Laboratory Handbook - Section 14 - MAINLY BIOLOGY, A - J

Transcription

14
MAINLY BIOLOGY, A - J
CONTENTS of this section:
14.1
14.1.1
14.1.2
14.1.3
14.1.4
14.1.5
14.1.6
14.1.7
14.1.8
14.2
14.3
14.3.1
14.3.2
14.3.3
14.3.4
14.3.5
14.4
14.4.1
14.4.2
14.4.3
14.4.4
14.4.5
14.5
14.5.1
14.5.2
14.5.3
14.5.4
14.6
14.7
14.7.1
14.7.2
14.7.3
14.7.4
14.8
14.9
14.9.1
14.9.2
14.10
14.11
14.11.1
14.11.2
14.12
14.12.1
14.12.2
14.12.3
14.13
14.13.1
14.13.2
14.14
14.14.1
14.14.2
14.14.3
14.14.4
14.14.4
2006
Page
Animals in school
1402
Choice of suitable species
1402
Bringing pets and other animals into schools
1408
Animal supply
1412
Keeping and handling animals
1413
Feeding and cleaning
1415
Hazards to handlers
1416
Health of animals
1417
Anaesthesia and euthanasia
1417
Animals in the wild
1419
Aquaria
1420
Cold-water aquaria
1421
Fresh-water tropical aquaria
1428
Marine aquaria
1429
Accessories and electrical safety
1430
Maintenance of accessories
1434
Body fluids and cells - human
1438
Blood
1438
Cheek cells
1441D
Saliva
1441E
Urine
1441F
Sweat
1441G
Breathing investigations
1441G
Spirometers
1441H
Alternatives to spirometers
1441K
Breathing rate measurements
1441L
Manometers - pressure gauges
1441M
Disposal
1441M
Dissection
1441N
Whole-animal dissection
1441N
Dissection of material from butchers etc
1441O
Preserved material for dissection
1441P
General precautions with dissections
1441Q
Enzymes
1441Q
Fermenters
1443
Safety
1444
Practical considerations
1447
Genetic engineering
1451
Greenhouses
1451
Automatic watering systems
1453
Automatic ventilation
1455
Habitat creation
1456
Planning the development of a new habitat
1457
Creating habitats
1458
Management of a new habitat
1459
Hygiene
1459
Good practice
1460
Human blood
1461
Incubators & other temperature-stabilised equipment
1462
Controlling the environment
1462
Incubators
1463
Other equipment for maintaining stable temperatures 1464
D-i-y systems
1465
Miscellaneous tips
1468
Mainly biology, A - J
1402
14.1
Animals in school
14.1.1
Choice of suitable species
© CLEAPSS 2006
General principles
There should be definite educational reasons for keeping animals. At the present time,
some people regard the keeping of any animals in the laboratory or prep room as
undesirable, though such feelings often only relate to vertebrates. It is important,
therefore, that the animals’ presence can be justified and that they are involved in
investigational studies or for teaching about animal care rather than just ignored or
employed as ‘decoration’. In addition, animals should be treated in ways that are sensitive to their needs and be seen by pupils to be treated humanely.
It must be remembered that keeping animals properly requires time and commitment
from hard-pressed members of staff. To decide to keep animals imposes a responsibility not to be accepted lightly; it must be clear who is to look after the animals and
there must be others prepared to take on the tasks of animal maintenance if the main
person responsible is absent. It must also be noted that it is illegal to release any
non-native species into the wild.
Nevertheless, science departments which rarely, or never, use living animals in teaching science cannot justifiably claim that they are providing students with a complete
scientific education; after all, biology is the study of life and living organisms. Keeping
and studying living animals, both invertebrates & vertebrates, provides opportunities
for interesting, motivational work. Involvement with animals is also important in
helping students to develop caring and responsible attitudes towards animals and to
provide guidance on the needs of pets kept at home. It is much more difficult to fulfil
this obligation if no or few living animals are encountered in school.
Bringing animals into school for a short period, rather then keeping them permanently, can be extremely valuable but requires careful consideration. This issue is discussed in detail in section 14.1.2.
Educational
purpose
Animals that serve a variety of purposes are better educational value. The inexperienced should, however, start with less-demanding species. Numbers of animals
should be kept low until experience is gained. Invertebrates will usually be less
demanding in their maintenance than vertebrates but provide many opportunities
for investigations.
Health hazard
Few animals present a health hazard; obtaining them from reputable sources,
providing proper care and practising good hygiene will, in most cases, eliminate
hazards or minimise them to insignificant levels. Avoid animals that carry a
higher health risk; see section 14.1.6 and the list at the end of this section (14.1.1).
Once on the premises, it is vital that small mammals cannot come into contact
with wild rodents etc; otherwise there is the possibility of disease being transmitted to healthy stocks.
Housing
Some animals need specialised housing which may not be available, affordable by
schools or easy to site satisfactorily. Use the largest cage, tank etc possible and
pay particular attention to creating an appropriate and aesthetic environment for
the animals and their display. Although in many cases, animals are actually best
maintained in sparse housing which is therefore easy to keep clean, strict adherence to such regimes may be counter-productive in a school context. Pupils may
view such environments as ‘unnatural’ or even ‘cruel’ to the animals.
