Entire Stream Team Water Quality Monitoring Manual

Transcription

Volunteer
Stream
Monitoring
Training
Manual
Miami Conservancy District
May 2005
Volunteer
Stream
Monitoring
Training
Manual
Third Edition
Published by The Miami Conservancy District 2004
Miami Valley Stream Team is sponsored by:
Miami Conservancy District
38 E. Monument Ave.
Dayton, OH 45402
(937) 223-1271
www.miamiconservancy.org
Montgomery SWCD
10025 Amity Road
Brookville, OH 45309
(937) 854-7645
www.montgomeryswcd.org
Five Rivers MetroParks
1375 E. Siebenthaler Ave.
Dayton, Ohio 45414
937-275-PARK
www.metroparks.org
This manual was written by Sarah Hippensteel and adapted from the second edition of the Hoosier Riverwatch Volunteer
Stream Monitoring Training Manual.
The second edition of the Hoosier Riverwatch Manual was written by Lyn Hartman and Mandy Burk in 2000. The first
edition was written by Sarah Hippensteel in 1997. Hoosier Riverwatch is a program of the Indiana Department of Natural Resources - Division of Soil Conservation.
The large-scale graphics were created by Sarah Beth Lauterbach unless otherwise indicated.
For more information on Miami Valley Stream Team,
the Great Miami River Watershed,
or The Miami Conservancy District, please visit our website:
www.miamiconservancy.org
This manual is printed on recycled paper.
Table of Contents
Welcome to Miami Valley Stream Team
How do Volunteers Get Started?
Chapter 1- Introduction
Water Quality Monitoring
Safety!
Chemical Safety
Chapter 2 - Designing your Water Study
Identifying Your Watershed
What is Water Pollution and Where Does it Come From?
What is Your Watershed Address?
Great Miami River Watershed
Setting Goals
Planning a Water Study
Quality Assurance and Quality Control
Chapter 3 - Habitat Study
What is the riparian zone?
Citizens Qualitative Habitat Evaluation Index (CQHEI)
Site Map
Chapter 4 - Chemical Monitoring
Eight Chemical Tests
Chemical Parameters and Test Procedures
DISSOLVED OXYGEN
E. coli
pH
BIOCHEMICAL OXYGEN DEMAND (5-Day)
WATER TEMPERATURE CHANGE (1 mile)
TOTAL PHOSPHATE (PO4) and Orthophosphate
NITRATE (0-1, 0-10 mg/L)
Chemical Monitoring Worksheet & Data Sheet
Chemical Monitoring Data Sheet Instructions
Chapter 5 - Biological Monitoring
Benthic Macroinvertebrates
Taxonomic Key to Benthic Macroinvertebrates
How to Complete the Biological Monitoring Data Sheet
Chapter 6 - Data - What’s Next?
Data Analysis, Action & Evaluation
Guide for Water Quality Ranges
Pollution Indicators Table
Habitat Parameters for Selected Macroinvertebrates
Volunteer & Organization Registration
Stream Site Registration
Record-keeping Form
Appendix A - Equipment
Appendix B - Glossary
Appendix C - Suggested Reading
Appendix D - Watershed Group Contacts
Appendix E - References
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Chapter 1- Introduction
Welcome to Miami Valley Stream Team
The Miami Valley Stream Team is a volunteer water quality monitoring program. It was
started in the Great Miami River Watershed
by The Miami Conservancy District and Five
Rivers MetroParks to increase public awareness
of water quality issues and concerns by training volunteers to monitor stream water quality.
Miami Valley Stream collaborates with agencies
and volunteers to:
Increase public involvement in water quality
issues through hands-on training of volunteers in
stream monitoring and cleanup activities.
Provide water quality information to citizens
and organizations working to protect the Great
Miami Valley's rivers and streams.
The Miami Valley Stream Team will assist you
and your organization in understanding the importance of protecting local streams. Voluntary
participation is the key to the success of the watershed-wide stream monitoring and education
program. This manual provides information to
help you begin a successful water quality monitoring program.
Educate local communities about the relationship between land use and water quality.
How do Volunteers Get Started?
To become a trained volunteer - it is recommended that you thoroughly read this manual
and attend a Miami Valley Stream Team training
workshop. Workshops are free and open to the
public, and provide hands-on monitoring experience. Contact The Miami Conservancy District
at (937) 223-1271 for a current training schedule.
Training introduces citizens and educators
to water quality monitoring utilizing habitat,
chemical, and biological assessment methods.
The training focuses on the use of Hach chemical testing kits, seine nets, and quality assurance
and control techniques for good data collection.
The sessions are about 6 hours in length and take
place both inside and out of doors. Volunteers
are then able to perform stream testing and teach
others how to monitor.
Volunteer Stream Monitoring Training Manual
1-1
Water Quality Monitoring
What is trend monitoring?
Trend monitoring is the primary testing method
used by Miami Valley Stream Team. To get an
accurate picture of a stream's water quality, tests
have to be performed on a regular basis over a
period of years. Trend monitoring provides a
broad view of the stream allowing the seasonal
variations to be sorted out from long-term changes. In order to get useful data for trend analysis,
a group should consider the long-term commitment involved in this type of monitoring.
What parameters are used?
Water quality is determined by a variety of factors (See figure below). But due to time and
resource constraints, Miami Valley Stream Team
volunteers only monitor a fraction of the possible
parameters. These parameters are listed below:
Habitat - land use, substrate, flow, depth, riparian vegetation, stream shape, erosion
Chemical - dissolved oxygen, nitrate nitrogen,
total phosphate, orthophosphate, turbidity, pH,
biochemical oxygen demand (BOD), temperature change
Biological - benthic macroinvertebrates and fish
identification
1-2
Safety!
Safety is the critical first step in any volunteer
stream monitoring program. All volunteers
should read the following safety precautions
prior to beginning any monitoring activities.
Take a buddy along! Always monitor with
at least one partner. Always let someone else
know where you are, when you intend to return,
and what to do if you do not return on time.
Honor private property rights. Never cross
a landowner’s property without permission.
Never wade in swift or high water. Do
not wade if depth is greater than knee-deep. Do
not monitor if the stream is at flood stage. Any
stream is dangerous in times of flooding.
Never drink the water in a stream. Bring
water from home and be very wary when eating
and drinking if your hands have been in contact
with stream water.
Beware of polluted streams that are known
to be unsafe for handling. Check with your
County Health Department or the Ohio Environmental Protection Agency for information
on bacterial and/or toxic contamination of local
waterways. As a rule, treat every stream as if it
were polluted - wear waders, rubber gloves, and
protective eyewear.
Have a first aid kit on hand. At least one
team member should have first aid/CPR training.
Listen to weather reports. Never monitor if
severe weather is predicted or if a storm occurs.
Be very careful when walking in the
stream. Wear shoes that are in good condition
and have traction. Rocky-bottom streams can
be very slippery and may contain deep pools.
Muddy-bottom streams may also prove dangerous where mud, silt, and sand have accumulated
in sinkholes. If you must cross the stream, use
a walking stick to steady yourself. Watch for
barbed wire fences or sharp, rusty objects (e.g.,
car bodies, appliances) that may pose a particular
hazard.
Do not walk on unstable stream banks.
Disturbing these banks-including the vegetation
growing upon them-can accelerate erosion and
lead to a collapse.
Beware of animals. Watch for irate dogs,
farm animals, wildlife (e.g., snakes), and insects
such as ticks, mosquitoes, and hornets. Know
what to do if you are bitten or stung.
Beware of plants. Watch for poison ivy,
poison oak, sumac, and other skin-irritating
vegetation.
If you drive, park in a safe location. Be
sure your car doesn’t pose a hazard to other drivers and that you are not trespassing. If you are
sampling from a bridge, take special precautions.
Watch out for passing traffic and never lean over
the bridge unless you are firmly anchored.
Develop a safety plan. Find out the location
and telephone number of the nearest telephone
and write it down. Locate the nearest medical
center and write down directions for traveling
there. Have each volunteer monitor complete a
medical form that includes emergency contacts,
insurance information, and pertinent health information such as allergies, diabetes, epilepsy, etc.
1-3
Chemical Safety
The chemical reagents supplied in the testing
kits are laboratory grade reagents. Some of the
chemicals are concentrated, some are irritating,
some are poisonous and some will just make you
itch. Please read thoroughly the directions and
the Materials Safety Data Sheets provided with
each chemical test kit.
Wear safety goggles and rubber gloves.
Avoid contact between chemical reagents and
your skin, eyes, nose, and mouth. Never use
your fingers to stopper a bottle when shaking a
solution.
Do not mix chemicals indiscriminately.
Use only the designated chemicals in specified
amounts when performing tests.
Provide wash water at the monitoring site to
wash any chemicals from the eyes or the body.
Know chemical clean-up, disposal, and
first aid procedures. Wipe up all spills when
they occur. Use sealed plastic containers filled
with an absorbent material (e.g., kitty litter) to
store waste before disposal. Separate hazardous
nitrate waste (Hach kit) from all other waste. If
accidental consumption of chemical reagents
occurs, contact your local poison control office
listed below.
A first aid kit may not be enough. In addition, carry such safety equipment as life buoys,
life jackets, river rescue throw bag, a flashlight, a
whistle,a cell phone, and insect repellant.
Phone numbers for EMERGENCY only:
Ohio Poison Control Center
HACH Company
Rocky Mountain Poison Center
800-222-1222
800-227-4224
800-623-5716
First Aid Kit
Your first aid kit should contain the following items (at a minimum):
Telephone numbers of emergency personnel
Several Band-Aids for minor cuts
Antibacterial soap or alcohol wipes
First aid cream or ointment
Several gauze pads 3-4" square for deep wounds with excessive bleeding
Aspirin or other pain reliever/fever reducer
A needle and tweezers for removing splinters
A first aid manual that outlines diagnosis and treatment procedures
A single-edged razor blade for minor surgery and cutting tape to size
A 2" roll of gauze and a triangular bandage for large wounds
A large compress bandage to hold a dressing in place
A 3" wide elastic band for sprains, applying pressure to bleeding wounds
If a participant is sensitive to bee stings, include their doctor-prescribed antihistamine
An eyewash to flush chemicals
1-4
Chapter 2 - Designing your Water Study
Identifying Your Watershed
The first step in developing a water study design
is identifying your watershed. The watershed is
the total area of land that drains into a particular
waterbody (wetland, stream, river, lake, or sea).
Land uses and runoff in a watershed determine
the quality of surface water in smaller streams
and waterways and can then influence the water
quality of larger streams.
The ability of a stream to support beneficial
uses such as fishing, boating and swimming
is influenced by the major land uses in the
watershed, the nature of the stream channel, the
diversity of instream habitats, and the character
of the riparian area.
Approximately one percent of a watershed is
stream channels. The smallest channels in a
watershed have no tributaries and are called
first-order streams. When two first-order streams
join, they form a second-order stream. When
two second-order streams join, a third order
stream is formed, and so on. First and secondorder channels are often small, steep or intermittent. Stream orders that are six or greater constitute large rivers.
The stream channel is formed by runoff from
the watershed as it flows across the surface of
the ground following the path of least resistance.
The shape of the channel and velocity of flow are
determined by the terrain, unless changes have
been made by man. When the terrain is steep,
the swiftly moving water may cut a deep stream
channel and keep the streambed free of sediments. In flatter areas, the stream may be shallow and meandering, with a substrate comprised
largely of fine sediments.
2-1
What is Water Pollution and Where Does it Come
One reason many volunteers choose to monitor
is concern about pollution impairiming water
bodies. Volunteers can monitor for current pollution and develop a baseline to gauge future
pollution.
Water pollution can typically be placed in one
of two categories, point or nonpoint source
pollution. Point source pollution is easy to
identify because it is discharged from the end
of a pipe. It accounts for about 25% of all
water pollution. Point sources are regulated by
permits issued from the Ohio Environmental
Protection Agency (for more information http:
//www.epa.state.oh.us/).
Nonpoint source pollution originates primarily
from runoff and is more difficult to identify. It
can be defined as polluted run-off from land and
makes up about 75% of water pollution. Different types of pollution are described below and
shown in the figure below.
2-2
Point sources are indicated by a "P"; nonpoint
sources are "NP."
1. Organic Pollution - decomposition of onceliving plant and animal materials
2. Inorganic Pollution - suspended and dissolved solids (e.g. silt, salt, minerals)
3. Toxic Pollution - heavy metals and lethal
organic compounds (e.g. iron, mercury, lead,
PCB's) - some of these are transferred via the
atmosphere and air deposition
4. Thermal Pollution - heated water from
runoff (e.g. streets, parking lots) or point source
discharges (e.g. industries, nuclear or other power plant discharges)
5. Biological Pollution - introduction of nonnative species (e.g. zebra mussels, purple loosestrife, Eurasian Water Milfoil).
What is Your Watershed Address?
The Miami Valley Stream Team organizes data
from volunteer stream monitors by watershed location using the 11- Digit Hydrologic Unit Code
Areas delineated by the U.S. Geological Survey.
Hydrologic unit codes (HUCs) represent the
geographic boundaries of water as it flows across
the landscape. The Ohio Environmental Protection Agency also uses this watershed scale to
organize official data from Ohio's surface water
quality studies.
As you can see from the map on the following
page each watershed name also has an associated 11-digit number. This number is representative of the size of the watershed. The two largest
watersheds - the Upper Great Miami and the
Lower Great Miami are represented by 8-digit
numbers.
Knowing your "watershed address" is important
to understanding the influences on the water
quality in your stream or river.
Both your watershed name and 11-digit # are
required on all Stream Team data sheets.
Check the map on the following page and write your watershed address here:
Watershed Name: ____________________________________________________
Watershed # __ __ __ __ __ __ __ __
2-3
2-4
Great Miami River Watershed
The Great Miami River Watershed is located in
the southwest portion of Ohio. Major tributaries
of the Great Miami River include the Stillwater
River (676 mi2) and the Mad River (657 mi2),
both of which join the Great Miami River at
Dayton, Ohio. The total drainage area of the
Great Miami River watershed in Ohio is 4,124
square miles. The Great Miami River Watershed
includes all or part of 15 counties with its headwaters in Hardin and Auglaize counties and the
mouth in Hamilton County where it drains to the
Ohio River.
Dayton, with a population of 190,000, is the
largest city within the watershed. Other major
cities within the watershed exceeding 50,000
population include Springfield, Hamilton, and
Middletown. Cities with more than 20,000
people include Piqua, Troy, and Fairfield. Each
of these major population centers is located directly adjacent to one of the streams or rivers in
the watershed.
Approximately 79 percent of the total land area
is used for agricultural activities, primarily rowcrop production of corn, soybeans, and alfalfa.
Residential, commercial, and industrial land
uses comprise 13 percent of the area whereas
the remaining area consists of forests (7 percent) and water bodies or wetlands (1 percent).
Major industries, which are concentrated along
the Dayton- Cincinnati corridor, produce automobile parts, business and computer equipment,
chemicals, household goods, paper products, and
processed foods and beverages.
Surface Water
Water quality in the rivers and streams has
shown strong improvement since the passing of
the Clean Water Act in 1972. The following table
shows the number of stream miles that meet
water quality standards of the stream miles that
have been monitored by the Ohio EPA:
Fully attain standards 427.3
Partially attain standards
309.5
Non-Attaining of standards 285.3
Threatened
40.3
Total Miles Monitored
1063.0
The improved quality of the surface waters in
addition to the existence of several major lakes
provides many opportunities for water-based
recreation. Boating, swimming, and fishing are
a few of the many activities enjoyed on Acton
Lake, Indian Lake and Lake Loramie. The cold
water habitat of the Mad River is one of the few
trout fishing streams in Ohio.
Groundwater
The Miami Valley is fortunate to have one of the
largest and most productive aquifer systems in
the country. The Great Miami buried valley aquifer consists of ancient river valleys filled with
permeable deposits of sand and gravel capable of
storing vast amounts of groundwater. The buried
valley aquifer has sustainable yields of 500 to
3,000 gallons per minute. This aquifer system
was designated by US EPA as a Sole Source
Aquifer in 1988. An estimated 97% of the population in the watershed relies on groundwater for
their drinking water supply.
Geology and Soils
There are 2,360 miles of rivers and streams in
the Great Miami River Watershed. Major tributaries include the Great Miami River, Mad River,
Loramie Creek, Twin Creek, and the Stillwater
River, which is designated as a State Scenic
River.
The geology of the watershed consists of bedrock underlying unconsolidated, surficial sediments containing Ordovician-age interbedded
limestone and shale and Silurian-age shale,
limestone and dolomite. The dominant soils in
this watershed are Miamian, Crosby, Russell,
Kokomo, Blount, Pewamo and Glynwood.
2-5
2-6
Setting Goals
The next step in developing a water study design is creating a list of goals. These will differ for each group depending upon individual
interests. While developed for use in schools,
the following types of goals apply to all Miami
Valley Stream Team volunteers (modified from
the GREEN Standard Water Monitoring Kit
Manual):
Data Collection or Scientific Goals
Volunteers will:
- plan, implement and analyze a scientific investigation;
- develop field skills necessary for water quality
testing;
- strengthen observational, analytical and problem-solving skills;
- compile and compare water quality data;
- use and integrate several disciplines (chemistry,
biology, geography, math, etc.).
