Cfdb =
Cio =
Cidb =
final DO concentration in clear chamber, mg/L,
final DO concentration in light bottle, mg/L,
initial DO concentration in clear chamber, mg/L,
initial DO concentration in light bottle, mg/L,
final DO concentration in opaque chamber, mg/L,
final DO concentration in dark bottle, mg/L,
initial DO concentration in opaque chamber, mg/L, and
initial DO concentration in dark bottle, mg/L.
Calculate periphyton community respiration by:
where:
R = community respiration, mg O2/m2/d24h, and
t = length of exposure, h.
Determine the net periphyton comm unity productivity (PN) as the difference:
PN = PG − R
If the incubation time is different from one-half the photoperiod, modify the daily gross
production calculation as follows:
where:
tp = length of the daily photoperiod, h.
Community respiration and net production calculations for incubation periods other than
one-half the photoperiod are not changed.
3. Standing Water Productivity Measured by Carbon-14 Method
The approach is similar to that described above for the oxygen method. Transparent and
opaque chambers are placed over the substrate, carbon-14-labeled Na2CO3 is injected into the
chamber by syringe, mixed well, and allowed to incubate with the periphyton for one-half the
photoperiod. The concentration of dissolved inorganic carbon available for photosynthesis is
determined by titration. At the end of the incubation period, the periphyton is removed from the
substrate and assayed for carbon-14.5
1) Incubation chamber: See Section 10300D.2a.
2) Special equipment and reagents: See Section 10200I.
3) Carbon-14-labeled solution of sodium carbonate, having a known specific activity of
approximately 10 µCi/mL.
4) Other equipment and reagents: See Section 4500-CO2.
b. Procedure: At each station place a transparent and opaque chamber over the substrate and
add approximately 10 µCi carbon-14/L of chamber volume. Mix water in the chambers well,
taking care to avoid disturbing the periphyton. Determine concentration of dissolved inorganic
carbon as described in Section 2320. At end of exposure period, remove surface centimeter of
periphyton and sediment enclosed in the chamber, freeze, and store frozen in a vacuum
desiccator.
Immediately before analysis, expose sample to fumes of HCl for 10 to 15 min to drive off all
inorganic carbon-14 retained in the periphyton. Combust sample (or portion) by the Van Slyke
method6 or oxidize by heating in a closed system. Collect all CO2 for radioassay either by
flushing CO2 into a two-vial train of ethanolamine (2-aminoethanol) or alternative CO2
absorber, such as methoxyethanol (1:7)7 or flushing CO2 produced by combustion into a
gas-flow counter or electrometer. Alternatively, extract known amounts of periphyton biomass
with a tissue solubilizer,†#(143) using, for example, 1.0 mL in closed vials at 60°C for 48 h.8
Radioassay subsamples (100-µL) by liquid scintillation.
c. Calculations:
where:
PN = net primary productivity per unit area of substrate per unit time, mg C/m2/d,
a = 12C available = dissolved inorganic carbon, mg 12C/L = (total alkalinity −
phenolphthalein alkalinity) × 0.2406 = mg 12C/L,
b = 14C assimilated = [(radioactivity of sample in light chamber × k1) − (background
activity of dark chamber × k2)] × (isotope effect, 1.06). Express radioactivity as disintegrations
per second (dps), i.e., counts per second corrected to 100% radioassay counter efficiency.
k1 = correction factor to convert individually different light-chamber volumes to
1 L,
k2 = correction factor to convert individually different dark-chamber volumes to
1 L,
1.06 = isotope effect to correct for slightly greater mass of 14C than of 12C, which
results in a 6% slower assimilation rate,
c = 14C available = 14C activity added = (µCi 14C added) × (disintegrations of
14C/s/µCi) = 3.7 × 104 µCi 14C added, mL,
d = a dimensional factor to convert area of substrate sampled to m2, and
e = factor to expand incubation period to the total daylight period. After integration
by planimetry or electronic digitizer of the total amount of insolation for the day, determine
percentage of total represented by the incubation period.
4. Flowing Water Productivity Measured by Oxygen Method
Primary productivity of the periphyton community in a stream or river ecosystem can be
related to changes in DO. These changes are the integrated effects of photosynthesis, affected by
light levels and turbidity, that occur during the photoperiod by stream phytoplankton,
periphyton, and the submerged portions of macrophytes. Respiration results from metabolism of
plant communities, aquatic animals, and attached and free-floating microbial heterotrophs. Water
depth, turbulence, and water temperature all influence the process of reaeration. Oxygen also can
enter by accrual of groundwater and surface waters. Daily fluctuations in photosynthetic
production of oxygen are imposed on the relatively steady demand of respiratory activity.
However, this latter process may fluctuate greatly in streams receiving a significant load of
organic wastes, particularly under intermittent loads such as oxygen demand from urban
stormwater runoff. Respiration rates also may vary diurnally under certain conditions, but the
factors involved are not well understood.
The rate of change in stream DO (q) in grams per cubic meter per hour is represented by the
following function of the photosynthetic rate (p), respiration (r), reaeration (d), and accrual from
groundwater inflow and surface runoff (a):9
q=p−r+d+a
If the equation is multiplied through by depth in meters (z), the resulting values are in terms
of grams oxygen per square meter per hour. Figure 10300:2 illustrates this conceptual
relationship between q, primary productivity, and respiration of the stream plant community.
The procedure measures the time-variable oxygen concentrations in a stream over a 24-h
period. Compensations are made for oxygen changes due to physical factors (accrual and
reaeration) and the rate of oxygen change due to biological activity that is separated into
components due to respiration and primary production. The metabolic rates are the sum of the
activity of the entire stream community. Planktonic productivity and respiration can be separated
from overall community activity by the use of the light- and dark-bottle oxygen technique (see
Section 10200I). However, in most small streams planktonic production is insignificant. The
component of production and respiration due to macrophytes is very difficult to separate from
periphytic metabolic activity in systems where vascular plants are common.
Because periphyton attach to plant surfaces as well as nonliving substrates, radiotracer
techniques are required to separate the component of production due to macrophytes from that
due to attached algae.10 When vascular plants are present use techniques discussed in Section
10400 to estimate their contribution to net primary productivity.
Respiration by fish and benthic fauna also is difficult to quantitate directly and usually is not
separated from periphyton respiration. If compartmentalized animal metabolism is required,
calculate this contribution from laboratory respiration rates extrapolated to the field situation
based on animal population sizes.11,12
Estimate primary productivity in flowing water by either the free water demand method or
the chamber method.13,14 The first does not introduce artificiality to the system; however, it is
difficult to separate the components of metabolic activity except for the contribution due to
plankton. The chamber method measures periphyton activity alone.15-19
Depending on the hydrologic characteristics of the stream system, accrual and reaeration
may be significant. Accrual can be accounted for by simple mixing equations if estimates of the
accrued flow and its oxygen concentration are known. In practice, select for study reaches that
do not incur significant accrual. Measure reaeration rates either directly15-18 or by estimation
from physical and hydrodynamic features of the stream itself.17,18
a. Equipment:
1) BOD bottles, for light- and dark-bottle measurements. See Section 10200I.
2) DO meter and probe for measurement of DO.
3) Bottom chamber, 60 × 20 × 10 cm, with 32-cm lengthwise dividing baffle,
rheostat-controlled submersible pump, temperature thermistor, and DO probe.14 Use clear and
opaque plastic sleeves for covering chamber and petri dishes or other means of placing
periphyton within chambers.
4) Current meter, capable of detecting water current velocities ranging from 0.03 to 3 m/s in
water depths as shallow as 0.3 m.
5) Tape measure (30 m) and depth staff, or similar equipment, as required to measure stream
cross sections.
6) Fluorometer, capable of detecting fluorescent dye concentration at 0.5 to 100 µg/L
(required only if direct measurement of reaeration is made).
7) Liquid scintillation counter, capable of sensitive detection of 85Kr and 3H (required only
if direct measurement of reaeration is made).
b. Procedure:
1) Light- and dark-chamber method—Grow samples of typical periphyton communities on
artificial substrate or collect natural material. Transfer identical portions to both clear and
opaque chambers, taking care to use sufficient periphyton to make the ratio of chamber volume
to periphyton area equivalent to the ratio of stream volume to periphyton substrate area. Measure
current in the stream and match the circulation rate in the clear and opaque chambers to the
current. Measure DO concentrations initially in both clear and opaque chambers and after 1 to 3
h to estimate the rate of oxygen increase or decrease. Make concurrent measurements of
phytoplankton activity using light- and dark-bottle techniques as described in Section 10200I.2.