Further information on housing requirements for a wide range of vertebrates and
invertebrates is given in CLEAPSS Guide L56, Housing and Keeping Animals.
© CLEAPSS 2006
Feeding
1403
Long weekends and holidays can present problems if the animals need specialist
care. If, at these times, animals are looked after by people other than the normal
person responsible, ensure that the volunteers are fully instructed in the animals’
normal maintenance and in appropriate action for emergencies. Carnivores are
generally more of a problem than herbivores. Pupils usually accept animals eating
worms and maggots etc but they may be upset to think of snakes eating small
vertebrates such as chicks, mice etc. There is also the practical problem of maintaining a supply of live or recently-killed food, though much of the latter can be
stored in the freezer until needed. Problems are largely overcome if the animal
will eat tinned dog food, or meat from the butcher.
Some species suitable for keeping in school are discussed below, from which a choice
can be made to form the nucleus of a useful school collection. The list is not intended
to be complete in any sense; much depends on the knowledge and experience of school
staff. Further information is given in separate CLEAPSS Guides1 and in the books
detailed in Table 14.1. Out-of-print titles are often still available for purchase on the
Internet, for example, from Amazon, E-Bay and other on-line sources.
Table 14.1
Further information on animals
Title
Author
Date
ISBN
Publisher
Breeding the British Butterflies
Rearing and Studying Stick & Leaf Insects
Rearing Crickets in the Classroom
Silkmoth Rearer’s Handbook
Peter Cribb
Paul Brock
Brian Gardiner
Brian Gardiner
2001
2003
1981
1982
0900054662
0900054689
0900054638
0900054395
Amateur
Entomologists’ Society
Tarantula Keeper’s Guide
S and M Schultz
1998
0764100769
Barron’s Educational
Handbook on the Care and Management of
Laboratory Animals:
Vol 1 Terrestrial Vertebrates
Vol 2 Amphibious & Aquatic Vertebrates &
Advanced Invertebrates
UFAW
Gardening for Butterflies
Margaret Vickery
African Clawed Toad: a Guide to the Biology,
Care and Breeding of Xenopus laevis
Blackwell Publishing
1999
0632051310
1999
0632051329
1998
-
Butterfly Conservation
A M Leadley Brown
1970
Out of print
Butterworth
Care of Reptiles and Amphibians in Captivity
Garter Snakes: Care in Captivity
Keeping and Breeding Snakes
Chris Mattison
Roger Sweeney
Chris Mattison
1987
1992
1998
Out of print
Out of print
0713727098
Cassell / Blandford
Garden for Birds
Starting a Butterfly Garden
Nigel Matthews
Peter Cawdell
1992
1987
1851168052
185116801X
Chalksoft / SGC Books
Keeping and Breeding Tarantulas
Rearing Stick and Leaf Insects
Rearing Wild Silkmoths
Ronald Baxter
Ronald Baxter
Ronald Baxter
1993
2002
1992
0951921924
0951921932
0951921908
Chudleigh Publishing
Keeping Spiders, Insects and Other Land
Invertebrates in Captivity
Frances Murphy
2000
0952408325
Fitzgerald Publishing
Animals in Schools Vol 1: Vertebrates
Vol 2: Terrestrial Invertebrates
M Hogg
L Comber & M Hogg
1977
1979
Out of print
Out of print
Heinemann
Keeping Stick Insects
Dorothy Floyd
1987
0951246607
Small-Life Supplies
Birdkeeper’s Guide to Finches
Complete Encyclopedia of the Freshwater
Aquarium
Interpet Guide to Coldwater Fishes
Interpet Guide to the Tropical Aquarium
Pet Owner's Guide to the Leopard Gecko
Q & A Manual of Reptiles & Amphibians
David Alderton
1999
1902389867
Interpet Publishing
John Dawes
Dick Mills
Dick Mills
Noel Morgan
R & V Davies
2001
1999
1999
2003
1997
1842860410
1902389530
1902389514
1860541240
190238993X
1
L52 Small Mammals, L56 Housing and Keeping Animals, L71 Incubating and Hatching Eggs, L197 Giant African Land Snails,
L201 Giant Millipedes, L206 Tadpoles, L213 Science with Minibeasts: Snails and L227 Stick Insects.
1404
© CLEAPSS 2006
Vertebrates
Fish
‘Tropical’ - guppies; swordtails; mollies; platties; gouramis; cichlids; angel fish;
acaras; barbs; tetras and a variety of other species. ‘Cold water’ - goldfish: the
cheaper kinds are likely to be hardier and more suitable as those with large tails
and bulbous eyes require considerably more care. Koi are much more expensive
and less tolerant of an unsuitable environment. Short term: sticklebacks and
other small fresh water species; these require very clean conditions. Rearing trout
from eggs presents problems of feeding, aeration, disease etc but can be an interesting exercise. Adult trout are not suitable. For details of aquaria, see section
14.3 (Aquaria), CLEAPSS Guide L56 and references in Table 14.1.
Amphibians
Clawed toads1, Xenopus laevis or borealis; Axolotl, Ambystoma mexicanum; Tiger
salamander, Ambystoma tigrinum; Fire salamander, Salamandra salamandra.