Community Goals
Volunteers will:
- become actively involved in a community-supported water quality monitoring program;
- develop an awareness and responsibility to
their watershed as an individual and as a community;
- communicate findings and the results of their
actions to the community.
Environmental Eductional Goals
Volunteers will:
- become familiar with the river ecosystem;
- learn to recognize water quality problems and
their sources;
- understand relationships between land use and
water quality;
- make a responsible, action-oriented contribution toward protecting the river and watershed.
Planning a Water Study
The final step in developing a water study design is actually planning the stream study. This
involves choosing your sampling site(s) and setting a sampling schedule. Each sampling site
is a 200-foot stream segment. You should use
local landmarks (bridges, trees) or survey tape to
define the boundaries of your sampling site. You
must also ensure safety by considering accessibility, water depth, and private property rights.
See the Chapter 1 for these and other important
safety considerations.
Site Selection
Your sampling sites should reflect your individual goals and interests. If you are interested
in the affects of agriculture on water quality, you
may want to sample a stream with a primarily
agricultural watershed. If you want to determine
the affects of industrial discharge on stream
water quality, you may choose to monitor at two
points, one upstream of and one downstream
from the industry in question. It is up to you to
choose where you would like to monitor. If you
need help choosing a spot, your local watershed
group or county Soil and Water Conservation
District may have some suggestions (see Appendix B for a list of contacts).
Sampling Schedule
Finally, make a sampling schedule. Consider
how many people will be monitoring, how many
sites you, or your group plan to sample and
whether sampling is feasible year-round (due to
drought, ice cover, etc.). Think about the types
of tests that you will perform, the time requirements, and the goals you have set. A schedule
will help you organize yourself and/or your
group and make these goals more attainable.
2-7
Quality Assurance and Quality Control
Many volunteers strive to obtain the best data
possible. We think this is important, as YOU are
one of the primary users of the data. The following are some suggestions on how you can improve the quality of your water monitoring data.
A quality assurance and quality control (QA/QC)
plan can help ensure that test results are as accurate and precise as possible. Accuracy refers
to how close a measurement is to the true value.
Precision means the ability to obtain consistent
results. Reliability in both accuracy and precision is achieved by:
- Collecting the water sample as directed
- Rinsing bottles/tubes with sample water before
collecting the sample and with distilled water
after completing the test
- Performing tests immediately after collecting
the water sample
- Careful calibration, use, and maintenance of
testing equipment (check by using blanks and
standards)
- Running splits to test for operator error
- Following the specific directions of a testing
protocol exactly as described
- Repeating measurements to check for accuracy
and to understand any sources of error
- Minimizing contamination of stock chemicals
and testing equipment
- Storing kits away from heat and sunlight
- Checking expiration dates on chemicals and
replacing them before they expire
2-8
- Checking to be sure the numbers submitted to
the Stream Team database are the same as those
recorded on the data sheets.
Standards, Blanks and Splits
A standard is a sample of known concentration.
Standards can be purchased from Hach or other
chemical companies. A blank is a sample run
using distilled water. By testing standards and
blanks, volunteers can check for bad reagents
and equipment contamination. A split is one
sample tested twice (for example, two nitrate
tests preformed out of the same bucket of water
taken from a stream). Splits test for operator error, as both tests should yield the same result.
Calibration
Calibration is a procedure to check the accuracy
of testing equipment and to “reset” the equipment to a correct, known value. For example,
to ensure that a pH pen is functioning properly,
a solution of known value (a standard) is tested.
Some calibration procedures may be done in
the classroom or at home just before taking the
equipment into the field. However, in some
cases, it may be necessary to check the calibration again in the field. For most Stream Team
volunteers, the pH pen is the only equipment requiring regular calibration. However, if you use
other high-tech electrical equipment, calibration
is likely required.
Repeated Measurements
By repeating measurements, volunteers collect
better data for two reasons. First, streams and
rivers are variable. The water that flows past
a point in the stream is constantly changing.
Taking three measurements and averaging
the values captures some of the natural variation within the stream and provides a better,
more representative value. Second, taking
more than one measurement reduces the chance
of reporting incorrect data. By running the same
test three times, obvious incorrect values can be
excluded. If more than one person and/or set of
equipment are used, replicates provide an opportunity to test for both operator error and bad
reagents/chemicals. If one person obtains a value
that is considerably different, have them repeat the
test. If only one test is run, it is much more difficult to recognize errors.
3 Star Quality Assurance Guide
Habitat Assessment
Drawing a stream map and taking just a
few physical measurements.
Completing the CQHEI Habitat Evaluation.
Completing the QHEI Habitat Evalua-
Student Groups
tion.
Measurements can be taken by groups of 2-3
students. Tasks within a group include collecting
samples, processing samples, and recording data.
It is very useful to have multiple groups testing for
each parameter (for example, two groups measure
dissolved oxygen). This allows more students to
get involved in quality control. Groups of students conducting the same test should compare
results to determine if the data are similar. If there
are different results for the same sample, students
should check the procedures and redo the test to
determine what caused the difference. Quality
control should be an important part of data collection and the learning experience.
Quality Assurance Levels
Chemical Assessment
Using a less accurate testing method such as presence/absence indicators.
Use of the GREEN LaMotte chemical
testing kit; and/or running only one sample of
each test
Use of the Hach chemical testing kit; and
running each test at least three times and averaging the values (excluding obvious outlying
values)
Biological Assessment
So that volunteers can strive for higher levels of
data quality, we have developed a three star quality rating guide. This will give volunteers an
idea of where their data ranks in terms of quality
assurance. The more stars you get the better. It is
important to note that ALL volunteer data is useful, whether it’s used for education or to promote
erosion control practices within a community. We
do not ask you to report your quality level; this
information is provided to help you develop your
monitoring plan.
Noting the presence of groups of organisms but not counting the individuals; and/or
spending less than 45 minutes on the total biological sampling at your stream site
Reporting the actual number of macroinvertebrates collected (34 mayflies, 8 dragonflies,
etc.); and spending more than 45 minutes on
biological sampling and identification
Reporting the actual numbers of organisms collected; and spending more than 45
minutes on sampling and identification; and
completing the diversity index calculation.
2-9
Chapter 3 - Habitat Study
What is a Healthy Stream Habitat?
Studying the health, type, and condition of the
habitat along the stream banks and corridor is
crucial in determining the health trends of the
stream. The condition of land within and adjacent to the stream channel is critical to the health
of the stream and its ability to support aquatic
life.
A natural stream channel provides a variety of
habitats for many species of plants and animals.
Pools, riffles, undercut banks, and snags (fallen
limbs or small log piles) all provide different
types of habitat. The more types of habitat present in a stream system, the greater the potential
for aquatic plant and animal diversity.
A uniformly straight or deep channel provides
less potential habitat than a stream with variable flows and depths. Examples of healthy and
unhealthy stream habitats are shown in the figure
3-1
What is the riparian zone?
The term "riparian zone" refers to the areas
adjacent to a stream channel (see figure below).
The riparian zone is the strip of land between
the stream channel and the upland hills. Stream
riparian zones form an important transition zone
between land and freshwater systems. "Riparian vegetation" refers to the plants that occur
naturally on stream banks and along stream
channels.
Streamside vegetation is an important component of a stream ecosystem because it provides
streams with bank support and stabilization, ero-
3-2
sion and flood control, water quality protection,
fish and wildlife habitat, and scenic beauty.
Plant roots bind soil to stream banks and reduce
erosion, and deflect the cutting action of swift
flowing stormwater, expanding surface ice, and
strong winds. Streamside vegetation keeps the
water cool by providing shade, and it provides
habitat for aquatic and terrestrial creatures. In
addition, plant debris that falls in upland streams
is a major source of food for organisms in the
stream.
Citizens Qualitative Habitat Evaluation Index (CQHEI)
This index was developed by the Ohio Environmental Protection Agency as a "Citizens"
companion to the Qualitative Habitat Evaluation
Index (QHEI) used by the state's professional
staff. The data sheet and diagrams on the next
three pages were provided by the Ohio EPA.
This index provides a measure of the stream
habitat and riparian health that generally corresponds to physical factors affecting fish and
other aquatic life.
When completing the CQHEI, evaluate your entire stream site (200' section).
1. Stream/River Name: The official name of the stream that is being monitored. The official
name can be found by using a U.S. Geological Survey topographic map. In the case of unnamed
streams, indicate the next named stream into which the unnamed stream drains (e.g. tributary to
Pigeon Creek).
2. Watershed Name and Number: The name and 11-Digit Hydrologic Unit Code (HUC) of the
watershed where your sampling site is located.
I. Substrate (Bottom Type) - Max 24 pts
a) Size - Try to check only one box (the predominant choice).
If two types are close, check two boxes and average the points
b) Smothering - Check one box.
c) Silting - Check one box.
II. Fish Cover (Hiding Places) - Max 20 pts
* Check all the cover types that you see using the picture below as a guide. Add the points.
3-3
III.Stream Shape and Human Alterations - Max 20 pts
a) Curviness or Sinuosity of Channel - Check one box; or
check two boxes and average the points.
b) How natural is the site? - Check one box; or check two
boxes and average the points.
IV.Stream Forests and Wetlands (Riparian Areas)
&
Erosion - Max 20 pts
* For each of these categories, chose the most predominant
answer. If the sides of the stream bank have completely
different characteristics, check two different boxes and
average the score.
a) Width - This is not the width of the stream!
Estimate the width of the area containing trees or
wetlands on each side of the streambed.
b) Land Use
c) Bank Erosion
d) How much of the stream is shaded? - Check one box only.
V. Depth and Velocity - Max 11 pts
a) Deepest Pool - If your stream is a consistent depth (no
pools), select the maximum depth.
b) Check all the flow types that you see - In this category,
check more than one and add the points.
VI. Riffles/Runs (Where current is turbulent)- Max 15
a) Depth & Velocity of Riffles/Runs are - Check one box; or
check two boxes and average the points.
b) Size of Riffle/Run Substrates - Check one box; or check
two boxes and average the points.
MAXIMUM TOTAL POINTS IS 110. If the score is over 100,
consider it "extra credit." You will have an exceptional highquality stream.
3-4
Stream Site Map
Key
Site Map and Stream Flow (Optional)
3-7
Site Map
Drawing a map of your site location is an excellent first step in getting to know your 200 foot stream
segment. Continuing this tradition on an annual basis may also alert you to changes at your site
that may not have been obvious during regular sampling visits. An example map is shown below in
Figure 8.
3-8
Chapter 4 - Chemical Monitoring
Eight Chemical Tests
Many types of chemical tests can be performed
to assess varying aspects of stream water quality. However, volunteer monitoring programs
are faced with both financial and technical limitations. Given these constraints, Miami Valley
Stream Team trains volunteers to conduct eight
of the chemical tests considered by the National
Sanitation Foundation and The Field Manual
for Global Low-Cost Water Quality Monitoring
(Mitchell and Stapp, 1997) to be the most useful
in determining stream water quality (in relative
order of significance):
Dissolved Oxygen
E. coli
Read all of the instructions for each test
before
you perform the procedures.
Practice! The more familiar you are with the
tests, the easier they will be to perform, and
the more accurate your results will be.
Wear protective gloves and safety goggles.
Wash your hands when you are finished.
Obtain your water sample from the stream's
main stream flow (usually in the middle).
Take the sample 3-5 inches under the surface.
See more tips on the next page!
Rinse collection tubes or bottles with sample
water before collecting the sample.
pH
Water Temperature Change
Biochemical Oxygen Demand (5-Day)
Nitrates
Phosphates (Ortho and Total)
Turbidity
Hints For Performing Chemical Tests
Rinse testing tubes and bottles with distilled
water after completing each test.
Clean labware with non-abrasive detergents
or a solvent such as isopropyl alcohol. Use
a soft cloth for wiping or drying. Do not use
paper towels or tissue on plastic tubes as they
may scratch.
The units mg/L and ppm are
equal in fresh water.
They are used interchangeably
throughout this chapter!
4-1
Tips on Collecting Water Samples
How to Discard Chemical Waste
How you physically obtain the water sample
depends on the size, depth, and banks of your
stream. Most volunteers sample wadeable
streams. If you are wading, make sure that you
collect water from a point upstream of where
you are standing, being careful not to stir up any
sediment. Collecting water directly from the
stream with the container used for the chemical
test is the preferred method.
To discard chemical waste, label two separate
containers with secure lids (such as a recycled
margarine or milk containers). Label one "Hazardous Nitrate Waste" (HACH Nitrate Test Only)
and one "Non-Hazardous Chemical Waste."
Place all liquids and solids in the plastic containers along with several cups of clay cat litter.
Allow the liquid to evaporate. The chemical
waste is now in solid form.
Deep water or steep banks are dangerous (see
pictures below). Depending upon conditions at
your site, you may need to use alternative sampling techniques. If you have a bridge at the site,
you may be able to lower a sampling container
or bucket down to the stream. At some sites, you
may be able to sample with a rod from the edge
of the stream. Regardless of the method of collection, sample water should be collected from
the main stream flow.
You can dispose of the Hach Nitrate waste by
calling your local Solid Waste Management
District. Your county may have Household Hazardous Waste Collection Days. All hazardous
waste; including old paint, pesticides, and Hach
Nitrate waste can be taken in or picked up on
these days. They are designed for community
members and are free.
The sample must be collected in a clean container to avoid contamination. Ideally, bottles
and glassware should be acid washed (rinsed
with dilute 10:1 hydrochloric acid solution).
However, isopropyl alcohol and distilled water
are more readily available and can be used for
regular cleaning.
The waste from all the other tests is non-hazardous. Once in solid form, you can throw it
away with your regular trash.
Rinse your container three times with river water
before collecting your final sample. Lower your
container down 3 to 5 inches below the surface
of the water so that your sample is representative
of the whole stream.
Maybe I should re-read
Pictures from GLOBE 1997.
the chapter on SAFETY!
4-2
Chemical Parameters and Test Procedures
Water quality index (WQI)
The Chemical Monitoring Data Sheet utilizes a
Water Quality Index. The Water Quality Index
provides a simple analysis of the results of the
eight chemical tests. If you complete at least six
of the eight tests, you can derive a single score
that will let you know if the stream results are:
Excellent, Good, Medium, Bad, or Very Bad for
that particular monitoring session. You can also
use this value to track changes in your site over
time, or compare the quality with other stream
sites.
instructions. To find the Q-value: locate your
test result on the bottom of the appropriate chart
(x-axis). Draw a vertical line up from your test
result until it intersects the curved line (Q-line).
From this point of intersection draw a line across
to the left hand side (y-axis). Read the number
on the left side of the chart closest to intersection; this is the Q-value for that particular test
result. Record the Q-value in the second column
of the Advanced Chemical Monitoring Data
Sheet. You can also check the Q-value table if
your result is close to a given value.
What does a Q-value mean?
Each of the tests is weighted according to its
level of importance to the overall water quality
(in this particular index). Example: Dissolved
Oxygen has the highest weighting factor (0.18);
therefore, the results are the most important
value in determining the water quality rating using the index.
You can think of a Q-value as a "Quality-value."
It helps interpret your results in terms of the
overall health or water quality of your stream. It
is like a grade for your stream. The higher the
Q-value, the better the test results (100 is the
maximum value; 0 is the minimum).
How to use the Q-value charts
Times and Locations
In order to obtain a WQI Rating, you must first
determine the Q-value for each test. After completing a chemical test, use the results to find the
Q-value on a Q-chart. Each chemical test has
its own Q-chart immediately following the test
The table below provides estimated times for
performing each of the tests and whether they
must be completed on-site or off-site. If samples
are taken off-site, they must be kept on ice or
refrigerated until testing is completed (except
BOD). All tests should be completed as soon as
possible to obtain the best possible results.
4-3
DISSOLVED OXYGEN
Oxygen is as important to life in water as it
is to life on land. Most aquatic plants and
animals require oxygen for survival, and the
availability of oxygen affects their growth and
development. The amount of oxygen found
in water is called the dissolved oxygen (DO)
concentration. Oxygen dissolves readily
into the water from the atmosphere until the
water is saturated. Aquatic plants, algae, and
phytoplankton also produce oxygen as a byproduct of photosynthesis.
DO is an important measure of stream health.
Presence of oxygen in water is a positive sign,
while absence of oxygen from water is often
a sign that the water is polluted. Aquatic
organisms require different levels of DO.
However, dissolved oxygen levels below 3ppm
(parts per million = mg/L) are stressful to most
aquatic organisms. DO levels below 2 or 1ppm
will not support fish. Levels of 5 to 6ppm are
usually required for growth and activity of
aquatic life.
Some of the factors affecting DO are:
• Temperature (water can't hold as much
DO at higher temperatures)
• Altitude/atmospheric pressure
• Turbulence
• Plant growth
• Amount of decaying organic material
Two pieces of information are needed to
interpret Dissolved Oxygen levels – the
DO concentration (in ppm or mg/L) and the
water temperature. From these two values,
the percent saturation can be determined.
Percent saturation is the level of DO in the
water compared to the total amount of DO
that the water has the ability to hold at a
given temperature. Cold water can hold
more dissolved oxygen than warm water. For
example, water at 28°C is 100% saturated
with 8ppm dissolved oxygen. However, water
at 8°C can hold up to 12ppm DO before it
4-4
is 100% saturated. Thus, daily and seasonal
temperature changes, as well as changes caused
by thermal pollution, greatly impact the oxygen
levels and the aquatic life found in streams and
rivers.