Incubate light and dark bottles for the same time interval as the chambers.
Make several measurements during the photoperiod to define daily primary productivity. In
addition, collect sufficient natural substrate samples of the study reach to estimate periphyton
biomass (see Section 10300B). At end of incubation period harvest enclosed periphyton and
determine ash-free biomass (see Section 10300B).
2) Free-water diurnal curve methods—Measure, hourly or continuously, DO concentration
and water temperature for a 24-h period at one or two stations, depending on stream conditions,
precision desired, and availability of equipment. If similar conditions exist for some distance
upstream from the reach being studied, diurnal measurements of DO at a single station are
sufficient to estimate productivity. Where upstream conditions are significantly different from
those in the reach being studied, make measurements at the upstream and downstream limits of
the reach.
If the single-station method is used, measure depth at several points along the study reach to
define average depth. Map and/or make physical surveys to estimate magnitude of possible
sources of accrual via effluents or tributary streams and springs. If the two-station method is
used, measure the wetted cross-sectional stream area as well as current velocity at several points
to define flow (in cubic meters per second) and average cross-sectional area. Correct for
phytoplankton activity by light- and dark-bottle measurements (see Section 10200I.2).
3) Direct measurement of reaeration16—Under special circumstances it may be desirable to
estimate reaeration directly although the results may not be more accurate than those of the
empirical formulations usually used. The tracer gas technique is satisfactory, but is difficult and
requires sophisticated equipment not routinely available. Use this method with care and with full
recognition of its restrictions. Depending on stream flow, release 10 to 250 µCi 85Kr with 5 to
125 µCi 3H at the upstream end of the reach together with sufficient fluorescent dye to produce a
concentration of 10 µg/L when completely mixed across the river cross section. Make
fluorometric measurements at the downstream end of the reach until the dye peak appears, then
collect water samples to measure the 85Kr/3H ratio by liquid scintillation techniques. Record
time of travel for the dye peak from the injection point.
c. Calculations:
1) Chamber method—Calculation is analogous to that used for the bell jar technique
discussed in Section 10300D.2.
where:
Pn = hourly rate of net primary production, mg O2/m2/h,
Vc = volume of clear chamber, L,
B = average periphyton biomass estimated for the study reach, mg/m2,
t = incubation period, h,
W c = total biomass of periphyton contained in clear chamber, mg,
C′fc = final oxygen concentration in clear chamber, corrected for phytoplankton
metabolism, mg/L:
C′fc = Cfc − Cf lb
Cfc = final DO in clear chamber,
Cf lb = final DO in light bottle, and
C′ic = initial oxygen concentration in clear chamber corrected for light-bottle
measurement, mg/L:
C′ic = Cic − Cilb
Cic = initial DO in clear chamber, and
Cilb = initial DO in light bottle.
where:
r = hourly periphyton respiration rate, mg O2/m2/h,
Vo = volume of opaque chamber, L,
B = average periphyton biomass for the study reach, mg/m2,
W o = total biomass of periphyton contained in opaque chamber, mg,
C′io = initial oxygen concentration in opaque chamber, corrected for phytoplankton
respiration, mg/L:
C′io = Cio − Cidb
Cio = initial DO in opaque chamber, mg/L,
Cidb = initial DO in dark bottle, mg/L, and
C′fo = final oxygen concentration in opaque chamber, mg/L:
C′fo = Cfo − Cfdb
Cfo = final DO in opaque chamber, mg/L, and
Cfdb = final DO in dark bottle, mg/L.
For each pair of chamber measurements,
Pg = Pn + r
where:
Pg = hourly gross periphytic primary production, mg O2/m2/h.
PG is the area under the curve of primary production per hour through the photoperiod, mg
O2/m2/d (see Figure 10300:3). Also,
where:
R = total periphyton community respiration, mg O2/m2/d, and
n = number of observations.
Thus,
PN = PG − R
where:
PN = net periphytic production, mg O2/m2/d.
2) Free water methods
a) Calculation of reaeration or diffusion for both the single and upstream-downstream
methods—Calculate k2 from radiotracer data as follows:
and
where:
k2 = reaeration coefficient (base e), d−1,
KKr = base e transfer coefficient for 85Kr, d−1,
t = time of travel, d,
(CKr/CH)u = ratio of released radioactivities (µCi/mL) 85Kr to 3H at the upstream station,
and
(CKr/CH)d = ratio of radioactivities (µCi/mL) 85Kr to 3H at the downstream station.
The reaeration coefficient also can be calculated from an equation relating the rate of energy
dissipation in a stream to k2.16,17
where:
K = escape coefficient,
∆h = change in water surface elevation in a stream reach, and
t = time of flow through a stream reach.
This can be expressed in terms of hydrodynamic and physical data:
where:
K′ = 28.3 × 103 s/m ⋅ d for stream flows between 0.028 and 0.28 m3/s; 21.3 × 103 s/m
⋅ d for stream flows between 0.28 and 0.56 m3/s; and 15.3 × 103 s/m ⋅ d for stream flows above
0.56 m3/s,
k2 = reaeration coefficient, d−1, at 20°C,
20
∆H/∆X = slope, m/km, and
V = velocity, m/s.
Convert k2 to the temperature of the stream by the following equation:
20
where:
k2t = k2 at ambient water temperature, d−1, and
T = ambient water temperature, °C.
Convert to D in mg/L/h:
where:
Cs = oxygen concentration at saturation at ambient stream temperatures, mg/L.
b) Single-station method—Calculation of primary productivity and respiration from diurnal
oxygen and temperature measure ments at a single station is summarized in Figure 10300:4 and
Table 10300:I.
Tabulate hourly DO measurements and temperatures. Determine Cs (DO of air-saturated
H2O at each temperature from Table 4500-O:I) and compute uncorrected DO consumption,
milligrams per liter per hour, for each period:
Plot on the half hour, as shown in Figure 10300:4b.
Calculate the net primary production and respiration of phytoplankton as shown in Section
10200I. Determine the 24-h average hourly plankton respiration,
in milligrams per
liter per hour every half hour. Calculate the hourly net phytoplankton production and tabulate for
the approximate hours during the photoperiod. Plot as shown on Figure 10300:4c.
Calculate and tabulate k2t and substitute D for each Cs, as outlined in ¶ a), above. Plot as
shown in Figure 10300:4c.
Correct each ∆DO for diffusion and phytoplankton metabolism:
∆DOcorrected, mg/L/H = ∆DOuncorrected − D − Pp − Rp
Plot each point as shown in Figure 10300:4d.
Gross primary productivity of the benthic and attached algal populations is computed as the
area under the curve in Figure 10300:4d from sunrise to sunset. This is primary production in
grams per cubic meter per day. Multiply by the average depth for a reach, z meters, to obtain PG
in grams per square meter per day. Calculate community respiration:
R = 24 z F
where:
R = community respiration, g/m2/d,
z = depth, m, and
F = average hourly ∆DO for the dark period (without regard to sign), mg/L/h.
Calculate net primary productivity PN as:
PN = PG − R
c) Upstream-downstream method—Calculation of primary productivity and respiration for a
stream reach from upstream and downstream pairs of diurnal curves of oxygen and water
temperature is summarized in Figure 10300:5 and Table 10300:II. Alternatively, calculate as
below, with oxygen change expressed as the difference between stations rather than as change
per hour. The calculations are analogous. Multiply the area under a curve of oxygen change
between two stations, corrected for diffusion and plankton metabolism and expressed in
milligrams per liter, by the discharge in cubic meters per hour, and divide by the water surface
area between the two stations. This, multiplied by 24, yields gross primary productivity in grams
per square meter per day.
To compute gross primary productivity by this method, tabulate upstream and downstream
DO and average water temperature for the reach at each hour. Calculate ∆DO between upstream
and downstream stations for each hour as
∆DO = DOdownstream − DOupstream
Tabulate Cs and determine the planktonic activity. Correct for planktonic respiration by relating
average hourly dark bottle DO change to the time of travel in the stream reach; correct for
planktonic production by the hourly change in DO in the light bottle times the time of travel (see
Table 10300:II).