Observing the growth and development of tadpoles of the common frog or toad
often occurs and is recommended, provided that only a small quantity of spawn is
taken from the wild, the tadpoles are reared as carefully as possible and the
adults returned to their native pond if possible or another suitable site. [Taking
spawn of common species is not illegal; see section 14.2 (Animals in the Wild).]
Adults of native common frogs and toads do not usually thrive in captivity indoors
and should not normally be kept other than in a pond outdoors. If animals are
brought into school by children, they can be maintained for a short time in temporary housing. Experience has shown that the adults of frogs from North America,
particularly Rana pipiens and clamitans which are sometimes available, often
adapt well to the conditions of an indoor vivarium. Tadpoles of the North American bullfrog, Rana catesbiana, are sometimes purchased and may be attractive
because of their larger size and relatively slow growth. It should be noted, however, that adult bullfrogs - which the tadpoles will inevitably become - are large
animals and not easy to rear and feed.
Adult amphibians are carnivorous and often require moving, live food such as
maggots, mealworms, worms or crickets. Some, such as adult North American
bullfrogs, require vertebrate foods and are obviously less suitable. Clawed toads
can be fed an artificial pelleted diet and these animals, together with axolotls,
may readily eat small pieces of liver, heart etc from the butcher.
Reptiles
Reptiles are not the easiest animals to keep, requiring heated vivaria often with a
‘basking’ area where the animals can thermoregulate by moving in and out of the
heat given off by a lamp. Some have particular requirements with regards lighting; a source of ultra-violet radiation is essential for the health of many species.
Feeding may be a problem with some animals requiring vertebrates as food.
Nevertheless, there are several reptiles that are relatively easy to maintain; authoritative advice should be sought before keeping other species.
Despite the often-quoted statement that all lizards are difficult to rear, there is
one, the leopard gecko, Eublepharus macularius, which presents relatively few
problems. It does not need U-V radiation, and will feed on any dry invertebrates,
including crickets, mealworms, maggots, or animals from the garden such as
woodlice and spiders. It conveniently always defaecates in the same place, so can
easily be persuaded to use a dish for ease of cleaning! Food should be dusted with
a vitamin supplement and crushed cuttlefish ‘bone’ also provided. Males should
not be housed together but pairs readily breed. As an additional bonus, the leopard gecko cannot climb the sides of its tank and so escapes only rarely. Skinks are
also relatively easy to keep but are less suitable since they hide for much of the
time.
Secure caging is essential for keeping any snake. The common garter snake,
Thamnophis sirtalis, is relatively easy to keep, provided that it is given the correct
food. It can be offered earthworms occasionally but its main diet should be strips
of selected fish including trout, plaice or ‘lancefish’ (the latter obtainable frozen
from pet shops). Include the skin and bones when feeding the snakes. (Other types
1
See reference in Table 14.1 or Revised Nuffield Biology Teachers’ Guide 1, Nuffield Foundation, Longman, 1974, Appendix 3.
© CLEAPSS 2006
1405
of fish contain an enzyme that accumulates and ultimately kills the snake. It can
be destroyed by thoroughly boiling the fish but this then is often unpalatable to
the snake.) The corn snake, Pantherophis guttatus (Elaphe guttata), and the rat
snake, Pantherophis obsoletus (Elaphe obsoleta), are also undemanding animals
but they do require a rodent diet of mice and small rats which can be obtained
frozen from pet shops.
Terrapins are sometimes considered but are not normally suitable for the inexperienced as they are not the easiest animals to maintain, grow to a large size and
have been reputed to be carriers of Salmonella. Good hygiene, however, will
resolve issues of infection if these animals are kept. Tortoises, also reputedly
Salmonella carriers, are now very expensive and not often readily available.
Birds
Wild birds should never be caught or brought into school. Of the pet species bred
in captivity, members of the parrot family may be susceptible to psittacosis, a
disease that is rare but transmissible to humans. Free-flying flocks of budgerigars, pigeons, doves or other birds in aviaries may come into contact with wild
birds and so present similar risks from ornithosis. As a result it is difficult to
ensure that any captive-bred bird is completely free from disease though the
hazard must not be exaggerated.
It is usually preferable to observe birds in the wild (or school grounds) than to
confine them in cages but, if a school really wants to keep birds, the small seedeating species such as finches might be considered. Ensure, however, that the
distance between the wires in a cage is not too large; small finches can escape
easily! Budgerigars, obtained from a reliable source, may also be appropriate. It
may be possible for a local veterinary surgeon to check the health of a bird and so
identify if there might be risks from ornithosis. Much help in keeping birds can be
obtained from a local cage bird society.
If proper conditions, including an appropriate incubator and brooder, can be provided and maintained, schools might wish to hatch chicks or keep poultry including
ducks. See Table 14.1 and the CLEAPSS Guide L71, Incubating and Hatching
Eggs. Some schools have successfully kept quails. Before commencing work on
incubating eggs, it will be necessary to have found homes for the adult birds, if
these are not to be reared at school.
Mammals
Mouse, Mus musculus; rat, Rattus norvegicus; golden or Syrian hamster, Mesocricetus auratus; Russian hamster, Phodopus sungorus; Mongolian gerbil or jird,
Meriones unguiculatus (other species such as the Libyan or pallid jirds may be
available and are equally suitable); cavy or guinea pig, Cavia porcellus and rabbit,
Oryctolagus cuniculus. The choice of small mammal(s) to keep will be determined
by a variety of factors, as listed below. Further details are in CLEAPSS Guide
L52, Small Mammals.