High levels of bacteria or large amounts of
rotting organic material can consume oxygen
very rapidly and cause the percent saturation
to decrease. Conversely, water may become
supersaturated for short periods of time,
holding more than 100% of the oxygen
it would hold under normal conditions.
Supersaturation is often caused by high levels
of photosynthesis in streams overloaded with
aquatic plants and algae. Supersaturation may
also occur at the base of dams due to increased
pressure. Supersaturation can be harmful to
aquatic organisms, causing gas bubble disease,
a condition similar to “the bends”, which deep
sea divers may get if they surface too fast.
Problem
Lack of sufficient levels of dissolved
oxygen required by most aquatic
organisms for respiration. Some
organisms have adapted to low
oxygen in water or are able to ingest
air directly.
Causes
Rapid decomposition of organic materials, including dead algae, shoreline vegetation, manure or wastewater decreases oxygen concentrations.
High ammonia concentrations in the
stream use up oxygen in the process
of oxidizing NH4+ to NO3- (nitrification).
At higher temperatures, less oxygen
can dissolve in water.
Lack of turbulence or mixing to expose water to atmospheric oxygen
results in low dissolved oxygen concentrations in the stream.
Dissolved Oxygen
Instructions
These instructions are for use with the HACH
Co. Dissolved Oxygen (DO) Test Kit, Cat. No.
1469-00, Model OX-2P, for 60 mL sample.
1. After rinsing thoroughly with distilled water and
sample water, lower the DO bottle (or other clean
collection bottle) in an upside-down position
to a point well below the water’s surface (3-4
inches). Turn the bottle upright to an angle tilting
upstream. Allow water to flow into the bottle for
approximately 2 minutes until the bottle is full
and no air bubbles are present. While the bottle is
underwater, place stopper in the top. Remove the
bottle from the stream with the stopper in place.
Do not pour off the excess water around the rim
of the stopper. (Note: If pouring your sample
from a collection bottle into the DO bottle, be
careful not to agitate or splash the water into the
bottle.)
2. Add Dissolved Oxygen 1 Reagent and Dissolved
Oxygen 2 Reagent powder pillows to the DO
bottle (the order does not matter). Stopper the
bottle, being very careful not to introduce air
bubbles. (Note: Allow the excess water to spill
over into a waste container. If you get an air
bubble, start over with step one.) With your
thumb firmly holding the stopper in place, grip
the bottle and shake vigorously until the contents
are evenly mixed. A flocculent (floc) will form.
If oxygen is present in the sample, the floc will
appear brownish-orange in color. A small amount
of powdered reagent may remain, but will not
affect the test results. (Note: Oxygen in the
sample is now “fixed” and air bubbles will no
longer affect the test results.)
3. Allow the sample to stand until the floc has
settled below the DO bottle’s white line. The
upper half of the sample will be clear. Shake
the bottle again to remix contents and allow it to
resettle below the white line, again.
(Note: The floc will not settle in samples with high
concentrations of chloride. In general, allow a
maximum of five minutes for the floc to settle, if
no additional progress is made, continue with the
next step.)
4. Add the contents of Dissolved Oxygen 3 Reagent
powder pillow (located in the white tub). Carefully
replace the stopper and shake the bottle to mix.
The floc will dissolve, creating a yellowish-amber
color if DO is present. (Note: Small rust-colored
flakes may remain, but will not affect the test
results.)
5. Fill the sturdy, plastic 5.83 mL measuring tube
(1 cm width x 8.5 cm length) to its top with the
prepared sample, then pour into the square mixing
bottle. (Note: Do not discard the rest of the
fluid in the DO bottle until you have successfully
completed the rest of this test.)
6. Using the dropper located within the brown bottle
marked Sodium Thiosulfate Standard Solution,
add this solution drop by drop to the prepared
sample in the mixing bottle. Count each drop as
it is added and gently swirl to mix the solution
until it becomes colorless. Once the prepared
sample is clear, add one more drop to ensure a
complete color change. If there is no change in
color, do not count this last drop. (Note: Hold
the dropper vertically above the mixing bottle’s
mouth when adding drops to ensure the proper
volume of solution. Do not place the dropper
inside the mouth of the square bottle as you may
contaminate the dropper. Also, rinse any surface,
including your hands, that has contacted the
above chemical as it may “eat” holes in your
clothing and/or irritate your skin.)
7. Each drop added to bring about the color change
in Step 6 equals the presence of 1.0 mg/L of
dissolved oxygen. (Note: If the result of Step 6 is
3 mg/L or less, follow the Dissolved Oxygen LowRange 0.2 - 4.0 mg/L instructions provided on the
next page.)
8. Use the graph in Figure 9 (on the next page), to
calculate percent saturation. By running a straight
edge from the appropriate water temperature
reading to Dissolved Oxygen in mg/L, you will
be able to determine percent saturation along the
angled scale. Look below for a Fahrenheit to
Celsius and Celsius to Fahrenheit conversion.
C = (F – 32.0)/1.80 or F = (C x 1.80)+32
4-5
9. Record DO to the nearest 1.0 mg/L, and record
the Percent Saturation.
Example:
Dissolved oxygen = 8 mg/L
Water temperature = 16oC
Look on chart = 80% Saturation
10. Find the Q-Value (see page 4-3 in this chapter)
and record on the Chemical Data Sheet.
Dissolved Oxygen Lowrange (0.2-4 mg/L)
1. Use the prepared sample left from Step 5 of the
High-range test. Pour off the contents of the DO
bottle until the level reaches the white line (30
mL mark) on the bottle.
2. Add Sodium Thiosulfate Standard Solution one
drop at a time to the DO bottle. Count each
drop as it is added and gently swirl to mix the
solution until it becomes colorless. Once the
prepared sample is clear, add one more drop
to ensure a complete color change. If there
is no change in color, do not count this last
drop.
Typical range for DO =
5.4 to 14.8 mg/L
3. Multiply the number of drops used by 0.2 to
obtain the mg/L Dissolved Oxygen.
Dissolved Oxygen Q-
Example:
# drops used x 0.2 = mg/L Dissolved Oxygen
15 drops x 0.2 = 3 mg/L Dissolved Oxygen
4. Record DO in mg/L and Percent Saturation.
Dissolved Oxygen: % Saturation
4-6
Dissolved Oxygen % Saturation Chart
4-7
E. coli
Fecal coliform bacteria are found in the feces
of warm-blooded animals, including humans,
livestock, and waterfowl. These bacteria are
naturally present in the digestive tracts of
animals, but are rare or absent in unpolluted
waters. Fecal coliform bacteria typically
enter water via combined sewer overflows
(CSOs), poor septic systems, and runoff from
agricultural feedlots. The bacteria can enter the
body through the mouth, nose, eyes, ears, or
cuts in the skin.
E. coli is a specific species of fecal coliform
bacteria used in Ohio's state water quality
standards. Some strains of E. coli can lead
to illness in humans. While not all strains of
E. coli are pathogenic themselves, they occur
with other intestinal tract pathogens that may
be dangerous to human health. We test for
the presence of E.coli as an indicator of fecal
contamination.
Methods
There are several methods available to measure
coliform bacteria. See Appendix A, Monitoring
Equipment, for a list of suppliers. Folow the instructions that are provided with the test method
you choose.
Typical range for E. coli =
133 to 1,157 colonies/100 mL
E. coli Q-Values
E-coli: colonies/100 mL
4-8
pH
The pH test is one of the most common analyses
in water testing. Water (H2O) contains both hydrogen ions (H+) and hydroxide ions (OH-). The
relative concentrations of these ions determine
whether a solution is acidic or basic.
The activity of the hydrogen ions is expressed
in pH units (pH = power of Hydrogen). The
concentration of H+ ions is used to estimate pH.
The pH scale ranges from 1 (most acidic) to 14
(most basic), with 7 being neutral. If the solution has more H+ ions than OH- ions, it is acidic
and has a pH less than 7. If the solution contains more OH- ions than H+ ions, it is basic with
a pH greater than 7. It is important to remember
that pH is measured on a logarithmic scale; it is
reported as the negative log of the hydrogen ion
concentration (-log [H+]). A change of 1 pH unit
means a ten-fold change in the ion concentrations. For this reason, pH units are not normally
averaged; however, to simplify calculations,
Riverwatch allows volunteers to average pH.
The pH level is an important measure of water
quality because aquatic organisms are sensitive
to pH, especially during reproduction. Adult
organisms may survive, but young will not be
produced. A pH range of 6.5 to 8.2 is optimal
for most organisms (see Figure 10).
Many natural processes affect pH. Waterbodies
with higher temperatures have slightly lower
pH values. Also, algae blooms remove carbon
dioxide (CO2) from the water during photosynthesis, which may raise pH to 9 or more.
Runoff from abandoned mine lands can produce
acid main drainage which lowers pH. Lower
pH values increase the solubility of some heavy
metals, such as copper and aluminum, allowing
them to dissolve into the water and harm aquatic
organisms.
Most natural waters have pH values from 5.0
to 8.5. Freshly fallen acidic rainwater has a pH
from 5.5 to 6.0 due to the presence of CO2 in the
atmosphere. Alkaline soils and minerals (e.g.
limestone) can raise pH to 8.0 to 8.5. Seawater
usually has a pH close to 8.0.
4-9
pH Instructions
shake or stir to dissolve the powder completely.
Discard solution after one day.
Company’s portable pH pen, Cat. No. 4435000, Model Pocket Pal Tester.
Step 2: Calibration of the pH pen
1. For accurate results, always begin by
calibrating the pH meter. Follow the
instructions provided in the next column.
2. Once calibrated, place the pH meter into the
stream so that 1/3 of its length is submerged
below the water’s surface. Gently swirl the
meter in the water during the reading.
3. Turn the meter on and allow the digital
reading to stabilize. Once stabilized, record
the reading.
4. After each use, turn off the pH meter to
lengthen the life of the battery. (Note: The
pH meter uses two watch-type batteries.)
Typical range for pH = 6.6 to 8.3
(Note: This procedure can be performed before
you go out into the field. If the pH pen does
not have automatic temperature compensation,
the buffer solution should be at 25°C. Low
temperatures and low conductivity will make
calibration and testing of pH difficult with a pH
pen.)
1. If the pH pen has been stored dry, soak the
electrode in pH 7.0 buffer solution for 24 hours
before calibrating.
2. Rinse the electrode (glass probe) with distilled water using a squeeze bottle and blot dry
with a soft tissue. Do not touch the electrode
with your fingers.
3. Turn the pen on and immerse the electrode
entirely in the pH 7.0 buffer as shown in the
photo.
4. Gently stir the buffer solution with the probe
Calibration Instructions
Step 1: Prepare the Buffer Solution
Pre-mixed liquid buffer solutions can be stored
for 1 year, as long as they have not been contaminated. If you are using a pre-mixed buffer solution, measure 50 mL using a graduated
cylinder or the Hach square mixing bottle and
pour into a clean 100 mL beaker or well-cleaned
baby food jar labeled "pH 7.0 buffer." Discard
solution after one day.
If you are using the powder pillow buffer,
dissolve one pH 7.0 powder pillow in 50 mL
distilled water. Measure 50 mL distilled water
using a graduated cyclinder or the Hach square
mixing bottle and pour into a clean 100 mL beaker or well-cleaned baby food jar labeled "pH
7.0 buffer." Add one pH 7.0 powder pillow and
4-10
and wait for the reading to stabilize.
5. Use a jewelry or eyeglass screwdriver to turn
the small screw in the hole in the back of the
pen until the reading is exactly 7.0.
6. Remove the pH pen from the solution and
rinse the electrode with distilled water.
7. Recheck your pH pen in the Field
Take the pH buffer solution into the field with
you. If the value of the buffer solution is more
than + or - 0.2 pH units from the true value, go
through the calibration procedure again.
pH Q-Values
pH: Units
4-11
BIOCHEMICAL OXYGEN DEMAND (5-Day)
Biochemical oxygen demand (BOD5) is a
measure of the amount of oxygen used by
aerobic (oxygen-consuming) bacteria as they
break down organic wastes over five days.
Polluted streams, or streams with a lot of plant
growth (and decay), generally have high BOD5
levels. High levels indicate that large amounts
of organic matter are present in the stream.
Streams that are relatively clean and free from
excessive plant growth typically have low BOD5
levels.
In slow moving and polluted waters, much
of the available dissolved oxygen (DO)
is consumed by bacteria, which rob other
aquatic organisms of the oxygen needed to
live. Streams with higher DO levels, such as
fast-moving, turbulent, cold-water streams,
can process a greater quantity of organic
material. Therefore, interpretation of BOD5
levels depends upon the conditions of the
stream sampled, as some streams can “handle”
more waste than others. However, in general,
a healthy stream has high DO levels and low
BOD5 levels – be careful not to confuse the two!
The following is a rough guide to what various
BOD5 levels indicate:
Problem
High levels of organic matter and some
ions (ammonia in particular) can lead to
rapid exhaustion of dissolved oxygen.
Causes
Municipal wastewater and septic
tank effluent that has not been completely treated will use up oxygen.
Eutrophication and hot weather can
cause algae blooms. When bacteria
decompose the dead algae, oxygen
is consumed which increases BOD.
4-12
1-2 mg/L BOD5 - Clean water with little organic
waste
3-5 mg/L BOD5 - Fairly clean with some
organic waste
6-9 mg/L BOD5 - Lots of organic material and
bacteria
10+ mg/L BOD5 - Very poor water quality. Very
large amounts of organic material in water.
Instructions
In addition to a black (light-free) bottle, use the
HACH Company’s Dissolved Oxygen (DO)
Test Kit with Cat. No. 1469-00, Model OX-2P,
for 60 mL sample.
1. Rinse, then lower a stoppered black (light-free)
bottle below the water’s surface. Allow water to
flow into the bottle for approximately 2 minutes.
Ensuring that no air bubbles exist, replace the
stopper and remove the bottle from the water.
(Note: Preferably, use a sample bottle that has
a volume at least two times greater than the test
requires.)
2. Place the BOD sample in a light-free location
(e.g., desk drawer) at room temperature and allow it to sit undisturbed for 5 days.
3. After 5 days, remove the BOD bottle and perform
Steps 2 through 8 of the DO test. If results are <3
mg/L, follow the Low-range DO test instructions
(see pages 41-42).
4. Determine the BOD level by subtracting the mg/
L of the BOD sample from that of the original
DO sample taken 5 days prior.
Example: 11 mg/L (original DO)
- 6 mg/L (DO 5 days later)
5 mg/L (BOD5)
Typical range for BOD =
1.1 - 3.3 mg/L
BOD5 Q-Values
BOD5: mg/L
4-13
WATER TEMPERATURE CHANGE (1 mile)
Water temperature is very important to overall
water and stream quality. Temperature affects:
1. Dissolved Oxygen Levels – Colder
water can hold more dissolved oxygen
than warmer water, thus colder water
generally has higher macroinvertebrate
diversity. Warmer water has less dissolved
oxygen. Lower oxygen levels weaken fish
and aquatic insects, making them more
susceptible to illness and disease (See Figure
11).
2. Rate of Photosynthesis – Photosynthesis
by algae and aquatic plants increases with
increased temperature. Increased plant/
algal growth leads to increased death and
decomposition, resulting in increased
oxygen consumption by bacteria.
3. Metabolic Rates of Aquatic Organisms –
Many animals require specific temperatures
to survive. Water temperature controls their
metabolic rates, and most organisms operate
efficiently within a limited temperature
range. Aquatic organisms die when
temperatures are too high or too low.
Thermal pollution (temperature increases)
threaten the balance of aquatic systems. An
increase in temperature may mean that there is
not enough shade along the stream to prevent
the sun from warming it. There may also be a
point source of thermal pollution, such as heated
industrial discharge, entering the stream. If
water temperature decreases within a mile of
the sampling site, there may be a source of cold
water, such as a spring, entering the stream.
Problem
Aquatic organisms have narrow optimal
temperature ranges. In addition warmer
water holds less dissolved oxygen.
Causes
Loss of shading in the riparian zone
can allow water temperature to increase.
Runoff from roads, parking lots, and
dry fields can increase water temperature.
Discharges from municipal wastewaters and many industrial sources
have elevated temperatures.
Months:
4-14
Temperature Change Instructions
1. Place the thermometer below the water’s surface
(e.g., the same depth at which other tests are
performed). If possible, obtain the temperature
reading in the main streamflow.
2. Swirling gently, hold the thermometer in the
water for approximately 2 minutes or until the
reading stabilizes.
3. Record your reading in Celsius. (Note: If you are
using a thermometer that reads only in Fahrenheit, use the following equation to convert to
Celsius: C = (F – 32.0)/1.8
4. Choose a portion of the stream with roughly the
same degree of shade and velocity as in Step 1,
and conduct the same test approximately 1 mile
upstream as soon as possible. Using the same
thermometer will reduce the possibility of equipment error.
Temperature Change Q-Values
5. Calculate the difference between the downstream
and upstream results. Record the temperature
change in Celsius and note if the change is
positive or negative.
6. Find the Q-Value (see page 4-3 in
this chapter) and record on the
Chemical Data Sheet.