Calculate or tabulate k2 and convert it to the total oxygen diffusion for the reach. Because
diffusion, D, is expressed as milligrams per liter per hour, multiply it by the travel time to obtain
the diffusion correction.
Correct each hourly upstream-downstream ∆DO as shown in Table 10300:II. Integrate the
area under this ∆DO curve from sunrise to sunset to give P as in Figure 10300:5d.
where:
Q = flow, m3/h, and
A = reach area, m2 (average reach width × reach length).
and
Net production PN = PG − R
5. References
1. VOLLENWEIDER, R.A., ed. 1969. A Manual on Methods for Measuring Primary
Production in Aquatic Environments. IBP Handbook No. 12. F.A. Davis Co.,
Philadelphia, Pa.
2. SLADEEK, V. & A. SLADECKOVA. 1964. Determination of periphyton production by
means of the glass slide method. Hydrobiologia 23: 125.
3. KING, D.L. & R.C. BALL. 1966. A qualitative and quantitative measure of aufwuchs
production. Trans. Amer. Microsc. Soc. 82:232.
4. CLARK, J.R., D.I. MESSENGER, K.L. DICKSON & J. CAIRNS, JR. 1978. Extraction of ATP
from aufwuchs communities. Limnol. Oceanogr. 23:1055.
5. WETZEL, R.G. 1963. Primary productivity of periphyton. Nature 197: 1026.
6. WETZEL, R.G. 1964. A comparative study of the primary production of higher aquatic
plants, periphyton, and phytoplankton in a large shallow lake. Int. Rev. ges. Hydrobiol.
49:1.
7. LOEB, S.L. 1981. An in situ method for measuring the primary productivity and
standing crop of the epilithic periphyton community in lentic systems. Limnol.
Oceanogr. 26:394.
8. BEER, S., A.J. STEWART & R.G. WETZEL. 1982. Measuring chlorophyll a and 14C-labeled
photosynthate in aquatic angiosperms by use of a tissue solubilizer. Plant Physiol.
69:54.
9. ODUM, H.T. 1956. Primary production in flowing waters. Limnol. Oceanogr. 1:102.
10. ALLEN, H.L. 1971. Primary productivity, chemo-organotrophy, and nutritional
interactions of epiphytic algae and bacteria or macrophytes in the littoral of a lake.
Ecol. Monogr. 41:97.
11. HALL, C.A.S. 1972. Migration and metabolism in a temperate stream ecosystem.
Ecology 53:585.
12. NIXON, S.W. & C.A. OVIATT. 1974. Ecology of a New England salt marsh. Ecol.
Monogr. 43:463.
13. MCINTIRE, C.D., R.L. GARRISON, H.K. PHINNEY & C.E. WARREN. 1964. Primary
production in laboratory streams. Limnol. Oceanogr. 9:92.
14. THOMAS, N.A. & R.L. O’CONNELL. 1966. A method for measuring primary production by
stream benthos. Limnol. Oceanogr. 11:386.
15. COPELAND, B.J. & W.R. DUFFER. 1964. Use of a clear plastic dome to measure gaseous
diffusion rates in natural waters. Limnol. Oceanogr. 9:494.
16. TSIVOGLOU, E.C. & L.A. NEAL. 1976. Tracer measurement of reaeration. III. Predicting
the capacity of inland streams. J. Water Pollut. Control Fed. 48:2669.
17. GRANT, R.S. 1976. Reaeration-coefficient measurements of 10 small streams in
Wisconsin. U.S. Geol. Surv. Water Resources Publ. 76-96.
18. ODUM, H.T. & C.M. HOSKIN. 1958. Comparative studies of the metabolism of marine
water. Publ. Inst. Mar. Sci. Univ. Tex. 4:115.
19. BOTT, T.L., J.T. BROCK, C.E. CUSHING, S.V. GREGORY, D. KING & R.C. PETERSEN. 1978. A
comparison of methods for measuring primary productivity and community respiration
in streams. Hydrobiologia 60:3.
6. Bibliography
POMEROY, L.R. 1959. Algal productivity in salt marshes. Limnol. Oceanogr. 4:386.
CASTENHOLZ, R.W. 1961. An evaluation of a submerged glass method of estimating production
of attached algae. Verh. Int. Ver. Limnol. 14:155.
WHITFORD, L.A. & G.J. SCHUMACHER. 1964. Effect of a current on respiration and mineral
uptake in Spirogyra and Oedogonium. Ecology 45:168.
DUFFER, W.R. & T.C. DORRIS. 1966. Primary productivity in a southern Great Plains stream.
MCINTIRE, C.D. 1966. Some factors affecting respiration of periphyton communities in lotic
environments. Ecology 47:918.
CUSHING, C.E. 1967. Periphyton productivity and radionuclide accumulation in the Columbia
River, Washington, USA. Hydrobiologia 29:125.
HANSMANN, E.W., C.B. LANE & J.D. HALL. 1971. A direct method of measuring benthic primary
production in streams. Limnol. Oceanogr. 16:822.
SCHINDLER, D.W., V.E. FROST & R.V. SCHMIDT. 1973. Production of epilithiphyton in two lakes
of the experimental lakes area, northwestern Ontario. J. Fish. Res. Board Can. 30:1511.
NORTH AMERICAN BENTHOLOGICAL SOCIETY. 1974–1990 (annual). Current and Select
Bibliographics on Benthic Biology. Springfield, Ill.
WETZEL, R.G. & G.E. LIKENS. 1991. Limnological Analyses, 2nd ed. Springer-Verlag, New York,
N.Y.
10300 E.
Interpreting and Reporting Results
Although several systems have been developed to organize and interpret periphyton data, no
single method is universally accepted. The methods may be qualitative or quantitative.
Qualitative methods deal with the taxonomic composition of the communities in zones of
pollution, whereas quantitative methods deal with community structure using diversity indices,
similarity indices, and numerical indices of saprobity.
1. Qualitative Methods (Indicator Species and Communities)
The saprobity system developed by Kolkwitz and Marsson is a widely used method of
interpreting periphyton data. This scheme divides polluted stream reaches into polysaprobic, α
and mesosaprobic, and oligosaprobic zones, and lists the characteristics of each. The system has
been refined1,2 and enlarged by Fjerdingstad3,4 and Sladecek.5,6
Evaluation of the saprobity system requires microscopic evaluation of living indicator biota,
particularly for the sensitive sessile protozoans. Glass slides and other transparent substrates are
advantageous because they permit direct microscopic examination and identification. Removal
of periphyton from slides and preservation for subsequent examination may be acceptable for
diatoms, but observation of preserved material is not acceptable for most flagellated organisms.
2. Quantitative Methods
These methods use cell counts per unit area of substrate and numerical indices of pollution
or water quality. Considerable data on cell densities and species composition of periphyton
collected on glass slides in polluted rivers in England are available.7
Other indices include the Shannon-Weaver,8 Simpson’s,9 and Pinkham-Pearson.10 The
saprobity system11 also may be used where code numbers assigned for the saprobial value and
the abundance of individual species are used to calculate a Mean Saprobial Index. Results also
may be expressed by the truncated-log normal distribution of diatom species12,13 as well as the
Autotrophic Index (AI).14
3. References
1. KOLKWITZ, R. 1950. Oekologie der saprobien. Ver Wasser-, Boden, Lufthyg.
Schriftenreihe (Berlin) 4:1.
2. LIEBMANN, H. 1951. Handbuch der Frischwasser und Abwasserbiologie. Bd. I.
Oldenbourg, Munich, Germany.
3. FJERDINGSTAD, E. 1964. Pollution of streams estimated by benthal
phytomicroorganisms. I. A saprobic system based on communities of organisms and
ecological factors. Int. Rev. ges. Hydrobiol. 49:63.
4. FJERDINGSTAD, E. 1965. Taxonomy and saprobic valency of benthic
phytomicroorganisms. Int. Rev. ges. Hydrobiol. 50:475.