Will the mammals be handled regularly by pupils? (Rats, guinea pigs and rabbits
are best.)
Are low maintenance times and costs of paramount importance? (Gerbils score
here.)
Is an active animal with interesting behaviour required? (Again gerbils are most
suitable, while hamsters are less desirable as they are nocturnal.)
Is there limited space or size of caging available? (Guinea pigs and rabbits are less
suitable for keeping indoors since they need a lot of room and should ideally have
access to outdoor pens.)
Are the mammals needed for genetic breeding investigations? (Mice in particular
and some strains of gerbil have advantages here.)
Farm animals
These are often kept by schools that teach rural studies and therefore require
special facilities and staff with specialist knowledge.
For further information on vertebrates, refer to the UFAW Handbook on the Care and
Management of Laboratory Animals and other publications in Table 14.1.
1406
© CLEAPSS 2006
Invertebrates
Useful examples are too numerous for them all to be listed here. In addition to fruit
flies (Drosophila melanogaster) and flour beetles (Tribolium sp.) for genetics, a wide
range of insects and other invertebrates, many collected from the wild for short-term
studies or maintained as laboratory cultures, can be used in behavioural, physiological
and reproductive studies. See Animals in Schools Vol 2 Terrestrial Invertebrates cited
in Table 14.1. For more-detailed information on keeping a variety of invertebrates,
refer to CLEAPSS Guide L56, Housing & Keeping Animals.
Butterflies and
moths
It is difficult to house adult British butterflies and moths over an extended period
unless a large net-covered outside area can be created in which suitable plants
can be grown. For studies of life cycles, it is, however, possible to rear the eggs,
larvae and pupae indoors and then release the adults of native species in an
appropriate place where they would normally be found. For a particular species,
supplies of the appropriate food plant for the larvae to eat will be needed. For the
large white butterfly, Pieris brassicae, and painted lady butterfly, Vanessa cardui,
artificial diet is available and the animals could be reared throughout the year.
The eggs and larvae of tropical silk moths (eg, the Chinese oak silk moth Antheraea pernyi) are often available, easy to keep indoors and the adults will mate
without needing a large cage. Where pupils bring larvae into school, it is important for them also to note on which plant they were found, so that the animals can
be provided with appropriate food. The larvae and food plant should ideally be
enclosed in a nylon net cage to give adequate ventilation, but a ‘cylinder’ cage
constructed from a sheet of acetate and a biscuit tin may be suitable, if there is
limited condensation. Provide twigs and a layer of slightly moist peat for pupation
of the larvae; depending on the species they will climb or burrow before they
pupate.
Rather than containing butterflies, it may be preferable to attract them to the
school garden. Table 14.1 provides references for keeping lepidoptera indoors or
encouraging them into the garden. Visits to special living collections such as the
various Butterfly Houses around the country are exciting and informative.
Suppliers of eggs and larvae of butterflies and moths, such as Worldwide Butterflies, are an invaluable source of advice.
Stick insects,
locusts
These are herbivorous and so, in theory, are easy to keep1. Fresh grass and bran
will be needed for locusts and privet, bramble or other plants for stick insects,
depending on the species. A source of heat is necessary in, or near, locust housing
and, if locusts are to be bred, sand containers must be provided for the eggs.
However, keeping locusts in continuous culture is no longer recommended
because of the increased risk of allergies developing with long-term exposure to
the locust allergens. Small locust hoppers and stick insects are great escape
artists. Some species of stick insects have sharp spines or emit defensive chemicals; these should be handled with care.
It is fairly easy to improvise temporary housing for stick insects brought in by
pupils. Plastic sweet jars or soft-drinks bottles make suitable housing for some
stick insects. For animals kept over a longer period, greater air circulation is
needed and a cage with ventilation on at least two sides is required.
Bees
These need special housing and care by someone with specialist knowledge; they
can, however, create considerable interest. A local apiary society may be helpful2.
1
For rearing locusts see CLEAPSS Guide L56, Housing and Keeping Animals. CLEAPSS Guide L227, Stick Insects, provides
detailed information for rearing these animals.
2
Contact the British Beekeepers Association [see Section 1.1 (Addresses)] for details of local societies.
© CLEAPSS 2006
1407
Brine shrimps
and water fleas
Brine shrimps are easily hatched from eggs in salt water and kept in soft-drinks
bottles or aquaria. The animals provide excellent material for various studies and
there is good support material produced by the Brine Shrimp Project at Homerton
College. Water fleas (Daphnia), are usually readily available from aquarists and
again are useful for a variety of studies, particularly investigations on heart rate.
For details of keeping both types of crustacean, refer to CLEAPSS Guide L56,.
Housing and Keeping Animals.
Giant African
land snails
In the past, some imported from the Far East have been found to carry parasites
potentially harmful to humans. Animals available in the UK will, however, have
been bred in this country and are not hazardous since the parasite’s life cycle will
have been broken and the snails cannot become reinfected. Ensure that animals
have not been recently imported. See CLEAPSS Guide L197 for detailed information.