Example:
Downstream Temp (Site A)
- Upstream 1-mile Temp
(Site B)
= Temperature Change (+/-)
Water Temp Change:
C
o
4-15
TOTAL PHOSPHATE (PO4) and Orthophosphate
Phosphorus is essential to plant and animal
life, and its presence in the environment is
natural. Problems with phosphorus as a water
pollutant result not from its presence, but from
the addition of excessive amounts. Aquatic
ecosystems develop with very low levels of
phosphorus. The addition of seemingly small
amounts of phosphorus that would have littleto-no affect on terrestrial systems can lead to
problematic algal blooms when added to aquatic
systems.
Phosphorus enters surface waters in organic
matter (dead plants and animals, animal
waste), attached or adsorbed to soil particles,
or in a number of man-made products
(detergents, fertilizers, industry wastes).
Phosphorus is an important nutrient in
fertilizer because it increases terrestrial plant
growth (vegetation). When transported
into aquatic systems, phosphorus increases
aquatic plant growth (e.g. algae, weeds), as
well. When phosphorus levels are too high,
excess plant and algal growth creates water
quality problems. Plants begin to die and
decompose, depleting the dissolved oxygen
supply in the water. This can ultimately
lead to fish kills in some cases. Phosphorus
is also released from the decomposing
plants back into the water, continuing the
cycle. The reaction of the aquatic system
to an overloading of nutrients is known as
eutrophication.
This means that once it is present in an aquatic
system, phosphorus tends to remain there unless
physically removed. Additionally, over time
some of the other forms of phosphates can be
changed into orthophosphates, becoming readily
available for plant growth. For this reason, it is
important to test for all forms of phosphates or
total phosphate levels.
* Results of the total phosphate test (not the
orthophosphate test) are used in the Miami
Valley Stream Team Q-values and the Water
Quality Index calculations.
Phosphorus occurs in waters in the form of
phosphates (PO4). Phosphate levels higher
than 0.03ppm (mg/L) contribute to increased
plant growth. Orthophosphates are one
form of phosphates. Orthophosphates are
dissolved in the water and are readily available
for plant uptake. Thus, the orthophosphate
concentration is useful as an indicator
of current potential for eutrophication.
However, unlike nitrogen and other nutrients,
phosphorus does not have a gaseous phase.
4-16
Problem
Most fresh water is naturally deficient in
phosphorus, and this limits algal growth.
If excessive phosphorus enters the
surface water, it can support rapid algal
growth rates. When the algae die, their
decomposition uses up oxygen and may
produce odors and
algal toxins.
Causes
able, use Isopropyl alcohol and triple rinse
with distilled water. Wrap in aluminum foil to
retain cleanliness. This glassware should be
dedicated for the purpose of analyzing phosphorus samples.
Orthophosphate (OP) is one component that
makes up Total Phosphate (TP), thus OP < TP.
These tests have three levels: low, medium, and
high range. To help determine which level of
the Total Phosphate test to perform, you can
complete the Orthophosphate test first.
Phosphorus occurs naturally in soil.
Suspended sediments from soil erosion
and runoff are often a significant source
of phosphorus. These may enter the
stream via stream bank erosion or
runoff from forestry, agriculture, and
urban lands. In soils that contain it,
phosphorus can desorb from particles
and enter solution.
1. Rinse with distilled water and sample water and fill the square mixing bottle to the 20
mL mark with the water to be tested. Pour the
sample into a clean and rinsed 50 mL Erlenmeyer flask.
Phosphorus can come from manure
sources, such as treatment lagoons,
over fertilized fields, or waterfowl.
2. Open one Potassium Persulfate Powder
Pillow and add its contents to the flask. Gently swirl to mix.
Urban sources of phosphorus may
include: runoff from parking lots,
construction sites, inadequately treated
Total Phosphate
Instructions
Total Phosphate (Cat# 2250-01, Model PO-24)
test, for 25 mL sample, which is included in
the Stream Survey Kit (Cat# 27120-00). These
instructions are not compatible with the Orthophosphate test (for 5 mL sample) included in the
HACH Surface Waters Kit (Cat# 25598-00).
For greater accuracy and safety, it is recommended that this test be performed inside in a
well-ventilated setting.
All glassware to be used for this test should
be acid washed with a dilute hydrochloric acid
solution (10:1) and triple-rinsed with distilled
water before each use. If acid is not avail-
Total Phosphate (All Levels)
3. Add 2 mL of 5.25N Sulfuric Acid Solution
to the sample by twice filling the dropper to
the 1 mL mark and discharging the contents
into the flask. Gently swirl to mix. (Note:
Rinse thoroughly any surfaces, including your
skin, that may have contacted the acidic solution.)
4. Set up the boiling apparatus. (Note: A hot
plate or camping stove is easier and more
reliable than the fuel tablets provided with
the kit.)
5. Boil the sample for 30 minutes, occasionally
adding demineralized or distilled water to keep
the liquid volume near 20mL. Although it is important to let the volume drop low at least once,
do not bring the volume above 20 mL nor let it
boil to dryness.
6. Remove from heat source and allow to cool.
4-17
7.
Add 2 mL of 5.0N Sodium Hydroxide
Solution by twice filling the dropper to the 1 mL
mark and discharge the contents into the flask.
8.
Return sample to the square mixing
bottle. Add distilled water to return its volume
to 20 mL.
9.
Using the prepared sample, follow the
Ortho-phosphate test of the appropriate range,
except read the result as mg/L Total Phosphate
(PO4).
Typical range for Total Phosphate
= 0.01 to 0.17 mg/L
Orthophosphate Low-range test (0-1 mg/L)
1. Rinse and fill the square mixing bottle to the 20mL mark with the water to be tested.
2. Open one PhosVer 3 Phosphate Powder Pillow
and add its contents to the bottle. Gently swirl
to mix. Allow at least two, but not more than
ten minutes for full color development. (Note: If
phosphate is present, a blue color will develop.)
3. Place the mirrors onto the shelf in the color
comparator. Place the Phosphate (blue-violet)
color disk into the comparator. (Note: The
mirrors are used only during the Low–range
Orthophosphate and Total Phosphate tests.)
4. Fill one of the glass color viewing tubes to the
top line with prepared sample. Place the tube on
the right side of the comparator. (Note: Keep the
rest of the prepared sample in the square mixing
bottle until the test is complete. If the results are
greater than 1mg/L, you can use the same sample
for the 0-5 m/L test as explained in the shortcut.)
a color match is obtained. Divide the reading
from the scale window by 50 to obtain the
Orthophosphate (PO43-) in mg/L.
SHORTCUT FOR 0-5 mg/L METHOD!
If the result is > 1mg/L and it has been less
than 10 minutes, use this shortcut. Pour out the
prepared sample and the untreated water from
the test tubes until they reach the 5mL mark.
Remove mirrors from the comparator! Obtain a
color match and read the results using the color
comparator. Divide the reading from the scale
window by 10.
Orthophosphate Medium-range test (0-5
mg/L)
1. Follow steps #1 and 2 of the Low-range test.
2. Fill one of the glass tubes to the bottom line with
prepared sample (approx. 5 mL). Place the tube
on the right side of the comparator.
3. Fill the other glass tube to the bottom line with
untreated sample water and place on the left side.
4. Do not use the mirrors in the comparator. Hold
the comparator up to a light source and rotate the
disc until a color match is obtained. Divide the
reading from the scale window by 10 to obtain
mg/L Orthophosphate (PO43-). (Note: Holding
a piece of white paper 6-8 inches behind the
comparator may help in viewing the color.)
Orthophosphate High-range test (0-50
mg/L)
1. Rinse the square mixing bottle and dropper with
distilled water. Add 2.0 mL of sample by twice
filling the dropper to the 1.0 mL mark.
2. Add demineralized or distilled water to the 20 mL
mark on the mixing bottle. Swirl to mix.
5. Fill the other glass tube to the top line with
untreated sample water and place it on the left.
3. Open one PhosVer 3 Phosphate Powder Pillow.
Add the contents to the bottle. Gently swirl to
mix. Allow at least two minutes, but no longer
than ten minutes, for color development.
6. Do not place caps on the tubes. Orient the
comparator with the tube tops pointing to a
window or light source. Rotate the disc until
4. Fill one of the glass tubes to the bottom line with
prepared sample (approx. 5 mL). Place the tube
on the right side of the comparator.
4-18
5. Fill the other glass tube to the bottom line with
untreated sample water and place it on the left side.
6. Do not use the mirrors in the comparator. Hold
the comparator up to a light source and rotate the
disc until a color match is obtained. Read the mg/L
Orthophosphate (PO4-3) directly on the scale. (Note:
Holding a piece of white paper 6-8 inches behind
the comparator may help in viewing the color.)
Total Phosphate (PO4) Q-Values
Total Phosphorus: mg/L
4-19
NITRATE (0-1, 0-10 mg/L)
Nitrate Nitrogen 0-1 mg/L
Nitrogen makes up about 80% of the air we
breathe, and it is found in all living things.
Nitrogen occurs in water as nitrate (NO3), nitrite
(NO2), and ammonia (NH3). It enters the water
from human and animal waste, decomposing
organic matter, and runoff of fertilizer from
lawns and crops.
1. Rinse the plastic test tubes with distilled water
and the sample to be tested. Fill one test tube to
the lowest mark (the bottom of the frosted band,
approx. 5 mL) with sample water.
Nitrates are an essential nutrient for plant
growth. Similar to phosphates, these are a main
ingredient in fertilizers and can lead to increased
aquatic plant growth when transported into
aquatic systems. Unpolluted waters generally
have a nitrate level below 4ppm (mg/L). Nitrate
levels above 40ppm (mg/L) are considered
unsafe for drinking water. (See introduction
to Phosphates on page 55 for more detailed
discussion of eutrophication and impacts of
nutrients on water quality).
2. Add the contents of the NitraVer 6 Nitrate powder pillow to the sample to be tested. Stopper
the tube and shake for three minutes. Allow the
sample to sit undisturbed for an additional 30
seconds.
3. Add the contents of one NitriVer 3 Nitrite powder pillow to the sample. Stopper the tube and
shake for 30 seconds. A pink or red color will
develop if nitrate is present. Allow at least 10
minutes, but not more than 20 minutes, before
completing Steps 4 through 6.
4. Insert the tube of prepared sample into the right
top opening of the color comparator.
5. Fill a second test tube to the lowest mark with
untreated sample water and place in the left side
of the comparator.
Problem
Nitrogen works with phosphorus to
increase algae growth and cause
eutrophication.
Causes
Nitrogen can come from manure, such
as treatment lagoons and over fertilized
fields.
Nitrogen is the most abundant nutrient
in commercial fertilizers. Runoff from
agriculture, golf courses, and lawns is
high in nitrogen, especially if it rains
Nitrate Instructions
Co. Low Range Nitrate Test Kit 0-1, 0-10 mg/L
as Nitrate Nitrogen, Cat. #14161-00, Model NI14, for 5 mL sample. (This kit has been distibuted in grants since 1998.)
4-20
6. Place the pink disk into the color comparator.
Do not use the mirrors. Hold the comparator
up to a light source. Rotate the disc to obtain a
color match. (Note: Holding a piece of white
paper 6-8 inches behind the comparator may
help in viewing the color.) Read the mg/L nitrate nitrogen (N) through the scale window.
Multiply the reading on the scale by 4.4 to obtain results as mg/L nitrate (NO3).
7. Place waste in Hazardous Waste container. See
safety tips on disposal on page 4-2 of this chapter.
SHORTCUT FOR 0-10 mg/L METHOD!
If the concentration is > 1 mg/L and it has been
less than 20 minutes, you may use your prepared sample from the low-range test to perform the 0-10 mg/L test. Use an eye dropper
to measure 0.5 mL of prepared sample and add
it to an empty test tube, then fill this tube to the
lowest mark with distilled water. Obtain a result
using the comparator and multiply by 10. Then
multiply by 4.4 to obtain final result of mg/L
nitrate NO3.
Nitrate Nitrogen 0-10 mg/L:
1. Rinse the plastic test tubes and a plastic dropper
with sample water. Fill the dropper to the 0.5
mL mark with sample and discharge into one
rinsed test tube.
2. Fill the test tube containing 0.5 mL of sample
to the bottom of the frosted band with distilled
water.
3. Add the contents of one NitraVer 6 Nitrate
powder pillow to the sample to be tested.
Stopper the tube and shake for three minutes.
Allow the sample to sit undisturbed for an
additional 30 seconds.
4. Add the contents of one NitriVer 3 Nitrite
powder pillow to the sample. Stopper the tube
and shake for 30 seconds. A pink or red color
will develop if nitrate is present. Allow at least
10 minutes, but no more than 20 minutes, before
completing Steps 5 through 7.
5. Insert the tube containing the prepared sample
into the right opening of the color comparator.
6. Fill a second test tube to the lowest mark with
untreated sample water and place in the left side
of the comparator.
7. Place the pink disk into the color comparator.
Do not use the mirrors. Hold the comparator
up to a light source. Rotate the disc to obtain
a color match. (Note: Holding a piece of
white paper 6-8 inches behind the comparator
may help in viewing the color.) Multiply that
reading by 10 to obtain the mg/L nitrate nitrogen
(N) present in the sample. Then multiply by 4.4
to obtain mg/L nitrate (NO3).
Typical range for NITRATE (NO3)=
4.18 to 13.86 mg/L
Nitrate NO3 (the value after the result is multiplied by 4.4) is used in the Q-Value chart and
the Water Quality Index data sheet.
4-21
Nitrate (NO3) Q-Values
Nitrate (NO3): mg/L
4-22
TURBIDITY
Instructions
Turbidity is the measurement of the relative
clarity or "cloudiness" of the water. Turbid
water is caused by suspended and colloidal
matter, including clay, silt, organic and
inorganic matter, and microscopic organisms.
These materials scatter and absorb light, rather
than allowing it to shine through the water
column in a straight line. Turbidity should not
be confused with color, since darkly colored
water can still be clear and not turbid.
Turbid water may be the result of soil erosion,
urban runoff, algal blooms, and bottom
sediment disturbances caused by boat traffic or
abundant bottom feeding fish. It is an important
measurement, because light affects both the
biological and chemical reactions in a stream.
If a stream is very turbid, light will not
reach through the water column and many
reactions, especially photosynthesis, will be
limited. When water is turbid, the floating
particles absorb heat from the sun, raising
water temperature and thus lowering dissolved
oxygen levels. The particles can kill fish and
aquatic invertebrates by clogging their gills and
smothering their habitat.
Problem
When light transmission decreases,
algae can only grow on the surface of
the water. The water looks “dirty”, and
organisms on the stream bottom receive no light.
Causes
Most particles come from erosion
and runoff of soils - from fields,
parking lots, or the stream bank
itself.
You can assess turbidity with several types of
equipment, including a homemade Secchi disk
or turbidity tube, as well as a very accurate but
expensive electronic turbidimeter. Given the
financial constraints of the volunteer monitoring
program and the shallow depth of most of the
streams being monitored, Miami Valley Stream
Team teaches the use of a turbidity tube. See
Appendix A for information about making your
own equipment.
For use with a TURBIDITY TUBE:
1. Collect sample water in a bucket or other
container from which you can pour the water
into a calibrated turbidity tube. Do not allow
the sample to settle. (Note: For a more accurate
assessment of stream turbidity, avoid stirring
bottom sediments when sampling at mid-stream.)
2. Slowly pour the sample water into the tube
while looking vertically down into it. When the
water level reaches the point at which you can
barely see the “X” on the bottom of the tube,
stop pouring and record the result in centimeters
or inches. (Note: Placing the tube over a white
surface and allowing air bubbles to dissipate
will help in reading the result.)
4. Repeat the above steps to verify the result.
(Note: Allowing one or two additional people
to repeat the test may help in obtaining a more
accurate result.)
5. To convert the above reading from inches or
centimeters to Nephelometer Turbidity Units
(NTUs), refer to the Q-Value chart located on
the next page.
Typical range for TURBIDITY =
4.5 - 17.5 NTU
Construction activities can have a
large impact on the amount of sediment that enters a stream.
Algae and organic particles also
contribute to turbidity.
4-23
Turbidity Q-Values
ft
150 120 90 60
30
25
20
15
12.5
10
7.5
5
2.5
NTU
4-24
Turbidity: inches
or
cm or NTU/JTU
cm
OTHER RESULTS
Other water chemistry results could include tests
such as ammonia nitrogen, total solids, chlorine,
conductivity, alkalinity, hardness, heavy metals, or pesticides. Any chemical test results you
obtain should be recorded on the data sheet.
AMMONIA NITROGEN
Although this test is not included in the water quality rating, it is included in the HACH
Stream Survey test kit, and can be an indicator
of pollution. Ammonia nitrogen can enter water
systems from several sources including industry,
sewage, and fertilizers. It is a natural degradation product of excretion and decay of dead
organisms. Ammonia is a form of nitrogen that
plants use for growth, but at high concentrations
it is poisonous to animals, including humans.
The toxicity of ammonia increases as pH increases.
Chemical Monitoring
Worksheet & Data
The chemical monitoring worksheet can be taken
into the field to record the results of multiple
samples. Miami Valley Stream Team recommends that volunteers take multiple samples to
assure higher quality stream monitoring results.
Up to three samples can be recorded on this
worksheet. Obvious outliers (results that are
drastically different from other values) should
not be recorded or used in calculations. The average of the test results is calculated then used in
the first column of the Water Quality Index (the
Test Results Column).
1. Rinse test tubes with sample water and fill to the
5mL mark (bottom of the frosted band).