5. SLADECEK, V. 1966. Water quality system. Verh. Int. Ver. Limnol. 16:809.
6. SLADECEK, V. 1973. System of water quality from the biological point of view. Arch.
Hydrobiol. Ergebn. Limnol. 7:1.
7. BUTCHER, R.W. 1946. Studies in the ecology of rivers. VI. The algal growth in certain
highly calcareous streams. J. Ecol. 33:268.
8. SHANNON, C.E. 1948. The Mathematical Theory of Communication. Univ. Illinois
Press, Urbana.
9. SIMPSON, E.H. 1949. Measurement of diversity. Nature 163:688.
10. PINKHAM, C.F.A. & J.G. PEARSON. 1976. Applications of a new coefficient of similarity
to pollution surveys. J. Water Pollut. Control Fed. 48:717.
11. PANTLE, R. & H. BUCK. 1955. Die biologische uberwachung der Gewasser und der
Darstellung der Ergebnisse. Gas-Wasserfach 96:604.
12. PATRICK, R., M.H. HOHN & J.H. WALLACE. 1954. A new method for determining the
pattern of the diatom flora. Bull. Philadelphia Acad. Natur. Sci. 259:1.
13. PATRICK, R. 1973. Use of algae, especially diatoms, in the assessment of water quality.
In J. Cairns, Jr., ed. Biological Methods for the Assessment of Water Quality. ASTM
STP 528, American Soc. Testing & Materials, Philadelphia, Pa.
14. WEBER, C. 1973. Recent developments in the measurement of the response of plankton
and periphyton to changes in their environment. In G. Glass, ed. Bioassay Techniques
and Environmental Chemistry. Ann Arbor Science Publ., Ann Arbor, Mich.
4. Bibliography
FJERDINGSTAD, F. 1950. The microflora of the River Mlleaa, with special reference to the
relation of the benthal algae to pollution. Folia Limnol. Scand. 5:1.
BLUM, J.L. 1956. The ecology of river algae. Bot. Rev. 22:291.
YOUNT, J.L. 1956. Factors that control species numbers in Silver Springs, Florida. Limnol.
Oceanogr. 1:286.
BUTCHER, R.W. 1959. Biological assessment of river pollution. Proc. Linnean Soc. London
170:159.
HOHN, M.H. 1959. The use of diatom populations as a measure of water quality in selected areas
of Galveston and Chocolate Bay, Texas. Publ. Inst. Mar. Sci. Univ. Tex. 5:206.
HOHN, M.H. 1961. Determining the pattern of the diatom flora. J. Water Pollut. Control Fed.
33:48.
PATRICK, R. 1963. The structure of diatom communities under varying ecological conditions.
Ann. N.Y. Acad. Sci. 108:359.
SCHLICHTING, H.E., JR. & R.A. GEARHEART. 1966. Some effects of sewage effluent upon
phyco-periphyton in Lake Murray, Oklahoma. Proc. Okla. Acad. Sci. 46:19.
SLADECKOVA, A. & V. SLADECEK. 1966. Periphyton as indicator of reservoir water quality.
Technol. Water (Czech) 7:507.
TAYLOR, M.P. 1967. Thermal effects on the periphyton community in the Green River. Tennessee
Valley Authority, Div. Health & Safety, Water Qual. Br., Biol. Sect., Chattanooga, Tenn.
PATRICK, R. 1968. The structure of diatom communities in similar ecological conditions. Amer.
Natur. 102:173.
DICKMAN, M. 1969. A quantitative method for assessing the toxic effects of some water soluble
substances, based on changes in periphyton community structure. Water Res. 3:963.
BESCH, W.K., M. RICARD & R. CANTIN. 1970. Use of benthic diatoms as indicators of mining
pollution in the N.W. Miramichi River. Tech. Rep. Fish. Res. Board Can. 202:1.
NUSCH, E.A. 1970. Ecological and systematic studies of the Peritricha (Protozoa, Ciliata) in the
periphyton community of reservoirs and dammed rivers with different degrees of saprobity.
Arch. Hydrobiol. (Suppl.) 37:243.
ROSE, F.L. & C.D. MCINTIRE. 1970. Accumulation of dieldrin by benthic algae in laboratory
streams. Hydrobiologia 35:481.
WHITTON, B.A. 1970. Toxicity of zinc, copper and lead to Chlorophyta from flowing waters.
Arch. Mikrobiol. 72:353.
BURROWS, E.M. 1971. Assessment of pollution effects by the use of algae. Proc. Roy. Soc. Lond.
Ser. B. 177:295.
PATRICK, R. 1971. The effects of increasing light and temperature on the structure of diatom
communities. Limnol. Oceanogr. 16:405.
ARCHIBALD, R.E.M. 1972. Diversity of some South African diatom associations and its relation
to water quality. Water Res. 6:1229.
CAIRNS, J., JR., B.R. LANZA & B.C. PARKER. 1972. Pollution-related structural and functional
changes in aquatic communities with emphasis on freshwater algae and protozoa. Proc.
Acad. Natur. Sci. Philadelphia 124:79.
OLSON, T.A. & T.O. ODLAUG. 1972. Lake Superior Periphyton in Relation to Water Quality.
Water Pollut. Control Res. Ser., 18080 DEM 02/ 72. Univ. Minnesota School Public Health,
Minneapolis.
HANSMANN, E.W. 1973. Effects of logging on periphyton in coastal streams of Oregon. Ecology
54:194.
RUTHVEN, J.A. & J. CAIRNS, JR. 1973. Response of fresh-water protozoan artificial communities
to metals. J. Protozool. 20:127.
MIDWEST BENTHOLOGICAL SOCIETY. 1964–1973 (annual). Current and Select Bibliographies
on Benthic Biology. Springfield, Ill.
NORTH AMERICAN BENTHOLOGICAL SOCIETY. 1974–1991 (annual). Current and Select
Bibliographies on Benthic Biology. Springfield, Ill.
BAXTER, R.M. 1977. Environmental effects of dams and impoundments. Annu. Rev. Ecol.
Systematics 8:255.
WEITZEL, R.L., ed. 1979. Methods of Measurement of Periphyton Communities: A Review.
ASTM Spec. Tech. Publ. 690, American Soc. Testing & Materials, Philadelphia, Pa.
WETZEL, R.G., ed. 1983. Periphyton of Freshwater Ecosystems. Developments in Hydrobiology
17. Dr. W. Junk B.V. Publ., The Hague, The Netherlands.
STEVENSON, R.J. 1984. Epilithic and epipelic diatoms in the Sandusky River USA with emphasis
on species diversity and water pollution. Hydrobiologia 114:161.
KOSINSKI, R.J. 1984. The effect of terrestrial herbicides on the community structure of stream
periphyton. Environ. Pollut. Ser. A, Ecol. Biol. 36:165.
LINDSTROM, E.A. & T.S. TRASAN. 1984. Influence of current velocity on periphyton distribution
and succession in a Norwegian soft water river. Verh. Int. Ver. Limnol. 22:1965.
MCGUIRE, M.J., R.M. JONES, E.G. MEANS, G. IZAGUIRRE & A.E. PRESTON. 1984. Controlling
attached blue-green algae with copper sulfate. J. Amer. Water Works Assoc. 76:60.
PARKER, B.C., G.J. SCHUMACHER & L.A. WHITFORD. 1984. Some rarely reported algae of the
Appalachian Mountains, Eastern North America: Why so rare? Va. J. Sci. 35:197.
10400
MACROPHYTON*#(144)
10400 A.
Introduction
The macrophyton consists principally of aquatic vascular flowering plants, but it also
includes the aquatic mosses, liverworts, ferns, and the larger macroalgae. Like other primary
producers, these plants respond to the quality of the water in which they grow. The use of biota,
including macrophyton, is an increasingly important and recognized technique for assessing
aquatic habitats.1 Macrophyton often constitute a dominant factor in the habitat of other aquatic
organisms.
Freshwater forms range in size from the tiny watermeal (Wolffia spp.), about the size of a
pinhead, to plants such as the cattail (Typha spp.), up to 4 m high, and finally to cypress trees
(Taxodium spp.), up to 50 m high. Higher aquatic plants often are clustered in large numbers,
many in pure stands, covering extensive areas of shallow lakes, reservoirs, marshes, and canals.