Giant millipedes
These tropical herbivores provide an interesting example of the myriapod group of
animals. See CLEAPSS Guide L201 for detailed information.
Cockroaches and British and American species of cockroaches are great escape artists and may
cause an infestation of school premises, making the science department deeply
crickets
unpopular! Species of cockroaches from tropical areas are, however, unlikely to
escape or survive to breed if they do. Crickets are a useful source of food for
reptiles and amphibians, as well as being interesting animals to study. They are
readily available from suppliers of food for pet animals but can also be maintained
as cultures in schools. For details of maintaining cockroaches and crickets, refer to
CLEAPSS Guide L56, Housing and Keeping Animals.
Other
invertebrates
Garden ‘minibeasts’ are often kept for short periods and returned to their source
as soon as possible, therefore only requiring temporary shelter. Where native
British animals are kept indoors for longer periods, it is important to remember
that they are not well adapted to a warm, dry atmosphere. They must be kept
moist and in a cool position, preferably in the dark. Where this is difficult to
achieve indoors, their containers should be placed in a shaded position outside.
Woodlice and garden snails are tolerant of conditions indoors and can be maintained as cultures in schools; earthworms, however, cannot. Other than for shortterm observations of burrowing, earthworms should not be kept in a typical
wormery consisting of a sandwich of soil between two sheets of glass or Perspex.
They are best kept outdoors in a sturdy box of soil. Refer to CLEAPSS Guide L56
and also Table 14.1 for various references for keeping invertebrates.
Unsuitable animals
m The following animals are unsuitable for keeping in school either because it is illegal
to do so or they typically present unacceptable risks (for example, of injury or the
transmission of disease):
monkeys and apes;
parrots or parakeets;
crocodiles and alligators;
poisonous reptiles;
mammals and birds taken, even if legally, from the ‘wild’.
To this list should also be added all animals for which there is inadequate knowledge,
expertise and commitment for their proper care. The list of unsuitable animals will
therefore be much more extensive in some establishments than others. See also
section 14.2 (Animals in the wild) for reference to animals protected by the Wildlife
and Countryside Act.
14.1.2
1408
© CLEAPSS 2006
Bringing pets and other animals into schools
Whether or not animals are permanently kept in the science department, there are
occasions when animals are brought into school for a short period, possibly just for a
day. In these circumstances, staff may not have sufficient experience of keeping and
handling the animals concerned. For details of how to look after such visitors for more
than just a day or so, schools should refer to suitable specialist texts, including
CLEAPSS Guides where relevant (see the list of titles in the footnote on page 1403).
The purpose of the information in this section is to provide general guidance on coping
with short-term visitors.
Pets and farm animals
Children may want to take their own pets into school to show them to their friends or
they may be encouraged to bring them in because the animals fit in well with a
current teaching topic. (This may apply to staff members’ pets too!) This can be very
valuable educationally but it is important that suitable arrangements are made in
advance for the well being of the animals for the short time that they are on the
school premises. Some establishments organise an ‘animal club’ for keen pupils who
help to look after the school’s collection of animals. Club members then bring in their
own animals for short periods. Occasionally, a school may arrange an ‘animal day’ in
which the pupils in a whole class bring in their pets; these pose problems simply by
the sheer number and variety of visitors! In schools with links to the rural community,
it is quite common for certain farm animals, such as new-born lambs, chicks or ducks,
to be brought in for a short time.
Effective organisation
When animals are brought into school, they must be looked after as well as they
would be when they are at their normal residences. If this cannot be achieved,
they should not be brought in. Using the expertise of pet owners, especially for
exotic species, may provide the necessary guidance for the proper care of the
animals.
The procedure of bringing the animal to school should not be unduly stressful (for
the handler as well as the animal!). Thus it would not be a good idea to remove
fish from their normal aquarium and transport them in a polythene bag, empty
them into a new tank of chlorinated water at school and then repeat the upheaval
at the end of the day. Common sense should apply!
Where a variety of animals will be together, consider carefully the possibility of
unwanted interactions between certain individuals. This is a particular problem if
the pets will include cats and dogs; will they fight or show an unhealthy interest
in any fish or birds that are also in the room?
Housing and food Animals must be properly housed while they are on the school premises. Where an
animal requires special housing that is not easily transportable and cannot be
for the animals
provided by the school, it too should be left at home. It may, however, be possible
for a spare cage etc to be taken to school in advance so that the animal can be
transferred to familiar territory as soon as it arrives in a transportable carrier. If
animals cannot be brought into school in their normal housing, containers used to
transport them must be appropriate and retain the animal securely. The possibility of gerbils, snakes and spiders running loose on the school bus, or when they
reach school, must be prevented at all costs!
Animals must be housed separately; different species, or individuals of the same
species from different litters, must never be placed together in the same cage,
vivarium, tank etc. Apart from ensuring that the correct animal is returned to its
rightful owner, this will prevent diseases passing between individuals and stop
any aggressive conflict between incompatible animals.
© CLEAPSS 2006
1409
An animal is likely to require feeding and watering during its stay on the school
premises, although this may not be an issue for some animals, eg, snakes and
spiders, which feed irregularly. The school should ensure that the owner knows
that all necessary food and equipment must be brought in with the animal.