2. Add the contents of one Ammonia Salicylate
Powder Pillow to one tube. Cap and shake until
the powder is dissolved. Wait 3 min.
3. Add the contents of one Ammonia Cyanurate
Powder pillow to the prepared sample. Shake
until the powder is dissolved. Wait 15 minutes
for the color to develop.
4. Insert the tube containing the prepared sample
into the right opening of the color comparator.
Place the second test tube with untreated sample
water in the left opening.
5. Hold the comparator up to a light source. Rotate
disk until a color match is obtained, then read
the result in mg/L ammonia nitrogen.
4-25
Chemical Monitoring Work Sheet
Clear/Sunny
Overcast
Showers
Rain (Steady)
Storm (Heavy)
Clear/Sunny
Overcast
Showers
Rain (Steady)
Storm (Heavy)
Chemical Monitoring Data Sheet Instructions
As you complete each chemical test (or average your results from the Chemical Monitoring Work
Sheet), record the values in the first column of the chemical monitoring data sheet.
Test Results
__________ mg/L
Dissolved Oxygen __________ % saturation
E. coli
__________ colonies/100ml
pH
__________ units
B.O.D. 5
__________ mg/L
H2O Temp Change __________ change in°C
Total Phosphate __________ mg/L
Nitrate (NO3)
__________ mg/L
Turbidity
__________ NTU's
Use the Q-charts or Q-tables in this chapter to derive the Q-values for each test. Record them in the
second column.
Test Results
__________ mg/L
Dissolved Oxygen __________ % saturation
E. coli
pH
__________ units
B.O.D. 5
__________ mg/L
Total Phosphate __________ mg/L
Nitrate (NO3)
__________ mg/L
Turbidity
__________ NTU's
4-28
Q-Value
______
______
______
______
______
______
______
______
After the Q-values have been determined and recorded in the second column, multiply the Q-value
for each test by the Weighting Factor and record the value in the final Calculation column.
Test Results
Q-Value
Weighting
Factor
__________ mg/L
Calculation
Dissolved Oxygen
__________ % saturation
______ X
.18
=
_________
E. coli
______
X
.17
=
_________
pH
__________ units
______
X
.12
=
_________
B.O.D. 5
__________ mg/L
______
X
.12
=
_________
______
X
.11
=
_________
Total Phosphate
__________ mg/L
______
X
.11
=
_________
Nitrate (NO3)
__________ mg/L
______
X
.10
=
_________
Turbidity
__________ NTU's
______
X
.09
=
_________
Once the calculations are completed for each parameter, you can then sum the Weighting Factor
column and the Calculation column. Divide the total of the Calculation column by the total of the
Weighting Factor column to obtain the Water Quality Index (WQI).
TOTALS
Excellent
Good
Medium
90 - 100%
70 - 90%
50 - 70%
Bad
Very Bad
25 - 50%
0 - 25%
WATER QUALITY
INDEX RATING
If you complete all eight tests, the total of the Weighting Factor column is 1.00 (or 100%). If you are
missing one or two tests (but no more than two!) you can calculate an adjusted Water Quality Index
(WQI) Rating. Just follow the same procedures: Divide the total of the Calculation column by the
total of the Weighting Factor column for the tests you completed to obtain the adjusted WQI.
For example, if the E-coli and Total Phosphate tests were not completed above. The
total of the Weighting Factor column would be 0.72, and the total of the Calculation
column would be 54.46.
Total of Calculation column (Divided by) Total of Weighting Factor column
= Adjusted Water Quality Index Rating
54.46 / 0.72 = 75.64 Good !
CHEMICAL MONITORING DATA SHEET
Date ____/____/____
MM
DD
Begin Time _____:_____ (am/pm)
YY
End Time
# Adults __________________
_____:_____ (am/pm)
# Students __________
______
Certified Monitors' Names____________________________________
Organization Name ___________________________________________________________________
Watershed Name _______________________________________ Watershed # _________________
(Please do not abbreviate.)
Current Weather
Clear/Sunny
Overcast
Showers
Rain (Steady)
Storm (Heavy)
Weather in Past 48 hrs.
Clear/Sunny
Overcast
Showers
Rain (Steady)
Storm (Heavy)
WATER QUALITY INDEX (WQI)
You may perform as many of the following tests as you wish; however, at least 6 must be completed
to obtain a Total Water Quality Index value. Divide the total of the Calculation column by the total
of the Weighting Factor column to obtain the Water Quality Index rating.
Test Results
Q-Value
Weighting
Factor
Calculation
Dissolved Oxygen
__________ mg/L
__________ % saturation
______ X
.18
=
_________
E. coli
__________ colonies/100mL
______
X
.17
=
_________
pH
__________ units
______
X
.12
=
_________
B.O.D. 5
__________ mg/L
______
X
.12
=
_________
______
X
.11
=
_________
Total Phosphate
__________ mg/L
______
X
.11
=
_________
Nitrate (NO3)
__________ mg/L
______
X
.10
=
_________
Turbidity
__________ NTU's
______
X
.09
=
_________
TOTALS
Excellent
Good
Medium
90 - 100%
70 - 89%
50 - 69%
Bad
Very Bad
25 - 49%
0 - 24%
WATER QUALITY
INDEX RATING
Chapter 5 - Biological Monitoring
Benthic Macroinvertebrates
Benthic macroinvertebrates are animals that are
big enough (macro) to be seen with the naked
eye. They lack backbones (invertebrate) and
live at least part of their lives in or on the bottom
(benthos) of a body of water.
Macroinvertebrates include aquatic insects (such
as mayflies, stoneflies, caddisflies, midges, beetles), snails, worms, freshwater clams, mussels,
and crayfish. Some benthic macroinvertebrates,
such as midges, are small and grow no larger
than 1/2 inch in length. Others, like the three
ridge mussel, can be over ten inches long.
Why Do We Monitor Them?
Biological monitoring focuses on the aquatic
organisms that live in streams and rivers. Scientists observe changes that occur in the number
of types of organisms present in a stream system
to determine the richness of the biological community. They also observe the total number of
organisms in an area, or the density of the community. If community richness and community
density change over time, it may indicate the
effects of human activity on the stream.
Biological stream monitoring is based on the fact
that different species react to pollution in different ways. Pollution-sensitive organisms such
as mayflies, stoneflies, and caddisflies are more
susceptible to the effects of physical or chemical changes in a stream than other organisms.
These organisms act as indicators of the absence
of pollutants. Pollution-tolerant organisms such
as midges and worms are less susceptible to
changes in physical and chemical parameters in a
stream. The presence or absence of such indicator organisms is an indirect measure of pollution.
When a stream becomes polluted, pollution-sensitive organisms decrease in number or disap-
pear; pollution-tolerant organisms increase in
variety and number.
In addition to being sensitive to changes in the
stream's overall ecological integrity, benthic
macroinvertebrates offer other advantages to
scientists looking for indications of stream pollution.
Benthic macroinvertebrates are relatively
easy to sample. They are abundant and
can be easily collected and identified by trained
volunteers.
They are relatively immobile. Fish can
escape toxic spills or degraded habitats by
swimming away. Migratory animals may
spend only a small portion of their life cycles
in a particular stream before moving to larger
rivers, wetlands, or other streams. However,
most macroinvertebrates spend a large part of
their life cycle in the same part of a stream,
clinging to objects so they are not swept
away with the water's current.
Benthic macroinvertebrates are continuous
indicators of environmental quality. The
composition of a macroinvertebrate
community in a stream reflects that stream's
physical and chemical conditions over
time. Monitoring for certain water quality
parameters (such as the amount of dissolved
oxygen ) only describes the condition of the
water at the moment in time the samples
were taken.
Benthic macroinvertebrates are a critical part
of the aquatic food web. They form a vital
link in the food chain connecting aquatic
plants, algae, and leaf litter to the fish species
in streams. The condition of the benthic
macroinvertebrate community reflects the
stability and diversity of the larger aquatic
food web.
5-1
How Do We Collect Them?
Kick Seine Sampling Method
The kick seine method is a simple procedure for
collecting stream-dwelling macroinvertebrates.
It is used in riffle areas where the majority of
the organisms live. This technique can be quite
effective in determining relative stream health;
however, it is only as good as the sampling
technique. Two to three people work together
to perform the technique properly. Follow the
procedures as closely as possible.
1. Locate a "typical riffle." Such a riffle is a
more shallow, faster moving mud-free section of
stream with a stream bed composed of material
ranging in size from ten-inch cobbles to onequarter inch gravel or sand. The water ranges in
depth from approximately two inches to a foot,
with a moderately swift flow. Avoid riffles located in an area of a stream that has been recently disturbed by anything, including construction
of a pipeline crossing or roadway.
2. Once the riffle has been located, select an area
measuring 3 feet by 3 feet that is typical of the
riffle as a whole. Avoid disturbing the stream
bed upstream from this area.
3. Examine the net closely and remove any
organisms remaining from the last time it was
used.
4. Approach the sampling area from downstream!
5. Have one person place the net at the downstream edge of the sampling area. (It may take
two people to hold it in place.) The net should
be held perpendicular to the flow, but at a slight
downstream angle. Stretch the net approximately three feet, being certain that the bottom
edge is lying firmly against the bed. If water
washes beneath or over the net you will lose
organisms.
5-2
6. Another person comes upstream of the net.
Stand beside, not within the sampling area.
Remove all stones and other objects two inches
or more in diameter from the sampling area.
Hold each one below the water as you brush all
organisms from the rock into the net. You can
also place rocks on the bottom of the net to help
hold it down.
7. When all materials two inches or larger have
been brushed, step into the upstream edge of the
sampling area and kick the stream bed
vigorously until you have disturbed the entire
sampling area. Kick from the upstream edge
toward the net. Try to disturb the bed to a depth
of at least two inches. You can also use a small
shovel to disturb the bed. Kick for at least 2-3
minutes.
8. Carefully remove the net with a forward upstream scooping motion. DO NOT allow water
to flow
over the top of the net or you may
lose organisms.
9. Carry the seine to a flat area on the stream
bank. Remove leaves, rocks, and other debris.
Examine them for any attached organisms. Using fingers or forceps, remove organisms from
the net and place in a plastic container for later
identification. If nothing appears to be on the
net, leave it alone for a few minutes. The insects will begin to move around because they are
out of the water.
10. Perform steps 1-9 a total of three times in
the same riffle or different riffles within your
site.
11. Sort all the organisms collected from the
three samples according to body shape using ice
cube trays or petri dishes. Record the presence
of each type of organism and estimate the number of each type.
•
Undercut Banks (see picture CQHEI
instructions): Place the net below
the surface under the overhanging
vegetation. Move the net in a bottomup motion, jabbing at the bank several
times in a row to loosen organisms. It is
recommended to do 10 "jabs" with the
net.
•
Sediments: (Sampling technique
useful in areas of mostly sand and/or
mud). The person holding the net stands
downstream of sediment area with dip
net resting on the bottom. Another
person begins upstream, kicking and
disturbing sediments to a depth of about
two inches as they approach the net. The
“netter” sweeps the net upward to collect
organisms as the kicker approaches.
Finally, keeping the opening of the net at
least an inch or two above the surface of
the water, wash sediment out of the net
by moving the net back and forth in the
stream water. It is recommended to do 3
"jabs" with the net.
Dip Net Sampling Method
If there are no riffles at your stream site to
perform the kick seine sampling method, then
you should use the dip net to perform your
biological monitoring. Dip nets are useful for
sampling aquatic habitats other than riffles.
One dip net “jab” involves forcing the dip net
against the stream bottom repeatedly, starting
close to your body and finishing with arms fully
outstretched. However, sampling technique
differs depending on specific habitat conditions.
The following is a list of habitat-specific
sampling hints (modified from the Clinton River
Watershed Teacher Training Manual):
•
•
Leaf Pack: Look for leaves that are
brown and slightly decomposed (only a
handful of leaves is necessary). Place
the bottom of the net immediately
downstream from the leaf pack with
handle perpendicular to stream flow.
Gently shake the leaf pack in the
water to release organisms, and then
quickly scoop up the net, capturing
both the organisms and the leaves. It is
recommended to do 3 "jabs" with the
net.
Tree Roots, Snags (accumulations of
debris), and Submerged Logs: Select
an area approximately 3 by 3 feet in
size. Begin working downstream,
scraping the surface of roots, logs,
or debris with the net. You may also
disturb such surfaces with a large stick,
your foot, or by removing some of the
bark to expose hidden organisms. In all
cases, be sure that the net is positioned
downstream from the snag, root, or log,
so that dislodged material floats into it.
It is recommended to do 4"jabs" with
the net.
Take a total of twenty jabs in a variety of habitats.
After two or three jabs are performed with one
net, dump the collected materials from the net
into a shallow white container or bin - a dish
pan works well. The materials in the bin may be
quite muddy and turbid (depending upon your
stream habitat). If this is the case, once you find
macroinvertebrates in your sample, you may
want to place them into another clean container
with a small amount of clear water for easier
identification.
Combination Sampling Method
If your 200 foot sampling site has a variety of
habitats, including riffles, then you may perform
a combination of sampling methods. Record the
types of equipment used and the types of habitats
sampled on the Biological Monitoring Data Sheet.
5-3
How Do They Develop?
Most of the benthic macroinvertebrates you will
encounter are aquatic insects. Aquatic insects
have complex life cycles and live in the water
only during certain stages of development.
Complete Metamorphosis
Aquatic insects may go through one of two kinds
of development or metamorphosis. Those that
go through complete metamorphosis undergo
four stages of development: egg, larva, pupa,
and adult. They lay their eggs in water; eggs
then hatch into larvae that feed and grow in the
water. (These larval insects do not resemble the
adult insects; many appear wormlike.) The fully-grown larvae develop into pupae and then into
adults. The fully-formed adults of some species
(midges and flies, for example) emerge from
the water and live in the habitat surrounding the
stream. Others, such as riffle beetles, continue to
live in the stream as adults. After mating, adults
of all aquatic insect species lay eggs in the water,
beginning the life-cycle all over again.
5-4
Incomplete Metamorphosis
Aquatic insects that go through incomplete metamorphosis undergo only three stages of development; eggs, nymphs and adult. The eggs hatch
into nymphs (also called larvae). Nymphs feed
and grow in the water while they develop adult
structures and organs. The life cycle begins
again when adults lay eggs in the water.
Incomplete metamorphosis:
egg → nymph → adult
(mayfly, dragonfly, stonefly, true bugs)
Complete metamorphosis:
egg → larvae → pupa →
(true flies, beetles, caddisfly)
adult
What and How Do They Eat?
Macroinvertebrates may be categorized by their
feeding groups - the type of food they eat and the
manner in which food is obtained/collected.
Grazer
These organisms graze on algae growing on
rocks in the substrate or on vegetation. Grazers
include snails and water pennies.
Shredder
Shredders feed on coarse, dead organic matter
(leaves, grasses, algae, and rooted aquatic
plants), breaking it into finer material that
is released in their feces. Shredders include
stonefly nymphs, caddisfly larvae, cranefly
larvae.
Collector
Collectors feed on fine, dead organic matter,
including that produced by the shredders.
Filtering collector: filters particles out of flowing current. Examples include blackfly larvae
and net-building caddisflies.
Gathering collector: gathers matter while
crawling along the river bottom. Gatherers
include mayfly nymphs, adult beetles, midge
larvae.
Predator
Predators feed on other invertebrates or small
fish. Mouth parts are specially adapted to feed
on prey. Dragonflies and damselflies have
scoop-like lower jaws, the jaws of hellgrammites
(dobsonflies) are pincer-like, and water strider’s
mouth parts are spear-like. Also includes beetle
adults and larvae.
What Do They Look Like?
A simple key to bethic macroinvertebrates is provided on the following pages. The organisms are
grouped according to pollution tolerance, starting with the most intolerant families. The figure
below may help you identify the distinguishing
features of many of the organisms.
Figures from GREEN Standard Water Monitoring Kit
5-5
Taxonomic Key to Benthic Macroinvertebrates
The purpose of this taxonomic key is to assist volunteer monitors, who are not trained in taxonomy,
with the identification of benthic macroinvertebrates found in the Great Miami River Watershed.
This key is a simplified version of more complex keys. The taxonomic level of this key is intended
for use by citizen monitoring groups. When using this key please note that each couplet offers two
or three options. Each couplet is numbered and the numbers in bold refer to the next couplet (the
next set of numbers that you proceed to).
Please be aware that some macroinvertebrates may have missing body parts so you should look at more than one organism!