A few of the larger freshwater algae (Chara spp. and Nitella spp.) resembling higher plants in
size, form, and habit sometimes are included in the macrophyton. In marine water, the intertidal
rockweeds (Fucus spp. and Ascophyllum spp.) and offshore kelps (Fucus spp. and Macrocystis
spp.) are conspicuous. Vascular marine or estuarine plants, such as the eelgrass (Zostera spp.)
and the marshgrass (Spartina spp.), are essential to the aquatic ecosystem.
Three growth forms of macrophyton generally are recognized: floating, submersed, and
emersed. Floating plants may be rooted or free-floating; their principal foliage or crown floats
freely on the water surface. All or most of the foliage of submersed plants grows beneath the
water surface: nearly all submersed vascular plants have roots. Growing tips of submersed plants
may emerge to flower and some species can produce floating leaves. Emersed plants have their
principal foliage in the air at or above the water surface; they are attached by roots to the bottom
mud. In some cases the same species may grow as a floating or emersed type, or as a submersed
or emersed type. Submersed and emersed vascular plants typically are rooted to the bottom but
they may be found detached and floating.
The distribution and abundance of higher plants is subject to considerable spatial and
temporal variation. Among the many factors that determine their presence, density, and
morphology are sediment type, water turbidity, water current, nutrient concentrations, water
depth, shoreline disturbance, herbivore grazing, and human activities. Zonation in the littoral
zone of lakes and shallow, slow-moving streams is common. Emergent macrophytes generally
are found in the most shallow portion of the littoral zone. During periods of low water level they
may occupy the terrestrial as well as the aquatic habitat. The depth of inhabitation seldom
exceeds 1 m. Floating-leaved plants commonly are found in the shallower littoral areas in water
depths between 1 and 3 m. Submersed plants may occur from the edge of the shore to the
interface of the littoral and profundal zones, but rarely extend beyond a depth of 10 m because of
limitation of underwater light.
2. Reference
1. NORRIS, R.H., B.T. HART & M. FIULAYSAR, eds. 1995. Use of biota to assess water
quality. Austral. J. Ecol. 20:1.
3. Bibliography
BUTCHER, R.W. 1933. Studies on the ecology of rivers. I. On the distribution of macrophytic
vegetation in the rivers of Britain. J. Ecol. 21:58.
WELCH, P.S. 1948. Limnological Methods. Blakiston Co., Philadelphia, Pa.
MILLIGAN, H.F. 1969. Management of aquatic vascular plants and algae. In Eutrophication:
Causes, Consequences, Correctives. National Acad. Science, Washington, D.C.
BOYD, C.E. 1971. The limnological role of aquatic macrophytes and their relationship to
reservoir management. In G.E. Hall, ed. Reservoir Fisheries and Limnology. Spec. Publ.
Amer. Fish. Soc. No. 8.
HUTCHINSON, G.E. 1975. A Treatise on Limnology. Vol. III. Limnological Botany. John Wiley
& Sons, New York, N.Y.
WESTLAKE, D.F. 1975. Macrophytes. In B.A. Whitton, ed. River Ecology. Univ. California
Press, Berkeley & Los Angeles.
WILE, I. 1975. Lake restoration through mechanical harvesting of aquatic vegetation. Verh. Int.
Ver. Limnol. 19:660.
WOOD, R.D. 1975. Hydrobotanical Methods. University Park Press, Baltimore, Md.
RASCHKE, R.L. 1978. Macrophyton. In W.T. Mason, Jr., ed. Methods for the Assessment and
Prediction of Mineral Mining Impacts on Aquatic Communities: A Review and Analysis.
U.S. Dep. Interior, Harpers Ferry, W. Va.
WETZEL, R.G. 1983. Limnology, 2nd ed. Saunders College Publishing, Philadelphia, Pa.
DENNIS, W.M. & B.G. ISOM, eds. 1984. Ecological Assessment of Macrophyton: Collection, Use,
and Meaning of Data. ASTM STP 843, American Soc. Testing & Materials, Philadelphia, Pa.
10400 B.
Preliminary Survey
A macrophyte survey includes species identification, location, assessment of health, and
quantity. More detailed studies may involve functioning of aquatic plants in nutrient and heavy
metal uptake and turnover, use of plants as indicator organisms, and effects of plants on water
quality conditions.
Several sampling protocols are required to meet diverse survey needs. The usefulness of a
given study and the appropriate types of statistical analyses are determined and fixed initially.
To develop a good sample design, determine what information is desired, prevailing
environmental conditions, the life and growth form of the species being sampled, and the
methods for obtaining reproducible data that are comparable to other or future studies. In
defining reporting requirements, consider matters such as the use of scientific names; the
selection of appropriate descriptors such as frequency, density, biomass, cover, diversity,
productivity, and outer limit of vegetation growth; and the use of proper statistical techniques.
2. Pre-Field Investigations
During pre-field investigations assemble maps, charts, aerial photographs, taxonomic keys,
and past reports. Maps, charts, and aerial photographs help determine access routes, project size,
plant community distribution, habitat characteristics that may influence plant distribution, and
sampling obstacles or hazards. These items also provide a base for doing field work and
reporting results. They may be available locally from municipal engineering departments, zoning
boards, planning commissions, drainage districts, and land conservation commissions. At the
state level information may come from natural resource agencies, natural history survey
organizations, universities, and transportation departments. At the federal level, the Geological
Survey, Natural Resources Conservation Services, Bureau of Land Management, Forest Service,
Park Service, Fish and Wildlife Service, Tennessee Valley Authority, Bureau of Indian Affairs,
Army Corps of Engineers, Environmental Protection Agency, National Biological Service, and
the National Oceanographic and Atmospheric Administration have many maps, charts, and aerial
photographs available. A final source is private map companies that publish hydrographic maps
for fishermen and recreational boaters.
Past reports provide historical records useful for planning sampling logistics and interpreting
results. Comparable studies taken at different times provide a dynamic look at the vegetation. An
often-overlooked resource is a herbarium storing pressed and mounted plant specimens. These
generally are located in universities and natural history museums.
3. Field Reconnaissance
Sampling efficiency is improved and a sampling scheme can be refined during a field
reconnaissance. It provides an opportunity to learn the species of plants present and to sketch
their distribution. The species-area curve technique frequently is used to determine the likelihood
of finding more species during a preliminary survey. A field reconnaissance allows the
investigator to answer logistical questions that are the bane of all sampling efforts.
4. Bibliography
ONDOK, J.P. & J. KVET. 1978. Selection of sampling areas in assessment of production. In D.
Dykyjova & J. Kvet, eds. Pond Littoral Ecosystems: Structure and Functioning. Ecological
Studies 28. Springer-Verlag, Berlin.
MACEINA, M.J., J.V. SHIREMAN, K.A. LANGELAND & D.E. CANFIELD, JR. 1984. Prediction of
submersed plant biomass by use of a recording fathometer. J. Aquat. Plant Manage. 22:35.
Also see Section 10400A.2.
10400 C.
Vegetation Mapping Methods
Mapping vegetation stands may be necessary. Do this during the preliminary survey.1
1. Baseline Method
Vegetation maps constructed using the baseline method or the basepoint-stadia rod-alidade
method generally are limited to pure stands of floating or emersed littoral macrophyton in all
bodies of water. In clear water, the outline of pure stands of submersed vegetation can be
determined by using a viewing box (usually a wooden or plastic box with a watertight glass lens)
from the surface or by underwater observation by a diver using a snorkel or SCUBA
(self-contained underwater breathing apparatus). The baseline method and the basepoint-stadia
rod-alidade method provide accurate maps of vegetation in areas up to 1 × 10 5 m2 where most
of the vegetation outline is visible. The baseline method uses intercepting lines from each end of
a predetermined base line to closely spaced markers (i.e., chaining pins) around the stand. By
presetting the map scale, the ratio between the length of the base line and its reduction on the
map (drawn on a plane table) can be determined. The basepoint-stadia rod-alidade method is a
modification of the baseline method in that the distance between the vegetational outline and the
base point is determined with an alidade, range finder, or portable Loran-C unit. One unit on the
stadia rod, as viewed between the cross-hairs of the alidade, is equivalent to a distance of 3.05 m
between the stadia rod and the alidade. Chaining pins are not required when this method is used.