Providing a
suitable
environment for
the animals
The conditions in which the animals will be left while they are at school must also
be considered carefully. For example, could the various animals brought in be left
near windows and subjected to the stress of overheating by sunlight on a hot day?
The animals may be the focus of attention for only part of a day. If they are not to
remain in the room when other work is commenced (and therefore become a considerable distraction), can alternative, and appropriate, accommodation be found?
If the animals will be in a classroom or laboratory all day long, are there particular organisms that would be stressed by the noise and constant attention of a
large group of excited pupils?
Handling animals This will be one of the main attractions with animals brought into schools. It
usually generates a great deal of interest and excitement but handling sessions
need to be managed carefully so that they do not become out of control. It is likely
that small mammals that are pets will be accustomed to being handled and will
not be stressed by this. However, such animals will not be used to the attentions
of so many people at once and may react differently to handling in school than
they do at home. Children should be shown how to handle small mammals1 and
other animals properly; the technique that is right for one animal is not correct for
another. There is always a danger that animals may be dropped, particularly by
children with small hands and if the animal is agile or fast moving. Handling
small animals should therefore always be carried out over a table or a trough
filled with a soft material such as sand or sawdust, so that the animal will not fall
more than a very short distance and onto a suitable surface.
It should be appreciated that some animals are best not handled at all. Hamsters
that are nocturnal and asleep during the day will not necessarily appreciate being
disturbed. Any animals that appear to be nervous should just be observed and not
handled. Invertebrates, eg, stick insects and spiders, are delicate and fragile
animals, for these, handling should always be kept to a minimum, for example
when cleaning their cages. They could easily be damaged by over-keen children,
not used to handling such animals gently. It is not recommended that tarantula
spiders are handled; see Health & safety issues below. An animal may best be
handled only by its owner.
Keep control!
All of these considerations require careful organisation by the school, especially if
several animals are to be brought in on the same day. Staff should be aware of the
pet that a pupil intends to bring in and therefore the problems that it might pose.
Liaison with parents is clearly very important.
Health & safety issues
In many ways, these are of lesser importance than the need to ensure the well
being of the animals brought into school, although this will not be the immediate
perception of most people.
Hygiene
1
If animals will be handled, the most important issue is the maintenance of good
hygiene.
•
Children and adults should always wash their hands soon after handling any
animal (or coming into contact with the soil, bedding, water etc in an animal’s
housing) and this is best done in the classroom or laboratory, where it can be
seen to be done.
•
Cover cuts and abrasions on the exposed skin of hands and arms.
•
Suitable facilities and, usually, paper towels for drying hands hygienically
are needed.
The CLEAPSS guide L52, Small Mammals, discusses appropriate handling techniques for different species of small mammal.
Physical injuries
1410
© CLEAPSS 2006
There is always the danger of bites and scratches, and teachers or technicians
should check that animals to be brought in are docile, friendly and gentle in the
presence of (exuberant) children. This is particularly important with larger
animals such as cats and dogs that have the equipment to inflict a lot of damage!
Pupils should be warned of the hazards of some animals, eg, not tormenting a
sleeping hamster that might retaliate upon waking! Small fingers poked towards
the mouths of normally non-aggressive animals may be interpreted as an offering
of food and obligingly bitten. The bodies of some large spiders are covered with
many small, barbed hairs or bristles; these cause irritation when they penetrate
the skin, mucous membranes and especially the eyes. This is another reason for
pupils not to handle such spiders that are brought into schools, even if the (illinformed) owner indicates that handling the spider is not harmful. Do not touch
the face or eyes if contact is made with ‘hairy’ spiders, or material within their
housing, until the hands have been thoroughly washed. Some pugnacious spiders
reportedly shed a cloud of irritant bristles into the air but it is unlikely that
aggressive species will be kept as pets by most pupils.
Diseases,
parasites and
allergies
The likelihood of diseases being passed on to humans from pet animals is low.
Farm animals1, however, present a higher risk. In all cases, good hygiene will
reduce the risks even further. For cats and dogs, it is sensible to check that these
have been regularly taken to a vet and have been recently wormed and treated for
fleas. For larger animals, including cats and dogs, it is important to provide
appropriate toilet facilities! All wastes produced, whether accidental or routine,
must be handled and disposed of hygienically, and contaminated items and
surfaces properly washed and disinfected; see section 15.12.3.
Small mammals should not be allowed to come into contact with school stocks at
any time to guard against possible transfer of diseases to resident animals.
Allergic reactions to mammals, birds and a few other animals cannot be discounted. These might result from handling the animals or just from being near
them and be detected by the development of skin rashes, irritation to the eyes and
nose or breathing difficulties. Again, washing hands soon after handling animals
will help and it is important to stop children rubbing their eyes before this has
been done. Children known to have allergic reactions to specific animals must, of
course, have restricted access to those that may trigger a response. In most cases,
an allergic reaction will subside once the animal and the afflicted person are kept
apart; in extreme cases, seek medical advice.
Phobias
Schools should be aware that pupils and adults may have phobias relating to
certain animals, eg, snakes and spiders. This is not to say that such animals
should not be brought into schools; indeed they may even be valuable in helping
individuals to overcome their phobias. However, fears that may be expressed
should be respected and efforts made to segregate the animals from people with
such phobias.