CHOOSE ONE:
GO BELOW TO:
2
(1)a Has a shell(s)
(1)b Has no shell
5
(2)a Has a hinged double
3
shell
(2)b Has a single shell
4
(3)a Adult under 2 inches long
19
MUSSEL
(3)b About 2-4 inches long
Right-Handed Snail
(4)a Right-handed opening
RIGHT-HANDED SNAIL
(4)b Left-hand opening
LEFT-HANDED SNAIL
Left-Handed Snail
5-6
CHOOSE ONE:
(5)a Has a segmented body or looks like a tiny tick
GO BELOW TO:
6
(5)b Has an unsegmented body and has an "arrow
PLANARIA
shaped" head; 2 pigment spots (eyes)
Planaria
(6)a No obvious legs
7
(6)b Obvious legs
12
(7)a Has no obvious appendages (long, tubular body)
8
(7)b Has some appendages (small tubes, tiny bumps,
9
or feathery structures)
(8)a Has a smooth body and suckers
LEECH
Leech
(8)b Has a round body and a rat tail
RAT-TAILED MAGGOT
Rat-Tailed Maggot
AQUATIC WORMS
(8)c Has a rounded body
Aquatic Worms
CRANE FLY LARVA
(9)a Body black or brown; more than 1/3 inch long;
plump and catepillar-like
Crane Fly Larva
10
(9)b Has a distinct head
BLACK FLY LARVA
(10)a One end of body wider than other
end; two tiny feather structures on smaller end
Black Fly Larva
5-7
CHOOSE ONE:
GO BELOW TO:
(10)b No difference in diameter along body
11
BLOOD WORM MIDGES
(11)a Bright red body
Chironomid
(11)b Grey Body
OTHER MIDGES
WATER MITE
(12)a Has four pairs of legs
Water Mite
(12)b Has three pairs of legs
13
(12)c Has many pairs of legs
26
(13)a Has no wings or short wing pads on back
14
(13)b Has two pairs of wings that cover the
(14)a Has a flat, round body with legs
underneath (wings are not obvious)
abdomen
Water Penny Beetle
(14)b Not flat, has long body with legs
CADDISLY LARVA
Caddisfly Larva
(15)b Free-living
16
5-8
WATER PENNY BEETLE
15
(15)a
Lives in a tube or a case or
has two
hooks in its
last segment and is green
with 3 plates on back behind head.
(The "green caddis" builds a net &
tube, but will be washed into the
kick net as "free living")
23
CHOOSE ONE:
GO BELOW TO:
(16)a Abdomen possesses lateral filaments
21
similar in size to legs
(16)b Abdomen does not have "leg-like"
17
filaments (may have feathery "gills")
(17)a Always with
STONEFLY NYMPH
Stonefly
Nymph
only two
tail appendages and no
(17)b Usually has three tail appendages, and
18
with no lateral gills on abdominal
segments
(17)c Tail has no appendages
25
Mayfly
MAYFLY NYMPH
(18)a Has long, bristle-like tail appendages,
sometimes 2 or 3
DAMSELFLY NYMPH
(18)b Lower lip formed into extensible scooplike structure and has leaf-like tail
appendages
Damselfly
(19)a Small rounded shell
20
ZEBRA MUSSEL (EXOTIC)
(19)b Small triangular shell with alternating
cream and dark brown bands
Zebra Mussel
(20)a Numerous very fine concentric rows
of elevated lines, white or
cream
colored, with
smooth lateral teeth
(ridge lines on inside near point)
FINGERNAIL CLAM
Fingernail Clam
(20)b Numerous concentric elevated ridges,
yellowish brown to black shell with
serrated lateral teeth
ASIATIC CLAM (EXOTIC)
5-9
CHOOSE ONE:
GO BELOW TO:
(21)a Head narrower than widest body
BEETLE LARVAE
Beetle larva
segments
(21)b Head as wide or wider than other body
22
segments
(22)a Abdomen with single long filament at end
ALDERFLY
Alderfly
(22)b Abdomen ending with a pair of tiny
DOBSONFLY OR FISHFLY
hooked legs, large head with pincer-like
jaws
Dobsonfly Larvae
(23)a Oval shaped body, legs with feathery
ADULT WATER BUGS AND
WATER BEETLES
swimming hairs
Water bug
(23)b
RIFFLE BEETLE ADULT
All legs smooth, without hairs,
Riffle Beetle Adult
(25)a Lower lip formed into scoop like structure
(25)b Looks like a tiny millipede
Dragonfly Nymph
Riffle Beetle Larvae
DRAGONFLY LARVAE
RIFFLE BEETLE LARVAE
SOWBUG
(26)a Flattened top to bottom, crawling looks
like "roly-poly" or a "pill bug"
Sowbug
SCUD
(26)b Flattened side to side, swimming looks
like tiny shrimp
Scud or Side-swimmer
5-10
How to Complete the Biological Monitoring Data Sheet
Sampling Procedures
Equipment: Check one or both of the nets used to collect macroinvertebrate sample.
Habitat: Check each type of habitat sampled during this survey.
Pollution Tolerance Index
The macroinvertebrate index is divided into Pollution Tolerance Groups (PT Group) 1, 2, 3, and 4.
These PT groups represent the different levels of pollution tolerance. The higher the group number,
the higher the pollution tolerance level. Record the number of macroinvertebrates you find here.
PT GROUP 1
Intolerant
Stonefly Nymph
________
Mayfly Nymph
________
PT GROUP 2
Moderately Intolerant
PT GROUP 3
Fairly Tolerant
Damselfly Nymph ________
Midge Larvae
PT GROUP 4
Very Tolerant
________
Left-Handed Snail ________
Dragonfly Nymph ________
Black Fly Larvae ________
Aquatic Worms
________
Caddis Fly Larvae ________
Sowbug
________
Planaria
Blood Midge
________
Dobsonfly Larvae ________
Scud
________
Leech
Riffle Beetle
________
Crane Fly Larvae ________
Water Penny
________
Clams/Mussels
________
________
Rat-tailed Maggot ________
________
Right-Handed Snail ________
The next row is the # of Taxa. Insects that have the same body shape all belong to the same taxa (see
the back of your PTI macroinvertebrate data sheet for general body shape/taxa). To find the total
number of taxa for each PT Group you need to add the number of types of organisms. It is possible
to have a particular PT group without any numbers, therefore it will score a zero.
Do not make the mistake of adding the numbers of organisms together.
# of TAXA
______
# of TAXA
______
# of TAXA
______
# of TAXA
______
The next row is the group scores. Multiply each # of taxa by its weighting factor.
# of TAXA
Weighting
(x 4)
______
______
# of TAXA
(x 3)
______
______
# of TAXA
(x 2)
______
______
# of TAXA
______
(x 1)
______
Factors:
5-11
Then total all of the group scores to get the POLLUTION TOLERANCE INDEX RATING.
# of TAXA
______
# of TAXA
______
# of TAXA
______
(x 4) ______
(x 3)
______
(x 2)
______
23 +
17 - 22
11 - 16
10 or Less
Excellent
Good
Fair
Poor
# of TAXA _______
(x 1)
______
POLLUTION TOLERANCE
INDEX RATING
(Add the final index values for each group.)
Other Biological Indicators
Check the appropriate box if you find native mussels, zebra mussels, or submerged aquatic plants at
your site. Estimate the percentage of algae covering rocks or the stream bottom at your site. Write your
Diversity Index score if you perform the procedure.
Native
Mussels
Zebra Mussels
Submerged
Aquatic Plants
_______ % Algae
Cover
_______ Diversity
Index
Example of a complete Pollution Tolerance Index:
POLLUTION TOLERANCE INDEX (PTI)
PT GROUP 1
Intolerant
Stonefly Nymph
PT GROUP 2
PT GROUP 3
Fairly Tolerant
________ Damselfly Nymph ________ Midge Larvae
PT GROUP 4
Very Tolerant
________ Left-Handed Snail _______
_
Mayfly Nymph
________
Dragonfly Nymph ________ Black Fly Larvae ________
Aquatic Worms ________
Caddis Fly Larvae________ Sowbug
________ Planaria
________
Blood Midge
_______
________
________ Rat-tailed Maggot ________
_
Dobsonfly Larvae ________ Scud
Riffle Beetle
________ Crane Fly Larvae ________
Water Penny
________ Clams/Mussels ________
Weighting
Leech
Right-Handed
Snail________
Factors:
23 or More
17 - 22
11 - 16
10 or Less
5-12
Excellent
Good
Fair
Poor
POLLUTION TOLERANCE
INDEX RATING
BIOLOGICAL MONITORING DATA SHEET
Date ____/____/____ Begin Time _____:_____ (am/pm)
MM
DD
YY
End Time
# Adults __________
_____:_____ (am/pm)
# Students__________
Certified Monitors' Names____________________________________
Organization Name ___________________________________________________________________
Watershed Name _______________________________________ Watershed # __ __ __ __ __ __ __ __
Stream/River Name ____________________________ Site Location:
(Please do not abbreviate.)
Check Methods Used
Check Habitats Sampled
Kick Seine Net (3 times)
Riffles
Undercut Banks
Sediment
D-Net (20 jabs or scoops)
Leaf Packs
Snags/Vegetation
Other
POLLUTION TOLERANCE INDEX (PTI)
PT GROUP 1
Intolerant
Stonefly Nymph
PT GROUP 2
PT GROUP 3
Fairly Tolerant
________ Damselfly Nymph ________ Midges
PT GROUP 4
Very Tolerant
________ Left-Handed Snail _______
_
Mayfly Nymph
________
Dragonfly Nymph ________ Black Fly Larvae ________
Aquatic Worms ________
Caddis Fly Larvae________ Sowbug
________ Planaria
________
Blood Midge
_______
________
________ Rat-tailed Maggot ________
_
Dobsonfly Larvae ________ Scud
Riffle Beetle
________ Crane Fly Larvae ________
Water
Penny
Weighting
________ Clams/Mussels ________
Leech
Factors:
Right-Handed
Snail_______
23 or More
Excellent
17 - 22
Good
11 - 16
Fair
10 or Less
Poor
POLLUTION TOLERANCE
INDEX RATING
Native
Mussels
Zebra Mussels
Submerged
Aquatic Plants
_______ % Algae
Cover
_______ Diversity
Index
5-13
Chapter 6 - Data - What’s Next?
Data Analysis, Action & Evaluation
Analysis involves looking at data and trying to
explain or understand what you’ve found. Often,
collection of data over time reveals patterns
and trends that are extremely useful in data
analysis. It is important to remember that the
data you have collected are interrelated – habitat
evaluation helps to explain macroinvertebrate
presence, which depends upon chemical
parameters, etc. A simple but important question
is: Do my results make sense? If not, what
does not fit? How can this be explained? The
following are useful questions to ask during data
analysis:
How do my results compare to the state
water quality standards?
Are there any noticeable patterns?
Do the results indicate a source of pollution
in the watershed?
Do the test results indicate important water
quality issues facing the community?
What does macroinvertebrate sampling
reveal that is not reflected in chemical testing?
Do the test results seem to correspond to
land use?
Do the CQHEI, Pollution Tolerance Index,
and Water Quality Index scores agree?
Take Action
The following list is a sumary of how to take action to address a water quality issue in your river,
stream, or watershed. However, don’t attempt it
alone - you may want to join a community-based
watershed organization in your area. There are
many groups that are working to locate and correct water resource problems. A list of groups in
the Great Miami River Watershed is included in
Appendix F of this manual.
List any problems that you discovered during
sampling. You may decide that you want to
help resolve a problem that you have identified.
First, you must define who or what is affected
by the problem. For example, fecal coliform
contamination impacts the stream community
and is a threat to human health.
Second, determine the possible actions that you
could take. You may choose to educate others by
speaking to neighbors, at school, or by writing
to the newspaper. You may choose to take
direct action by making lifestyle changes (e.g.
start recycling), organizing a stream cleanup, or
planting vegetation to stabilize stream banks. An
effective way to take action with children is to
write songs or theatrical productions that deal
with the issue. Finally, you may take political
action by speaking at a public meeting or by
writing or visiting public officials.
Third, create an action plan comprised of the
actions you feel will best help solve the problem.
Your plan needs to be realistic and achievable
with available information, have a designated
time frame, and yet still be challenging and
interesting to you and your group. Work locally
with people in your community.
Finally, implement your plan. Divide
tasks among group members and interested
participants and set timelines for each step, as
well as an overall deadline. Record meetings
and monitor your progress. We encourage
volunteers to use their data to take action at a
local level.
Evaluate the River Study
Evaluation of your river study is important,
as it helps to identify successes and improve
future monitoring efforts. Consider whether
or not you were able to meet the goals you set
prior to beginning stream monitoring. Was
time a major limitation? Did you take on too
many sampling sites? Did you feel comfortable
using the equipment, or would another training
workshop be helpful? What did you learn? If
you developed an action plan, was it successful?
Volunteer Stream Monitoring Manual
6-1
Guide for Water Quality Ranges
Dissolved Oxygen (% Saturation)
Total Phosphate (mg/L)
91-110
Excellent
<.10
Excellent
71-90, >110
Good
.11-.16
Good
51-70
Fair
.17-.58
Fair
<50
Poor
.59-2.99
Poor
>3.0
Very Poor
E.Coli (colonies per 100ml)
<50 colonies
Excellent
*Nitrate Nitrogen (N) (mg/L)
51-200 colonies
Good
<0.3
Excellent
201-1000 colonies
Fair
.4-.8
Good
>1000 colonies
Poor
.9-1.9
Fair
>2.0
Poor
pH (Units)
* Not multiplied by 4.4
6.5-7.5
Excellent
6.0-6.4, 7.6-8.0
Good
Turbidity (NTUs or ft/inches)
5.5-5.9, 8.1-8.5
Fair
1-10 or >3'
Excellent
<5.5, >8.6
Poor
10.1-40 or 1-3'
Good
40.1-150 or 2"-1' Fair
BOD (mg/L or ppm)
>150 or < 2"
<2
Excellent
2.0-4.0
Good
Total Solids (mg/L)
4.1-10
Fair
<100
>10
Poor
100-250
Good
6-2
Poor
Excellent
Pollution Indicators Table
From the Field Manual for Global Low-Cost Water Quality Monitoring by Stapp and Mitchell, 1995.
6-3
Habitat Parameters for Selected Macroinvertebrates*
* The values provided are preferred ranges for most species of these groups of organisms.
From GLOBE Manual 1997.
6-4
6-5
6-6
Volunteer & Organization Registration
The Registration page is a form that only needs
to be completed once, or if there are changes or
updates. Each volunteer participating in the Miami Valley Stream Team program must register
themselves and their stream sites before data will
be accepted into the database.
1. Certified Monitor’s Name: The name
of the volunteer who has attended a Miami
Valley Stream Team training session and was
present during data collection.
2.
Organization Registration
Organization registration helps track overall
participation by groups.
1. Organization Name and Contact Information: Same as above.
2. Homepage URL
Organization Name: The name of the
organization, agency, corporation, school,
class, troop, or group performing the volunteer monitoring activities. PLEASE do not
abbreviate the name.
3. Contact Information (Address, City,
State, Zip, Phone, Fax, E-mail): Please
also provide your phone, fax, and e-mail (if
applicable), so that we may contact you with
timely information if necessary.
4. Training Workshops Attended: Only
certified trained monitors may enter data in the
statewide Internet database. Exceptions may be
made on an individual basis for attendance at
other training workshops (e.g., GREEN).
5. Year: Year training was attended
6. Location: List closest town and site (e.g.,
town park, state park, school, 4-H building)
7.
Instructor: Name of training instructor
6-7
VOLUNTEER STREAM MONITOR REGISTRATION
Certified Monitor’s Name __________________________________
Organization Name ________________________________________________________________
Address _________________________________________________________________________
City _______________________________________ State ______
Phone (
) ____________ Fax (
Zip ____________________
) _____________ E-mail __________________________
TRAINING WORKSHOPS ATTENDED
Year ________ Location _______________________ Instructor ________________
ORGANIZATION REGISTRATION
Organization Name ________________________________________________________________
Name of Primary Contact(s)__________________________________________________________
Address _________________________________________________________________________
City _______________________________________ State ______
Phone (
)___________ Fax (
Zip ____________________
)____________ Homepage URL ______________________
STREAM SITE REGISTRATION
Stream/River Name _______________________________________ Site ID__________________
Nearest City/Town __________________________ County _____________________ State _____
Description of Location_____________________________________________________________
Watershed Name ____________________________________ Watershed #___________________
Latitude (North) ___________________________ Longitude (West)________________________
Source of Latitude / Longitude Data___________________________________________________
6-8
Stream Site Registration
1. Stream/River Name: The official name of
the stream that is being monitored. The
official name can be found by using a U.S.
Geological Survey topographic map. In
the case of unnamed streams, indicate the
next named stream into which the unnamed
stream drains (e.g. tributary to Mad River).
2. Nearest City/Town: The nearest community
to your sampling site.
3. County and State: The county and state in
which your sampling site is located.
4. Description of Location: Brief explanation of
your site location in relation to nearby roads,
bridges, dams, other landmarks or other
waterways.
5. Watershed Name and Number: The name
and 8-Digit Hydrologic Unit Code (HUC)
of the watershed where your sampling site is
located.
6. Latitude and Longitude: Please provide
geographic data in degrees, minutes, and
seconds.
Internet Site: Many sites on the Internet allow
you to pinpoint your latitude and longitude
using computer generated maps. A few of these
internet sites are listed below.
TopoZone - http://www.topozone.com
Map Blast - http://www.mapblast.com
U.S. Census Bureau - http://
www.census.gov/cgi-bin/gazetteer
USGS Geographic Names Info. System
- http://mapping.usgs.gov/www/gnis
Topographic Map: You can approximate
your site location using a topographic map.
Information on obtaining a topographic map is
provided in Appendix A.
Weather
Before you enter data into the website, you
are required to enter the weather conditions
during the time of sampling, and 48 hours prior
to sampling. Please submit the worst weather
conditions during this time period as they may
significantly impact water quality.
7. Source of Latitude and Longitude Data:
GPS: One of the most accurate methods for
determining site location is a Global Positioning
System (GPS) receiver. This device picks up
signals from satellites orbiting the Earth and
instantly displays the latitude and longitude
(and altitude, if desired) of your location. To
work properly, a GPS receiver must be able to
"see" the sky in order to locate the satellites. A
cliff, or even a dense forest can interfere with
your ability to get a good read-out. Hand-held
models are available at some outdoor stores, and
the prices are getting lower as the technology
improves.