In practice, more readings closely spaced along the vegetational outline usually are taken using
the basepoint-stadia rod-alidade method.
It is not necessary to measure all distances and angles; some can be determined
trigonometrically. After all angles and distances are calculated, fill in irregularities through
inspection and use of other maps and photographs.
The technology for using global positioning systems (GPS), especially when linked to
geographical information systems (GIS), is evolving rapidly.2 These systems probably will have
wide applicability to mapping aquatic vegetation, thereby rendering older surveying techniques
obsolete or at least time-consuming and tedious.
2. Line Intercept Method
The line intercept method is preferable for mapping mixed stands and/or large areas. Select
sampling points at equal intervals along a base line. Choose interval length by the degree of
accuracy desired: the closer the sampling points, the more accurate the map. Run transects
perpendicularly from the base line to the boundary of the plant stand. Use an intercept line
(transect line) of plastic-coated wire rope to prevent stretching. If line flotation is a problem, use
lead weights applied at regular intervals to sink the line and act as interval markers on the rope to
designate sampling units. Use 0.5-m segments (in dense vegetation) to 5-m sampling units (in
sparse vegetation) for determining plant species that vertically intercept the line at each segment.
During underwater surveys, record data with a wax or soft lead pencil on a writing board
constructed of plastic overlays. Construct the vegetation map by placing points where plant
species are found within an outline map (or aerial photograph) of the sample area. Determine the
area that a single species or total vegetation occupies by planimetry or digitization and computer
calculation.
Determine frequency from line intercept or quadrat data, or with a set sampler consisting of a
2-cm steel tube, 2 m long, to which five 0.75-cm by 25-cm steel rods are attached on 40-cm
centers. Record vegetation touching each of the five points within 2.5 cm of the distal tip. If
more than one plant species is touching, record the plant nearest the tip. If no plant is touched,
record bare ground.
3. Remote Sensing
a. General considerations: Remote sensing techniques are used to detect, assess, and
monitor aquatic macrophytes. These techniques include analog aircraft and satellite serial
photography, digital aircraft and satellite multispectral scanning in the visible, infrared, and
thermal bands; microwave techniques, primarily side-looking airborne radar (SLAR); and shuttle
imaging radar (SIR). They provide a synoptic view of large areas, and allow quick surveys to
further delineate areas of interest and repeated viewing at relatively low cost.
In selecting remote sensing system(s) consider expected results, time for project completion,
and available resources. The larger the area, the greater the advantage of remote sensing. Remote
sensing also lends itself to studies over time, because each image is a historical record.
For determining general associations of a widespread macrophytic growth, ground
resolutions of between 30 and 80 m are recommended. Widespread multitemporal coverage is
available at scales ranging from 1:100 000 to 1:1 000 000 at a reasonable cost in the form of
paper prints, transparencies, and digital formats. Landsat Multispectral Scanner (MSS) and
Thematic Mapper (TM) have been used to identify areas with emergent vegetation or topped-out
submergent vegetation. Surface roughness is a requisite for an imaging return with SLAR and
SIR. Landsat provides limited capability for species discrimination, but availability, cost, and
repeat cycles make it useful in determining presence of large populations over time. For a
detailed vegetation survey, including discrete species identification, use much larger-scale
imagery, (1:10 000 or greater). High-altitude aerial coverage available through the National High
Altitude Mapping Program (NHAP) and other sources at original scales of approximately 1:60
000 to 1:120 000 provides both good initial areal coverage and the capacity for enlargement up
to five times.3
After determining scale, select film/filter or sensor combinations. These include black and
white imagery, color infrared, and black and white-infrared imagery; color infrared film used
with a yellow filter is widely applicable, but other combinations also are useful.4-7 For detailed
flight planning consider growth stage of plants, water depth and clarity, tidal conditions, cloud
cover, and sun angle.3,8,9 Available resources ultimately determine remote sensing activities.
b. Aerial photography:
1) Equipment—For all-format aerial photography, use a good-quality 35-mm single-lens
reflex (SLR) camera with manual or through-the-lens metering or any good 70-mm camera
system, preferably with a motor drive. Intervalometers providing for exact exposure intervals for
stereo photography are available for both systems. A 28-mm-focal-length lens with a 35-mm
SLR camera gives wide coverage at low altitudes; 40-mm and 80-mm lenses are successful with
70-mm camera systems. When photographing with black-and-white or color film, use a skylight
or haze filter. When photographing with color infrared film below 1700 m, use a Wratten 12
filter; at altitudes above 1700 m use a Wratten 15 filter. Time of year and condition of vegetation
also may influence the choice of yellow/orange filter. In turbid waters, color film*#(145)
provides better water penetration and is more useful in mapping submersed vegetation than is
color infrared film.4 Color infrared†#(146) yields more detail, is more useful in mapping
emergent vegetation or wetlands, and may provide more detail in clear, nonturbid waters.
Preferably mount camera in the belly of a high-wing, single-engine aircraft for low-altitude,
small-format photography. Belly mounts require special aircraft modification, but provide a
stable platform, protection for the camera, and good access. Alternatively, camera may be
mounted on the aircraft door.
2) Procedures—Because sun angle is critical in obtaining high-quality aerial photography,
schedule flight for a time when solar altitude is between 40 and 68 degrees.7
Set camera at designated ISO reading for the film (assume 100 to 125 for color infrared film)
and shoot in the automatic exposure mode. At a typical airspeed of about 190 km/h
(approximately 120 mph) a shutter speed of 1/250 or 1/500 is adequate. Determine proper f-stop
from an aerial exposure computer; in general, f 5.6 to 11 (f 8 is optimum) gives acceptable color
exposure. Proper exposure of color infrared film depends on such factors as time of day and
year, altitude, humidity, and type of landscape.
Process film through the manufacturer, a photo laboratory, or aerial photography service.
Development of color infrared film is available solely through the manufacturer.
c. Fathometry: Recording fathometers are best applied in water more than 1 m deep where
the instruments can determine accurately the height and distribution of subsurface macrophytes.
A recording fathometer can be mounted on most boats and can accurately determine
one-dimensional (percent cover) and two-dimensional (percent vertical area) profiles of
submersed vegetation. Fathometry is especially useful for determining the outer edge of plant
growth. Make linear and planimetric measurements on chart tracings that provide permanent
records for ready comparison over time. Calculate percent cover by dividing the linear
measurement for a macrophyte species or community by the total chart paper length for any
given transect. Dividing the area of the tracing with macrophytes by the total water area gives
percent vertical area. Use a fathometer accurate to the nearest 0.1 m.
Mount the transducer for the recording fathometer with brackets on the boat’s transom. Keep
speed and recorder speed constant to produce tracings of similar length and resolution. Only a
few minutes are required to replicate transects several kilometers long. Unless gross
morphological differences exist, species discrimination on the chart tracings is difficult or
impossible. Mark boundaries of monotypic colonies and community types with a fixed line on
the chart tracings. Dense vegetation mats that reach the surface may impede boat movement,
prevent the transducer signal from reaching the hydrosoil, and merge tracings of macrophytes
with the transducer line. Tracing patterns from water less than 1 m deep may be difficult to
interpret.10 Location of transects or points along a transect also can be determined using GPS
techniques.2
4. References
1. RASCHKE, R.L. 1984. Mapping—Surface or ground surveys. In W.M. Dennis & B.G.
Isom, eds. Ecological Assessment of Macrophyton: Collection, Use, and Meaning of
Data. Spec. Tech. Publ. 843, American Soc. Testing & Materials, Philadelphia, Pa.
2. KRESS, R. & D. MORGAN. 1995. Application of New Technologies for Aquatic Plant
Management. U.S. Army Waterways Experiment Sta., Corps of Engineers, Aquatic
Plant Control Research Program. Bulletin, Vol. A-95.
3. AVERY, E.T. & G.L. BERLINE. 1985. Interpretation of Aerial Photographs, 4th ed.
Burgess Publishing Co., Minneapolis, Minn.
4. ANDREWS, D.S., D.H. WEBB & A.L. BATES. 1984. The use of aerial remote sensing in
quantifying submersed aquatic macrophytes. In W.M. Dennis & B.G. Isom, eds.