Visiting animal schemes
There are commercial organisations or individuals that, for a fee, take a variety of
animals into schools. Some authorities have misgivings about such schemes; fearing
that the animals in the collection, although not deliberately ill-treated, may be
harmed or stressed by the regular movement from place to place and repeated
exposure to, or handling by, groups of enthusiastic pupils. Some schemes ensure that
they have several individuals for each species in the collection. In this way, when one
animal is being shown to children in schools, its companions are ‘resting’. Before
deciding to invite in a visiting animal organiser, it might be worth enquiring whether
such a system is in operation.
1
School visits to farms etc are not discussed here. There is, however, guidance on precautions to take issued by the HSE and
DfEE. If schools require copies of this guidance, they should contact the CLEAPSS Helpline.
© CLEAPSS 2006
1411
It is essential to obtain information about the animals that will be shown, to identify if
there are likely to be any problems, such as phobias to particular species or allergic
reactions. The discussion on handling animals and health & safety issues in the
previous section is also likely to be relevant here, and good hygiene will be crucial.
Various conservation trusts, in promoting their organisations or publicising their
work, may bring ‘rescued’ wild animals into schools. Make sure that such animals, eg,
hedgehogs or owls, have been kept in captivity for some time and that they have been
treated for diseases transmissible to humans (if applicable) and parasites, such as
fleas. It may not be appropriate that children handle these rescued animals.
Animals from local habitats
Land
invertebrates
Invertebrate animals, such as woodlice, snails and earthworms, are often brought
into schools for short-term studies. While kept indoors, ensure they are kept in
cool, damp and dark conditions, which simulate their natural surroundings. Note
that it is not easy to keep carnivorous invertebrates, such as centipedes or spiders,
for extended periods because of the difficulty of providing them with live food.
Aquatic animals
Fish, such as minnows, from local streams should be kept in pond water that is
cool and well aerated, and provided with a supply of live food. If kept for any
length of time, refer to specialist texts on cold-water fish for guidance.
If sampling animals from ponds, streams or rivers, there may be a possibility that
the water has been contaminated with the bacteria that cause Weil’s disease.
Good hygiene is essential. Separate information on Weil’s disease is available1.
Amphibians
In spring, the spawn of common species of frogs, toads and newts (ie, not the
great crested newt nor the natterjack toad) are often studied in schools. This is
acceptable and perfectly legal, but only small amounts of spawn should be taken
and the tadpoles are best reared2 in water that is aerated and filtered. In this
way, the maximum number of tadpoles will survive to become young adults for
release into a suitable environment (preferably the pond where the spawn was
taken from, if this is known).
Birds and
mammals
Because of health & safety concerns, wild birds and mammals should not normally be taken directly from the wild and brought into schools (see Animals to avoid
below). However, this recommendation does not stop pupils arriving with injured
animals that they have found. While such animals are on the school premises,
guard against the possible risk of disease and parasite transmission. Isolate the
injured animal from resident small mammals and birds and keep it in quiet,
dimly-lit conditions. Handle the animal as little as possible, with due regard for
personal health & safety. Wash hands immediately afterwards if it has not been
appropriate or possible to protect the hands by wearing suitable gloves.
Pupils may have unrealistic expectations of school staff; they often want the
injured animal to be nursed back to full health and released into the wild.
Unfortunately, this may not be possible without expert help and the most appropriate action is often the immediate humane killing of the animal. Staff in schools
may not have the expertise, facilities or willingness to achieve this, and may not
even be able to assess the condition of the animal. At the earliest opportunity, to
avoid prolonging the animal’s suffering, staff will normally need to take the
injured animal to a local vet or an animal welfare clinic run by the PDSA or
RSPCA.
The first time that an injured animal arrives is not the occasion for frantic telephoning to find the help required. Schools are therefore advised, in anticipation of
the arrival of an animal in distress, to check out the local assistance available.
Vets are commercial businesses and should be asked if they will charge to treat
such wild animals or put them to sleep; it is hoped that they would not.
1
See section 17.1.3 or CLEAPSS guidance leaflet PS1, Pond dipping and Weil’s disease.
2
See CLEAPSS Guide L206, Tadpoles, for details.
1412
© CLEAPSS 2006
Try to dissuade pupils from bringing in young animals that they have found
‘abandoned’; the animals’ parents are often not far away and will generally
retrieve their offspring as soon as the unwelcome intruder has moved on!
Animals to avoid
It goes without saying that certain animals, such as leopards, monkeys, crocodiles
and poisonous snakes, should not be brought into schools! Indeed, such animals
are governed by the Dangerous Wild Animals Act; this makes it an offence to
possess such animals without a licence.
Endangered species should also not be brought into school, though it may be
difficult for school staff to identify which organisms are fully protected by the
Wildlife & Countryside Act which makes it illegal to take them from the wild.
Under the Act, there are different levels of protection given to different species.
Those with full protection (which includes, for example, dormice and all bats, all
wild birds and their eggs, the great crested newt and natterjack toad, the smooth
snake and sand lizard, and a variety of rare invertebrates) obviously must not be
taken from the wild. Lower levels of protection make it illegal to kill or injure an
animal and, for some species, it is only illegal to take them from the wild and sell
them without a licence. For such animals, it is legal to bring them into school,
though other considerations, discussed above, may rule against this. For details of
protected animals, use the web site of the Joint Nature Conservation Committee:
www.jncc.gov.uk; click on ‘legislation’ and ‘Wildlife & Countryside Act’.