6-9
Record-keeping Form
(If you have more than one site, copy this recordkeeping form. Use a separate form for each site.)
YEAR:
Date of
6-10
Date(s) of
Data Entry
Completed by
Appendix A - Equipment
Check list
The following supplies may be useful in monitoring the water quality of your local river or stream:
Site Assessment
>
>
>
>
>
>
>
Maps (e.g., 7.5” topographic map, county property maps indicating property boundaries)
Compass and survey tape for marking boundaries
Clipboard, writing utensils, and laminated copies of chemical, biological, and habitat data sheets
Tape measure or twine marked in one-meter/foot lengths
Stopwatch for measuring stream flow
Apple, orange, or other biodegradable object that can be floated to measure stream flow
Yardstick or other device to measure depth
Biological Assessment
>
>
>
>
>
Kick seine net, dip net, or other tools for collecting benthic macroinvertebrates
Sieve and trays for sorting biological samples (ice cube trays work well)
Tweezers, hand lens, magnifying glass, and possibly a microscope
Glass vials or jars filled with isopropyl alcohol or white vinegar for storing insects
Handmade Hester-Dendy Substrate Sampler to use in waterways too deep to enter on foot
Chemical Assessment
>
>
>
>
>
>
>
Chemical water quality testing equipment will vary with the type of monitoring you wish to pursue.
Equipment for each test will vary in range, sensitivity, and cost depending on the use of chemical or electronic
materials. Some equipment can be made by hand.
If using an electronic pH meter, need pH buffer and a small screwdriver for calibrating.
Handmade extension sampling rod
Distilled water for rinsing sampling bottles and tubes
Secchi disk or handmade turbidity tube
2 containers filled with kitty litter to store waste materials (one for nitrate and one for other waste)
Material Safety Data sheets for every chemical being used
Safety
>
>
>
>
Boots or waders (WARNING: Never put children in chest-high waders because they can fill with dangerous
amounts of water if submerged.)
Rubber gloves and protective eyewear
First Aid kit that includes eyewash
Washing water, soap, and a towel
Other Supplies
>
>
>
>
>
Drinking water
Calculator
Camera for documenting site
Computer and Internet access for entry of water quality data
Trash bags or other waste containers for a streambank clean-up
A-1
Where to Purchase Equipment
CHEMICAL TESTING KITS
Hach Co.
PO Box 389
Loveland, CO 80539
(800) 227-4224
www.hach.com
* Stream Survey Water
Testing Kit #27120-00
LaMotte
PO Box 329
Chestertown, MD 21620
(800) 344-3100
www.lamotte.com
* water quality testing kits
Earth Force
1908 Mount Vernon Ave
Alexandria, VA 22301
(703) 519-6877
www.earthforce.org
* GREEN Standard and
Low-Cost Water Kits
NETS
Nichols Net and Twine Co.
2200 Highway 111
Granite City, IL 62040
(800) 878-6387
* Stream Monitor Kick Seine
* Aquatic Dip Net (Like “D”-net)
Ben Meadows Co.
PO Box 80549
Chamblee, GA 30366
(800) 628-2068
* D-frame Net #224902
* Transparency Tube #22196
E-COLI TEST
RIVERS CURRICULUM
Micrology Laboratories
PO Box 340
Goshen, IN 46527
(888) EASY-GEL
www.micrologylabs.com
* E-coli test kit- EASYGEL
Dr. Bob Williams
Southern Illinois University
(618) 650-3788
www.siue.edu/OSME/river
*Biology, Earth Science,
Chemistry, Mathematics, Language Arts, Geography
BOOKS and MANUALS
Kendall Hunt Publishing
PO Box 1814
Dubuque, IA 52004
(800) 228-0810
* Field Manual for Global LowCost WaterQuality Monitoring
by Mitchell and Stapp
Jones and Bartlett Publishers
40 Tall Pine Drive
Sudbury, MA 01776
(800) 832-0034
*Aquatic Entomology
by McCafferty
U.S. EPA - OWOW
401 M Street, SW
Washington DC, 20460
(800) 490-9198
*Volunteer Stream
Monitoring Methods Manual
Doc. #841B97003
Adopt-A-Stream Foundation
600-128th Street SE
Everett, WA 98208
(206) 316-8592
* Streamkeeper's Field Guide by Murdoch and Cheo
This list contains just a few of the many science equipment suppliers available. It is not intended to be an endorsement of any product or
company.
A-2
How to Clean and Care for Equipment
Nets
To ensure that no contamination occurs between sampling sites, make sure that all nets and organism
collection equipment have been cleaned of all organisms and matter. Be sure to rinse them thoroughly before transporting to another location.
pH Meter
Using the HACH Company’s portable pH meter catalog number 44350-00, Pocket Pal Tester. After each use,
turn off the pH meter. This will lengthen the life of the battery. If the batteries die, they can be replaced with
appropriately-sized watch-type batteries. If the bulb (glass electrode) cracks, the pH meter must be replaced and
should be properly disposed.
Turbidity Tubes
(From the Minnesota Citizen Stream Monitoring Program, "Stream Reader" Spring 2000)
If you monitor a stream that is on the murky side, chances are the walls of your turbidity tube have clouded up.
Try cleaning the inside of your tube by filling it three-quarters full with tap water, add a couple drops of dish
soap, and push a clean, soft rag or washcloth down the tube with the end of a broom handle, scrubbing the sides.
If you take the stopper out of the bottom, be sure to fit it back into the tube securely.
If your tube has a release tube and valve, it may become crimped. Try moving the position of the clamp on your
release valve from time to time. By doing this, the tube won't break down and get crimped in any one spot.
A-3
How to Make Your Own Equipment
Not all of your water monitoring equipment has to be purchased through a catalog or at a store. Nets
and other sampling supplies can be made at home.
Kick Seine Net #1
Materials:
>
3 foot by 6 foot piece of nylon or fiberglass screening (white, if you can find it)
4 strips of heavy canvas (6 inches by 36 inches)
2 broom handles or wooden dowels (6 feet long)
> finishing nails
> sewing machine and thread
> hammer
> iron and ironing board
>
>
Directions:
1. Fold screening in half (3 foot by 3 foot).
2. Fold edges of canvas strips under 1/2 inch and press with iron.
3. Sew 2 strips at top and bottom of screening, then use remaining 2 strips on the sides of the screening to
make casings for handles. Sew bottom of casings shut.
4. Insert handles into casings and nail into place with finishing nails.
A-4
Kick Seine Net #2
Directions:
1. Fold one 8 x 122 cm strip of fabric over one of the long screen edges and sew, reinforcing the edge.
2.
Repeat for the other long edge.
3. Attach screen to poles with staples, making
the poles even with the bottom of the screen and
extending to form handles at the top.
4. Wrap screen around poles several times and
staple again to reinforce the edges.
Dip Net
Directions:
1. Cut a net shape from the 36 x 53 cm pieces of nylon
screen (see diagrams) and sew them together.
2. Edge the open end of the net with heavy fabric, leaving an opening to form a casing to insert the hanger.
3. Cut hooks from hangers and untwist the wires.
4. Use duct tape to tape the hangers together to make
your frame heavier.
5. Insert a wire through the casing and twist ends back
together at opening.
6. Drill a hole in the tip of the wooden handle large
enough to insert the ends of the hangers into the hole
in the pole. Secure the net to the pole by using the
hook you cut from the hanger and using the pipe
clamp or duct tape to secure the hook to the pole.
A-5
Turbidity Tube
For instructions on how to correctly use the turbidity tube see Chapter 5 Chemical Monitoring.
Directions:
1. Obtain a 1 and 1/2 inch plumbing cap and a length of florescent light bulb tube cover (2 feet works great). On
the inside of the plumbing cap, use a permanet black marker to draw the diagram pictured below.
2. Place the plumbing cap over one end of a clear tube. Cap should fit tightly so water cannot leak out. If necessary - use glue to secure the cap.
3. Use a marker and meter stick to make a scale on the side of the tube, beginning at the disk with 0 cm. Or mark
on a piece of tape and stick it to the outside of the tube.
A-8
Appendix B - Glossary
A
acid mine drainage: Waters of low pH (less than 6) from mining areas.
algae: Small plants which lack roots, stems, flowers, and leaves; living mainly in water and using the sun as an energy
source.
alkalinity: A measurement of water’s ability to neutralize acid.
aquatic habitat: All of the areas in a stream, lake or wetland that are occupied by an organism, population or
community.
aquifer: Any geological formation containing water, especially one that supplies water for wells, springs, etc.
B
banks: The portion of the stream channel which restricts the movement of the water out of the channel during times of
normal water depth. This area of the stream is characterized as being the exposed terrestrial areas on either side of the
stream.
benthic: An adjective which describes all things associated with the bottom, or sediments of a stream.
bedrock: Unbroken solid rock, overlain in most places by soil or rock fragments.
biochemical oxygen demand (BOD): An empirical test in which standardized laboratory procedures measure the
oxygen required for the biochemical degradation of organic material, and the oxygen used to oxidize inorganic
materials, such as sulfides and ferrous iron.
C
channelization: The straightening of a stream or the dredging of a new stream channel to which the stream is diverted. A
channelized stream is straight with little or no meanders.
class: A taxonomic rank which falls under the taxonomic rank of Order.
cobble streambed: A watercourse predominately lined with naturally rounded stones, rounded by the water’s action.
Size varies from a hen’s egg to that used as paving stones.
complete metamorphosis: The type of insect development that includes four stages; egg, larva, pupa, adult.
conservation practice: An engineered structure or management activity that eliminates or reduces an adverse
environmental effect of a pollutant and conserves soil, water, plant, or animal resources.
D
drainage basin: The total land area draining to any point in a stream. A drainage basin is composed of many smaller
watersheds.
Dissolved Oxygen (DO): The amount of oxygen dissolved in water. Generally, proportionately higher amounts of
oxygen can be dissolved in colder waters that in warmer waters.
E
ecology: The relationship between living things and their environments or the study of such relationships.
effluent: A discharge of partially or completely treated pollutants into the environment; generally used to describe
discharge into the water.
emergent plants: Plants rooted in the bottom of the watercourse, that rise above the water surface.
erosion: The wearing away of the land surface by wind or water.
eutrophic: A highly productive water body, can be caused or accelerated by the input of large amounts of nutrients from
human sources.
eutrophication: Natural eutrophication is the process of lake aging. Cultural eutrophication occurs when nutrients are
added from agricultural runoff, sewage, or other sources.
B-1
Escherichia coli (E. Coli): A bacterium of the intestines of warm-blooded organisms, including humans, that is used as
an indicator of water pollution for disease producing organisms.
F
floodplain: An area on both sides of a stream where flood waters spread out during high rains. The surface may appear
dry for most of the year, but it is generally occupied by plants that are adapted to wet soils.
fecal coliform bacteria: The portion of the coliform group which is present in the gut or feces of warm-blooded animals.
The presence of fecal coliform bacteria in water is an indication of pollution and potential human health problems.
food chain: A transfer of energy in a sequence of organisms (algae, fish, etc.) in a community in which each member of
the chain feeds on the member below it.
H
habitat: The area in which an organism lives.
herbaceous vegetation: Plants having a stem that remains soft and succulent during the growing, not woody.
I
incomplete metamorphosis: The type of insect development that consists of three stages; egg stage, a nymph stage and
an adult stage.
indicator organism: Organisms which respond predictably to various environmental changes, and whose presence or
absence, and abundance, are used as indicators of environmental conditions.
inorganic: Any compound not containing carbon.
intermittent stream: A watercourse that flows only at certain times of the year, receiving water from springs or surface
sources; also, a watercourse that does not flow continuously, when water losses from evaporation or seepage exceed
available stream flow.
invertebrate: An organism without a backbone.
J
JTU’s - Jackson Turbidity Units: a unit of measurement commonly used in electronic turbidity meters that indicate how
far light can penetrate into a water sample before the cloudiness of the sample cuts the light. Similar to NTU’s or
Nephelometer Turbidity Unit.
L
lake: A body of fresh or salt water of considerable size, whose open-water and deep-bottom zones (no light penetration
to the bottom) are large compared to the shallow-water (shoreline zone, which has light penetration to its bottom.
M
macroinvertebrates: Animals lacking backbones that are large enough to be visible without the aid of a microscope.
meanders: Sinuosity, or snake-like curving of a natural stream channel.
metamorphose: To change into a different form, such as from an insect pupa to an adult.
methemoglobinemia: The presence of methemoglobin in the blood, making the blood useless as a carrier of oxygen.
Methemoglobin, a compound closely related to oxyhemoglobin, is found in the blood following poisoning by certain
substances, such as nitrate. Young babies, both human and animal, are particularly susceptible to methemoglobinemia,
leading to a condition known as “blue baby” which if untreated can cause death.
mollusk: Soft-bodied (usually hard-shelled) animals such as clams or mussels.
B-2
N
nitrogen: A limiting nutrient for the aquatic environment. Nitrogen is considered to be limiting because it is needed by
the plants and animals in the stream in moderate amounts. When present in higher amounts, such as large amounts of
fertilizer runoff from local farm fields, large algal blooms occur which cause a depletion of dissolved oxygen.
nonpoint source pollution: A type of pollution whose source is not readily identifiable as any one particular point, such
as pollution caused by runoff from streets and agricultural land.
nutrient: Any substance which is necessary for growth of living things.
nymph: A juvenile, wingless stage of an insect.
NTU - Nephelometer Turbidity Units: a unit of measurement commonly used in electronic turbidity meters that indicate
how far light can penetrate into a water sample before the cloudiness of the sample cuts into the light. Similar to
Jackson Turbidity Units.
O
order: Taxonomic grouping of related families of organisms.
organic material: Any compound containing carbon.
P
pathogenic: Capable of causing disease.
pH: The measurement of acidity or alkalinity on a scale of 0 - 14. A pH of 7 is neutral, less than 7 is acidic, and more
than 7 is alkaline (basic).
phosphorus: An essential plant nutrient that, in excessive quantities, can contribute to the eutrophication of water
bodies.
photosynthesis: Process by which green plants use sunlight to produce food.
perennial stream: A watercourse that flows continuously throughout the year and whose upper surface generally stands
lower than the water table in the area adjacent to the watercourse.
pollution sensitive organisms: Those organisms which cannot withstand the stresses applied on the aquatic environment
by pollution.
pollution tolerant organisms: Those organisms which can withstand many of the stresses applied to an aquatic
environment by pollution.
point source pollution: Pollutants originating from a “point” source, such as a pipe, vent, or culvert.
pond: A body of fresh or salt water, smaller than a lake, and where the shallow-water zone (light penetration to its
bottom) is relatively large compared to the open water and deep bottom (no light penetration to the bottom).
pools: In a watercourse, an area often following a rapids (riffle), which is relatively deep with slowly moving water
compared to the rapids.
pupa: The stage of an insect in which it is enclosed in a protective case while changing from larva to an adult.
R
riffle: In a watercourse, an area often upstream of a pool, which is relatively shallow with swiftly moving water
compared to the pool.
riprap: Any material (such as concrete blocks, rocks, car tires or log pilings) which are used to protect a stream bank
from erosion.
riparian zone: An area, adjacent to and along a watercourse, which is often vegetated and constitutes a buffer zone
between the nearby lands and the watercourse.
runoff: Water from rain, snowmelt, or irrigation that flows over the ground surface and runs into a water body.
B-3
S
sediment: Soil, sand, and minerals washed from land into waterways.
sedimentation: The process by which soil particles (sediment) enter, accumulate and settle to the bottom of a waterbody.
septic odor: The sulfur (rotten egg) smell produced by the decomposition of organic matter in the absence of oxygen.
sewage: The organic waste and wastewater produced by residential and commercial establishments.
sewage treatment plant: A facility designed to remove organic pollutants from wastewater.
silt: Fine particles of soil or rock that can be picked up by air or water and deposited as sediment.
siltation: The process of silt settling out of the water and being deposited as sediment.
submergent rooted plant: An aquatic plant whose roots are in the watercourse’s bottom with the upper part of the plant
submerged below the surface of the water.
substrate: The surface upon which an organism lives or is attached.
species: A unit of classification for a group of closely related individuals.
stream bed: The bottom of a stream where the substrate and sediments lay.
stream depth: A measurement of the depth of a stream from the water’s surface to the stream bed.
stream flow: The amount of water moving in a stream in a given amount of time.
T
tolerant species: An organism that can exist in the presence of a certain degree of pollution.
topographic map: A map representing the surface features of a particular area.
total coliform bacteria: A group of bacteria that are used as an indicator of drinking water quality. The presence of total
coliform bacteria indicates the possible presence of disease-causing bacteria.
total suspended solids: Whole particles carried or suspended in the water, such as silt, sand or small algae or animals,
that cause a green or brown color in the water. These substances can be filtered out of the water and weighed.
total dissolved solids: Substances that are dissolved in the water which can color the water brown or yellow. Tannic
acids that leach from tree roots or from decomposing leaves can color the water brown to black due to dissolved
chemicals. This color does not disappear by filtering the water.
toxicity: A measurement of how poisonous or harmful a substance is to plants and animals.
turbidity: The presence of sediment in water, making it unclear, murky or opaque.
trend data: Data or measurements of a stream system which will show how particular characteristics changed over time.