Ecological Assessment of Macrophyton: Collection, Use, and Meaning of Data. Spec.
Tech. Publ. 843, American Soc. Testing & Materials, Philadelphia, Pa.
5. LONG, K.S. 1979. Remote Sensing of Aquatic Plants. Tech. Rep., Waterways
Experiment Sta., U.S. Army Corps of Engineers, Vicksburg, Miss., Vol. A-79–2.
6. BREEDLOVE, B.W. & W.M. DENNIS. 1984. The use of all-format aerial photography in
aquatic macrophyton sampling. In W.M. Dennis & B.G. Isom, eds. Ecological
Assessment of Macrophyton: Collection, Use, and Meaning of Data. Spec. Tech. Publ.
843, American Soc. Testing & Materials, Philadelphia, Pa.
7. BENTON, A.R., JR. 1976. Monitoring aquatic plants in Texas. Tech. Rep. RSC-76, Texas
A & M Remote Sensing Center, College Station, Tex.
8. COLWELL, R.H., ed. 1983. Manual of Remote Sensing, 2nd ed. American Soc.
Photogrammetry & Remote Sensing, Falls Church, Va.
9. SLAMA, C., ed. 1980. Manual of Photogrammetry, 4th ed. American Soc.
Photogrammetry & Remote Sensing, Falls Church, Va.
10. MACEINA, M.J. & J.V. SHIREMAN. 1980. The use of a recording fathometer for
determination of distribution and biomass of hydrilla. J. Aquat. Plant Manage. 18:34.
5. Bibliography
SCHMID, W.D. 1965. Distribution of aquatic vegetation as measured by line intercept with
SCUBA. Ecology 48:816.
OSBORNE, J.A. 1977. Ground Truth Measurements of Hydrilla verticillata Royle and Those
Factors Influencing Underwater Light Penetration to Coincide with Remote Sensing and
Photographic Analysis. Res. Rep. No. 2, Bur. Aquatic Plant Research & Control, Florida
Dep. Natural Resources, Tallahassee.
10400 D.
Population Estimates
1. Sampling Design
The design of a sampling program depends on study aims, collection methods, variation and
distribution of vegetation, personnel and funds available, and accuracy expected. Variation in
space usually is not random; distribution is determined by water depth, shoreline activity,
sediment type, or other factors. The parametric statistic for estimating the true population mean
assumes that the population being sampled has a normal distribution and that all sample units
have the same probability of being selected. Avoid fixed sampling stations in sampling programs
to determine population means, unless they are chosen at random at the beginning of the study.
Because normally distributed plant populations may not be a characteristic of contiguous plant
communities, use parametric statistics with caution.
The simple random sampling design is best applied to homogeneous, noncontiguous plant
communities. The number of stations required to obtain an estimate of the true population mean
with a predetermined level of confidence and permissible error can be determined by applying
the data from a pilot study to the following equation:
where:
N = number of sampling stations,
t = Student’s t at a given probability level; because N is unknown, set t = 2.0; t is
approximately equal to 2.0 for N > 30,
S = standard deviation,
x = estimator of true population mean usually determined by conducting a pilot study;
and
d = permissible error of the final mean; d = 0.1 is recommended for vegetation studies
(± 10%).
An estimate of sampling program cost may be obtained as the sum of initial fixed cost (such
as cost of equipment purchase) and variable cost (cost per sample multiplied by number of
samples).
Apply stratified random sampling to populations having many homogeneous stands. This
design is best applied to populations with obvious gradients and, in practice, to gain precision by
the minimized variance within strata. Determine placement of strata by a pilot study. To
maximize precision, place stratum boundaries around homogeneous areas; generally, the fewer
strata, the greater precision. Allocate sampling in stratified random sampling design according
to:
where:
Ni = number of samples in stratum I,
N = total number of samples,
Wi = a weight reflecting the size (number of quadrats, for example) of stratum i, and
Si = standard deviation of sampled characteristic within stratum I.
Means for population measurement taken along randomly placed stations on a transect line
do not represent large areas of lake populations unless the transect line is placed randomly.
Arbitrarily placed transect lines within a sampling area may or may not reflect the true variation
of the vegetation within.
2. Collection Methods
a. Field inventory/reconnaissance:
1) Manual collection—If water depth, clarity, temperature, flow, and other circumstances
permit, collect specimens by hand. Under ideal conditions, manual collection by wading,
snorkeling, or with SCUBA in deeper water habitats permits a detailed and comprehensive
evaluation of the macrophyte community.
2) Drag chains—Construct drag chains by welding sharpened U-shaped hooks to a short
length (0.6 to 1.0 m) of medium-weight chain. Attach chain to a rope and pull it through the
water. Attach a float to the end of the rope to prevent its loss if the chain is snagged and/or
dropped. The drag chain can be used readily from a slow-moving or stationary boat and is most
efficient in collection of submersed macrophyte species with tall growth forms.
3) Rakes and tongs—Rakes with various handle lengths and oyster tongs may be useful in
collecting macrophytes. A rope may be attached to the rake handle for sampling in deep water or
to facilitate sampling over a wider radius.
4) Grab samplers—Devices developed for sampling benthic organisms, such as the Ekman,
Ponar, and similar grab samplers (see Section 10500B.3), may be used to collect macrophytes.
The light weight of the Ekman grab makes it preferable for the rapid and numerous samplings
often required for survey inventories.
5) Recording fathometers—Use to determine height and distribution of subsurface
macrophytes. Species with similar morphology usually cannot be distinguished from chart
tracing; use supplemental methods to identify species.
b. Quantitative sampling: Numerical data collected to describe vegetation commonly
include such measures of abundance as density, frequency, cover, and biomass/standing crop.1-3
Collect these data from plots or quadrats or, less frequently, by plotless sampling techniques.
The choice of analytical method depends on vegetation density and types, water depth, flow,
height of vegetation in the water column, and nature of the sediment.
1) Line intercepts4—This plotless sampling technique entails use of a weighted nylon or
lead core line laid along the bottom between two known points or oriented by a compass reading.
For dense floating mat vegetation, a floating line may be laid on top of the mat. A surveyor
measures the linear distance occupied by various species that underlie the transect line. Express
these as a percentage of the total line length for individual species as well as for all species
combined. If frequency data are desired, mark the line in increments (e.g., 1 m) and treat species
presence/absence in a manner similar to data from quadrat sampling. The line intercept has been
used to characterize and map aquatic communities5 and to correlate distribution of macrophytes
with selected environmental factors.6 Line transects also are useful for determining patterns of
plant distribution.7 In aquatic environments, the line-intercept method is time-consuming and
may require a diver equipped with SCUBA. Problems arise in determining whether a plant
underlies the transect line.8
2) Belt transects—This technique is similar to the line transect and is useful for biomass or
density determinations. Data are collected along a fixed line, but from a two-dimensional plot or
belt. The belt can be treated as a series of contiguous quadrats or quadrat location may be
selected on the basis of a fixed interval or water depth. Use floating or sinking frames.
3) Quadrats—Quadrats can be used for such population and community estimates as
frequency, cover, density, and biomass. Quadrats can be any two-dimensional shape but are
typically round or rectangular. The sampling area of quadrat samplers can be of any size, but
typically varies from 0.1 to 1 m.2
With the exception of frames, most sampling devices described have been used to obtain
estimates of above-ground biomass (standing crop).9 Above-ground biomass generally is used
because of the difficulty in collecting underground plant parts, such as rhizome and roots.
Without the underground parts, however, the data are of limited value for estimates of primary
production.
a) Manual samplers—These are relatively simple devices for sampling macrophytes, such as
cutting shears. Although they can be used in deep water and manipulated by a diver, they work
best in shallow water. They are relatively inexpensive and can be constructed easily or purchased
from commercial sources.
Frames are suitable for sampling in shallow water. For sampling short, erect plants, use a
square sinking frame constructed of metal. For dense or tangled vegetation, a square assembly
frame with pins or wing nuts at the corners or a fixed-corner three-sided frame may be useful.
Decide whether to include only macrophytes rooted within the frame or also overlapping plants.
In deep water, difficulty and bias may occur in sampling tall submersed vegetation. For
macrophytes forming a dense floating mat, use a floating frame constructed of wood or PVC
pipe.