U
urban runoff: Water which has drained from the surface of land which is used for urban uses, such as paved roads,
subdivisions an parking lots.
W
wastewater: Water carrying unwanted material from homes, farms, businesses and industries.
water quality: The condition of the water with regard to the presence or absence of pollution.
watershed: The entire surface drainage area that contributes water to a stream or river. Many watersheds which drain
into a common river make a drainage basin.
woody vegetation: Plants having a stem or trunk that is fibrous and rigid.
B-4
Appendix C - Suggested Reading*
Water Quality Monitoring Resources
*All prices subject to change.
Field Manual for Global Low-Cost Water Quality Monitoring. 2nd ed. 1997. M.K. Mitchell and W.B. Stapp. 334
pp. $20. Illustrated guide to methods for conducting most common water quality monitoring tests, including turbidity,
phosphorus, nitrogen, fecal coliforms, insect collection, and watershed land use analysis. This book is used by most
school and volunteer groups in Indiana as standard methods for water testing. ISBN# 0-7872-2375-1 Available from:
Kendall Hunt Publishing Co. P.O. Box 1840, Dubuque, IA, 52004.Tel. (800) 338-8309.
Streamkeeper’s Field Guide: Watershed Inventory and Stream Monitoring Methods. T. Murdoch and M. Cheo.
1996. $29.95 Adopt-A-Stream Foundation, Everett, WA. 296 pp. ISBN 0-9652109-0-1. Excellent manual on citizen
assessment and monitoring of streams and watersheds. Available from: The Adopt-A-Stream Foundation, 600 128th
Street SE, Everett, WA 98208. Tel. (206) 316-8592. (The foundation also has guides on wetland assessment and several
beautiful posters on streams, wetlands, and salmon.)
Pond and Brook: A Guide to Nature in Freshwater Environments. M.J. Caduto, 1990. Excellent introduction to
aquatic biology, from wetlands and deep lakes to streams and vernal ponds, for the amateur naturalist, including handson projects and activities. ISBN 0-87451-509-2. $12.95. Available from: Patricia Ledlie Bookseller, Inc., Buckfield,
Maine 04220. Tel or FAX (207) 336-2778 (and at most larger bookstores).
Rapid Bioassessment Protocols for Use in Streams and Rivers: Benthic Macroinvertebrates and Fish. EPA/440/489-001. & Macroinvertebrate Field and Laboratory Methods for Evaluating the Biological Integrity of Surface
Waters. EPA/600/4-90/030. 256 pp. These two publications explain the standard methods used by EPA for sampling
insects and fish in streams. Available from: Clean Lakes Program, Assessment and Watershed Protection Division (WH553), U.S. Environmental Protection Agency, 401 M Street, S.W., Washington, D.C. 20460, Tel. (513) 569-7562.
The Volunteer Monitor. National newsletter of water quality monitoring in which issues address home construction of
monitoring equipment, collecting and analysis of data, and networking. Available from: River Network, 520 SW 6th
Ave., Suite 1130, Portland, OR 97204, [email protected], Tel. (503) 241-9256.
Stream Resources
Better Trout Habitat: A Guide to Stream Restoration and Management. C.J. Hunter. 1991. Island Press. 320
pp. ISBN: 0-933280-77-7. A less technical illustrated guide to stream restoration techniques, including streambank
stabilization and fish habitat rehabilitation. Most work addressed in the book would require state or federal permits on
private or public land and should be designed by a competent professional engineer. Available from: Island Press, Box
7, Covelo, CA 95428. Tel. (800) 828-1302. (Catalogs from Island Press include an excellent listing of unusual books
on environmental issues.)
Entering the Watershed: A New Approach to Save America’s River Ecosystems. B. Doppelt, M. Scurlock, C.
Frissell, and J. Karr. 1993. Island Press. 462 pp. ISBN 1-55963-275-5. Describes current and proposed laws and
regulations for protection of stream resources. Available from: Island Press, Box 7, Covelo, CA 95428. Tel. (800) 8281302.
Stream Ecology: Structure and Function of Running Waters. J.D. Allen. 1995. Chapman & Hall, New York, NY.
388 pp. ISBN 0-412-35530-2. Technical description of the scientific concepts guiding research in stream ecosystems.
Available from: Patricia Ledlie Bookseller, Inc., Buckfield, Maine 04220. Tel. or FAX (207) 336-2778.
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Stream Hydrology: An Introduction for Ecologists. N.D. Gordon, T.A. McMahon, and B.L. Finlayson. 1992. John
Wiley & Sons, New York, NY. 526 pp. ISBN: 0-471-93084-9. Technical book on engineering hydrology, fluvial
geomorphology, and hydraulics with examples of their biological implications. Appropriate for students and researchers
with some training in ecology or engineering. Available from: Patricia Ledlie Bookseller, Inc., Buckfield, Maine 04220.
Tel. or FAX (207) 336-2778.
Managing Habitats for Conservation. W.J. Sutherland and D.A. Hill. 1995. Cambridge University Press, New
York, NY. 399 pp. ISBN 0-521-44776-3. Valuable reference to options and solutions for managing wildlife habitat
in different kinds of ecosystems. Useful for land managers, landscape architects, and conservationists. Available from:
Patricia Ledlie Bookseller, Inc., Buckfield, Maine 04220. Tel. or FAX (207) 336-2778.
Riparian Landscapes. G.P. Malanson. 1993. Cambridge University Press, New York, NY. 296 pp. ISBN 0-521-384311. Valuable reference on options and solutions for managing riparian habitat. Useful for land managers, landscape
architects, and conservationists. Available from: Patricia Ledlie Bookseller, Inc., Buckfield, Maine 04220. Tel. or FAX
(207) 336-2778.
IDENTIFICATION KEYS
Aquatic Entomology: The Fishermen’s and Ecologists’ Illustrated Guide to Insects and Their Relatives. W.P.
McCafferty. Jones and Bartlett. 448 pp. ISBN 0-86720-017-0. $55. Illustrated keys to the majority of the aquatic
insect species that are found in Indiana and useful ecological information on each species. McCafferty is a professor at
Purdue University, so many of the species are common to the midwest. Available from: Jones and Bartlett Publishers,
Inc., 20 Park Plaza, Boston, MA 02116. Tel. (800) 832-0034.
The Fishes of Ohio.
A Guide to Common Freshwater Invertebrates of North America. J. Reese Voshell. ISBN 0-939923-87. A brand
new volumeabout macroinvertebrates! This book is smaller in size (not information) and price than many others.
Available from: McDonald and Woodward Publishing.
Freshwater Invertebrates of the United States. R.W. Pennak. 1989. 628 pp. A very thorough volume with keys to
species or genus of each group. $45. Available from: Reiter’s Scientific & Professional Books, 2021 K Street, NW,
Washington, DC 20006. Tel. (202) 223-3327.
Aquatic Plant Identification Deck. University of Florida. Sixty-seven laminated “cards” riveted together with a clear
photograph of the aquatic plant on the front and general information about identification and habitat of the species on
the back. Some tropical species are not relevant, but most of the species are found in Indiana. $8. Available from:
University of Florida, Tel. (904) 392-1799 or (904) 392-1764.
A Manual of Aquatic Plants. N.C. Fassett. 1957. University of Wisconsin Press, Madison, WI. 405 pp. ISBN0299-01450-9. Well-illustrated technical key to aquatic plants in the Midwest region. Available from: Patricia Ledlie
Bookseller, Inc., Buckfield, Maine 04220. Tel or FAX (207)336-2778.
How to Know the Freshwater Algae. G.W. Prescott. 293 pp. ISBN 0-697-04754-7. $20. and How to Know the
Aquatic Plants. G.W. Prescott. 158 pp. ISBN 0-697-04775-X (spiral bound) or 0-697-04774-1 (cloth). $20. Illustrated
keys to the majority of the algae and aquatic plant species in the US. Order both from larger book stores.
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Newcomb’s Wildflower Guide. L. Newcomb. 1977. Little, Brown, and Company, Boston, MA. 490 pp. ISBN: 0316-60442-9. One of the easiest and best organized illustrated keys to use for identificationof wildflowers, flowering
shrubs, and vines. Available from: Patricia Ledlie Bookseller, Inc., Buckfield, Maine 04220. Tel. or FAX (207) 3362778.
Field Guide to Freshwater Mussels of the Midwest. K.S. Cummings and C.A. Mayer. 1992. Illinois Natural History
Survey. 194 pp. ISBN: 1-882932-00-5. Illustrated keys to the majority of the mussel species in the Midwest. Includes
color photos of all species. A booklet of color pictures based on this book is also available from U.S. Fish and Wildlife
Service in Ft. Snelling (Minneapolis), Minnesota. Available from: Illinois Natural History Survey, Natural Resources
Building, 607 East Peabody Drive, Champaign, Illinois 61820.
LAKE MANAGEMENT RESOURCES
The Waterfront Property Owner’s Guide. 1979. 58 pp. Information on how water quality problems are caused
and provides suggestions on actions that the individual property owner can take to protect and improve water quality
conditions in lakes and streams. Available from: The East Central Florida Regional Planning Council, 1011 Wymore
Road, Suite 105, Winter Park, FL 32789, Tel. (305) 645-3339.
Life on the Edge...Owning Waterfront Property. 1994. 95 pp. Gives advice on selecting waterfront property and
stewardship responsibilities of shoreline property owners in protection of water quality, open space, and natural beauty.
Sections on aquatic plants and federal, state (Wisconsin), and local laws pertaining to waterfront property. Available
from: University of Wisconsin-Extension, Lake Management Specialists, College of Natural Resources, University of
Wisconsin, Stevens Point, WI 54481, Tel. (715) 346-2116.
A Primer on Limnology. Second Edition. 1992. B.A. Monson. 54 pp. Introduction to physical, biological, and
chemical structure of lakes, lake classification, human influences, and process for organizing a lake study. Available
from: Water Resources Research Center, College of Natural Resources, University of Minnesota, Room 302, 1518
Cleveland Avenue, N., St. Paul, MN 55108.
Lake Smarts: The First Lake Maintenance Handbook. 1993. S. McComas. 215 pp. Guide to affordable projects to
help clean up, improve, and maintain lakes and ponds, including aquatic plant control, sediment, on-site waste disposal,
undesirable fish, and waterfowl management. Developed for Midwestern states. $18.95 paperback. Available from:
Terrene Institute, 1717K Street, N.W., Suite 801, Washington, D.C. 20006-1504, Tel. (202) 833-8317.
Our Nation’s Lakes: A National Catalog of Lake Information Materials. Wisconsin Lake Management Program,
1990, 37 pp. Listing of information available from natural resource agencies across the country. Pub. WR-242-90.
Available from: Wisconsin Department of Natural Resources, Bureau of Water Resources Management, P.O. Box 7921,
Madison, WI 53707, Tel. (608) 267-3579.
Monitoring Lake and Reservoir Restoration. R.E. Wedepohl, et al., EPA 440/4-90-007, 1990, and “The Lake and
Reservoir Restoration Guidance Manual” by H. Olem and G. Flock (eds), EPA 440/4-90-006, 1990. Developed to help
users identify, describe, and define lake problems, evaluate available management practices, develop and implement a
site-specific management plan. Available from: Clean Lakes Program, Assessment and Watershed Protection Division
(WH-553), U.S.Environmental Protection Agency, 401 M Street, S.W., Washington, D.C. 20460.
Volunteer Lake Monitoring: A Methods Manual. EPA 440/4-91-002, 1991, 124 pp., and “National Directory of
Volunteer Environmental Monitoring Programs” EPA 841-B-94-001, 1994, 531 pp. Methods for monitoring lake
conditions, including algae, aquatic plants, dissolved oxygen, and other characteristics. Listing of groups involved in
water quality monitoring. Available from: Clean Lakes Program, Assessment and Watershed Protection Division
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(WH-553), U.S. Environmental Protection Agency, 401 M Street, S.W., Washington, D.C. 20460.
Lake Leaders Handbook. 1995. Contains the “largest collection of specialized lake management information for
citizen leaders ever brought together in one document” to support responsibilities of lake organization leadership,
including topics on formation and operation of lake associations, motivation of volunteers, planning for the lake future,
understanding government (!), running a proper meeting, insurance coverage, grants, educational programs, lake
management, land use regulations, directory of lake managers, and publications list. Available from: UWEX-LAKES
PROGRAM, ATT: Robert Korth, College of Natural Resources, University of Wisconsin, 2100 Main Street, Stevens
Point, WI 54481-3897.
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Appendix D - Watershed Group Contacts
Friends of the Great Miami River
515 Wyoming Ave
Cincinnati, OH 45215
(513) 761-4003
Middle Great Miami Watershed Alliance
201 W. Main Street
Troy, Ohio 45373
(937) 440-3945
Honey Creek Watershed Project
4191 River Ridge Road
Dayton, OH 45415
(937) 898-3495
Stillwater Watershed Project
1117 South Towne Court
Greenville, OH
45331
(937) 548-1752
Indian Lake Watershed Project
324 Co. Rd. 11
Bellefontaine, OH 43311
(937) 593-2946
Three Valley Conservation Trust
5920 Morning Sun Rd.
Oxford, OH 45056
(513) 524-2150
Loramie Valley Alliance
822 Fair Rd.
Sidney, OH 45365
(937) 492-4768
Upper Great Miami Watershed Protection
Project
P.O. Box 3
Port Jefferson, OH 45360
(937) 456-8143
Lower Mad River Protection Project
4400 Gateway Blvd.
Springfield, OH
45502
(937) 328-4600
Mad River Steering Committee
1512 S. US Hwy 68, Suite B100
Urbana, OH 43078
(937) 484-1526
Watershed Enhancement Program
c/o Miami Valley Regional Planning Commission 40 W. 4th Centre, Suite 400
Dayton, OH 45402
(937) 223-6323
Wolf Creek Project
10025 Amity Rd.
Brookville, OH 45309
(937) 854-7645
D-1
D-2
Appendix E - References
Blair, Jane, ed., 1996. Student Watershed Research Project: A Manual of Field and Lab Procedures.
Third Edition. Saturday Academy. Oregon Graduate Institute of Science and Technology.
Cruz, Javier, 2000. Streamwalk Training Manual. Thames River Basin Partnership Initiative.
Earth Force/GREEN, 2000. Water Studies for Younger Folks: A Water Activities Manual for
Elementary School Students. (Based on the Field Manual for Water Quality Monitoring. Mitchell,
M. and W. Stapp.)
Farthing, Patty, et.al., 1989. The Stream Scene- Watersheds, Wildlife,and People. Oregon Department
of Fish and Wildlife, Watershed Education Project.
Faulds, Ann M. et.al., Macroinvertebrate ID Flash Cards. Earth Force - GREEN.
Global Learning and Observations to Benefit the Environment (GLOBE), 1997. Teacher's Guide.
GREEN/LaMotte, 1999. Water Monitoring Kit Manual. Code 5848.
Karr, James R., et.al. 1986. Assessing Biological Integrity in Running Waters, A Method and its
Rationale. Illinois Natural History Survey Special Publication 5.
Kopec, J. and S. Lewis, 1989. Stream Monitoring: A Citizen Action Program. Ohio Department of
Natural Resources, Division of Natural Areas and Preserves; Scenic Rivers Program.
Larson, Meg, 2000. Teacher Training Manual. Clinton River Watershed Council.
Loren, et. al., Save Our Streams Monitor's Guide to Aquatic Macroinvertebrates. Izaak Walton
League of America.
McCafferty, P.W., 1981. Aquatic Entomology: The Fishermen's and Ecologist's Guide to Insects and
their Relatives. Jones and Bartlett Publishers, Inc. Massachusetts.
Merritt, R.W. and K.W. Cummins, 1984. An Introduction to the Aquatic Insects of North America,
Second Edition. Kendell/Hunt, Dubuque.
Minnesota Pollution Control Agency, Spring 2000. Stream Reader: Newsletter of the Minnesota
Pollution Control Agency. Citizen Stream-Monitoring Program.
Mitchell, M. and W. Stapp, 1997. Field Manual for Water Quality Monitoring. Kendall-Hunt
Publishers, Iowa.
Murdoch, T. and M. Cheo, 2001. Streamkeeper's Field Guide. The Adopt-a-Stream Foundation,
Everett, Washington.
Natural Resource Conservation Service, 1996. Water Quality Indicators Guide - Surface Waters.
Kendall-Hunt Publishers, Iowa.
E-1
Ohio Environmental Protection Agency: Division of Surface Water, 1996. Biological and Water
Quality Study of the Middle and Lower Great Miami River and Selected Tributaries, Columbus,
Ohio.
Ohio Environmental Protection Agency: Division of Surface Water, 1995. Biological and Water
Quality Study of the Upper Great Miami River and Selected Tributaries, Columbus, Ohio.
Rand, G. and A. Petrocelli, 1985. Fundamentals of Aquatic Toxicology. Hemisphere Publishing
Corp., New York.
Stapp, W. and M. Mitchell, 1997. Field Manual for Global Low-Cost Water Quality Monitoring.
Kendall-Hunt Publishers, Iowa.
U.S. Environmental Protection Agency: Office of Water, 1995. Volunteer Stream Monitoring: A
Methods Manual, Washington, D.C.
E-2

Entire Stream Team Water Quality Monitoring Manual

Transcription

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