Box samplers are useful for sampling where water is shallow and the bottom consists of
unconsolidated sediments. The sampler consists of an open-ended box with a metal cutting
flange at the bottom and lateral handles; a sampler constructed of 7-mm plexiglass with
dimensions 0.5 m × 0.5 m × 0.6 m and aluminum cutting flange and corner reinforcements
weighs about 12 kg. With modifications, a box sampler can be used in deep water.10
Benthic dome (BeD) samplers11 may be used for sampling in deep flowing waters. The
sampler consists of a plastic dome with a stainless steel circular collar that can be pushed into the
substrate. It weighs approximately 11 kg and has a sampling area of 0.25 m.2
Various samplers12,13 developed for macroinvertebrate sampling also may be used to collect
macrophytes. These include the Surber sampler, suitable for shallow rivers with moderate
current [see Section 10500B.3b1)]; the stovepipe (cylindrical) sampler, suitable for wadable
waters with unconsolidated sediment bottoms [see Section 10500B.3c4)]; and the Ekman grab
sampler, best suited for soft sediment bottoms with short, erect vegetation [see Section
10500B.3a6)].
b) Mechanically operated samplers—Mechanical sampling devices are costly and complex,
and require a floating platform with winches, cables, and booms. The samplers described below
are useful in deep water. They may decrease sample collection time, increase accuracy of
above-ground and total biomass estimates, and be subject to less bias than many manual
methods.
CAUTION: Use extreme caution for safe operation.
The Louisiana box sampler is an open-ended 35-cm-high box made of sheet metal or similar
material that samples a 61- × 61-cm quadrat (sampling area = 0.37 m2).14 It can be used from a
V-hulled or pontoon boat and is hoisted above the water with a cable and boom. A quick-release
mechanism lets the sampler fall free through the water column. Aquatic vegetation is trapped
against the bottom and severed by cutting edges along the base of sampler. A nylon net sack over
the top retains severed plant fragments. A diver inserts a horizontal cutting blade in a slot at the
level of the substrate before the sampler is hoisted to the surface. Manual insertion of the cutting
plate by a diver makes use of the Louisiana box sampler comparatively efficient. In soft
sediments, the sampler may penetrate too deeply and require lifting before the cutting plate can
be inserted. Rocks, stumps, roots, and other debris may prevent complete closure of the cutting
door.
The Osborne sampler15 is a stainless steel box having outside dimensions of 50 cm × 50 cm
× 60 cm high and a sampling area of 0.25 m.2 The sampler weighs 110 kg and is operated by
winch and cable from a pontoon boat. After hoisting and suspending the sampler alongside the
pontoon boat, a quick-release mechanism allows free-fall through the water column. Tempered
steel blades along the bottom edge of the sampler cut vegetation during the descent. A wire mesh
screen fastened to the top prevents loss of plant fragments. A hinged slotted door is closed with a
lift cable and the sampler is winched to the pontoon boat platform for removal of macrophytes
and sediments. Because the sampler penetrates and collects sediments, the sample includes roots
and rhizomes and can be used to estimate total biomass as well as above-ground biomass.
Efficient operation and accurate biomass estimates require an unconsolidated substrate free of
rocks and other debris.
The Waterways Experiment Station (WES)16 sampler is made of perforated stainless steel
and operated from a pontoon boat with an overhead beam that allows it to be hydraulically raised
and lowered through a circular opening in the pontoon’s platform. Two types are available: one
is cylindrical with a sampling area of 0.28 m2 and the other is square with sampling area of 0.39
m.2 Rotating cutting blades at the base of each sampler sever vegetation as the samplers are
lowered. The bottom cutting plate of each is closed hydraulically. A major advantage of the
WES sampler is its capability to obtain plant samples from any depth. The Louisiana Box and
Osborne samplers, once released, free-fall to the substrate, whereas the hydraulic operation of
the WES sampler controls its descent. The size and weight of the trailer and pontoon boat for the
WES sampler restrict its use in certain water bodies and require an improved ramp for launching.
Although the WES square sampling head is reported to provide a more accurate estimate of
above-ground biomass than the circular one, a substantial underestimate of actual above-ground
biomass still is reported.16
3. Sample Preparation and Analysis
a. Biomass:
1) Fresh weight (wet weight)—Wash samples free of silt and debris, place in a nylon bag
(mesh size 0.75 cm) and spin in a garment washer at 560 rpm for 6 to 7 min to remove excess
moisture. Weigh sample to nearest 0.1 g.
2) Dry weight—Dry subsample (not less than 10%) in a forced-air oven at 105°C for 48 h or
until a constant weight is achieved. The coefficient of variation for a series of subsamples should
not exceed 10%. Calculate dry weight by dividing dry weight of subsample by fresh weight of
subsample times fresh weight of sample.
3) Ash-free dry weight—Transfer dried subsample to a covered and preweighed crucible.
Ignite at 550°C for 6 h. The amount of ash is the weight of material remaining after combustion.
Calculate ash-free dry weight by determining the ratio between ash and dry weight times dry
weight of sample (see Section 10300C.5).
b. Chlorophyll content: Extract fresh plant material with acetone made basic with MgCO3.
Grind the plant material and centrifuge at 2500 rpm for 10 to 15 min. Wash residue with acetone
and add filtered washings to extract. Dry overnight in a container with anhydrous Na2SO4.
Dilute with 90% acetone and water. Determine chlorophyll content (see Section 10200G).
c. Carbon content: Most plants (entire) contain 46 to 48% carbon on a dry-weight basis. A
factor (46.5%) can be used to calculate carbon content and make comparisons.
d. Caloric content: Determine energy content by bomb calorimetry.
e. Species identification:
1) Sample preparation—Use fresh specimens for identification wherever possible. Avoid
immature plants or plants lacking flowers. Because aquatic plants contain from 80 to 95% water
and have less supportive tissue than terrestrial forms, a different procedure is required for drying,
preserving, and mounting them. Collect plants during peak growth when flowers and/or fruits are
present, if practical. Collect the entire plant (stems, rhizomes, leaves, roots, flowers, and fruits).
After collection, either press plants in the field17-19 or wrap specimen in several layers of
paper and submerge in water. Label wrapped specimens with date and location of collection on
an index card and place sample and card in a plastic bag. Preferably use an ice chest containing
crushed ice for storage in the field. Press plants as soon as practical. They can be kept for several
days under refrigeration at 4°C.
Clean plant of all silt and residue. Prepare a mount by centering the plant on 100% rag
herbarium paper. Place emergent plants immediately on paper because they take on a natural
posture. Place a limp plant in a shallow pan of water and slide the herbarium paper under it; with
a slow motion, raise the paper at a 30° angle while keeping the plant centered. Leaves and stems
should lie flat on the paper. Drain off excess water, cover with wax paper to prevent plant from
sticking to blotters, and place in a plant press between paper and blotters. Place plant press in a
dryer. Dry plants at room temperature, but change blotters at least every other day until the plant
is sufficiently dry for permanent mounting.
To prepare a wet mount place specimen in an airtight glass vessel filled with 1 part 10%
formalin, 3 parts water, and a trace of powdered copper sulfate. Plants will remain lifelike and
retain their color for many years in this condition.
2) Identification—A stereomicroscope is needed to identify many plants, especially aquatic
grasses and sedges. Observe vegetative and floral structures by dissecting them, under
magnification, with forceps and fine needle probes.
Preferably identify to species. Numerous references are available to assist in identifying
aquatic macrophytes (see Section 10900E.4).
3) Plant label—An important part of the species collection is the label that identifies the
plant, the collector, the location of the collection, and the date of the collection.19 Attach label to
the sheet with the mounted plant. The mounted plant is a permanent record that is most useful
when placed in an herbarium where it can be utilized by others.
4. Data Presentation
Express fresh weight (wet weight), dry weight, and ash-free dry weight as grams or
kilograms per square meter. Data are best expressed as ash-free dry weight of total biomass.
Determine significant digits for dry weight and ash-free dry weight from the accuracy of the
scale used to obtain fresh weight: do not use more significant digits than those used for
expressing fresh weight. Report pigment as grams chlorophyll per gram dry plant matter and
caloric value as gram calories per gram dry plant matte

Printing - Standard Methods for the Examination of Water and

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