Wild birds and mammals, dead or alive, are best not brought into schools directly
from the wild (even if legal to do so) because they may be harbouring diseases or
carrying parasites such as fleas, lice or worms that could be transmissible to
humans (or resident animals). They may also inflict physical injuries. See, however, the discussions above. Old birds’ nests could also be a source of parasites and
it is best to seal these in plastic bags. If the trapping of wild mammals is to be
studied, when identifying the animals caught they must be handled with care,
wearing gloves that provide protection from contamination and bites or scratches.
14.1.3
Animal supply
To obtain healthy animals, especially small mammals, a supplier with a good reputation should be used; this often excludes market stalls. Accreditation schemes for the
supply of certain small mammals that were recommended in the past are no longer
open to schools. If there is any doubt about the health of an animal, particularly small
mammals or birds, it should be taken to a vet before being brought into school.
General suppliers Blades Biological, Philip Harris Education and Sciento are examples of the few
companies that specialise in supplying schools which sell a range of invertebrates
and vertebrates, both living and preserved. It should be noted, however, that the
variety of animals available from such sources may decrease, particularly for vertebrates. Schools will therefore increasingly have to rely on local sources such as
pet shops or on specialist suppliers that sell to the general public. CLEAPSS
Guide L56, Housing and Keeping Animals, gives details of several of these suppliers and the types of animals that they sell.
Small mammals:
use pet shops
and breeders’
societies
None of the schools’ suppliers now sells live small mammals. Pet shops vary in the
quality of the animals they sell and the conditions in which they are kept. If such
sources are to be used, especially for small mammals, it is recommended that
schools should purchase only from shops that are evidently well run and clean,
keep their animals in first-class conditions and have knowledgeable & dedicated
staff. It may be possible to find reliable local sources of small mammals from
specialist breeders’ societies.
Fertile eggs of
domesticated
birds
CLEAPSS Guide L71, Incubating and Hatching Eggs, advises on suppliers of
fertile eggs of chickens, ducks and other birds. Some suppliers may be traced
through advertisements in the monthly magazine Cage and Aviary Birds.
© CLEAPSS 2006
1413
Reptiles and
amphibians
Blades Biological may offer a small range of amphibians and reptiles. In addition,
there are companies that specialise in the breeding and supply of these animals
and they can also be obtained locally from pet shops; see Yellow Pages. Animal
Allsorts (Reptile Centre), Coast to Coast Exotics and Peregrine Livefoods are
examples of sources; see CLEAPSS Guide L56 for further suppliers.
Fish
Fish and trout eggs can be obtained from local aquarists or fish farms; see Yellow
Pages under Aquarium and Pond Supplies and Fish Farms & Hatcheries and
advertisements in the monthly magazine Practical Fishkeeping.
Invertebrates
As well as schools’ suppliers, including Blades Biological, Philip Harris Education
and Sciento, which stock a range of invertebrates including Drosophila and
Tribolium, locusts, stick insects, snails and cockroaches, there are a number of
specialist suppliers of invertebrates for display, investigatory work or as a source
of animal food. Several of these are listed in Table 14.2; Guide L56 has a more
extensive listing. Maggots (blowfly larvae) and Daphnia (water fleas) can be
obtained locally from fishing tackle shops and aquarists.
Table 14.2
Suppliers of
invertebrates
Supplier
Organisms available
Bugs Direct UK
Livefoods Direct
A wide range including snails, beetles, cockroaches, millipedes,
tarantulas and stick insects.
Lepidoptera and some other invertebrates.
An extensive collection including stick insects, cockroaches, snails,
spiders and other species.
Many animals are supplied including land hermit crabs, stick insects,
millipedes, snails, spiders, locusts, crickets, etc.
Mealworms, crickets, locusts.
Monkfield Nutrition
Crickets, locusts, mealworms.
Peregrine Livefoods
Rep-Tech
Spiders, stick insects etc as well as live foods: mealworms, crickets,
etc.
Mealworms, crickets, locusts.
Small-Life Supplies
Stick insects, cockroaches, lepidoptera, snails, etc.
Southcoast
Invertebrates
Worldwide Butterflies
A wide selection including land hermit crabs, stick insects, millipedes,
snails, spiders, locusts, crickets, etc.
Lepidoptera and stick insects.
Butterfly Connections
V Cheeseman
Faunology
14.1.4
Keeping and handling animals
m It
is a sensible precaution for staff working regularly with animals, particularly in
rural studies courses, to receive anti-tetanus vaccinations at appropriate intervals. No
one with an unprotected open wound should normally be allowed to handle animals.
New staff should receive practical training in the various techniques for handling
safely and appropriately the animals kept in schools; in most cases this will be from
experienced staff. Guide L52 gives guidance on handling small mammals. Suitable
protective animal-handling gauntlets should be used where appropriate. Pupils will
also need training and close supervision when handling animals. Small mammals that
are handled regularly are most unlikely to bite.
Emotional
disturbance
m Hygiene
Housing
This can be caused if pupils perceive the treatment of animals, including invertebrates, to be cruel or inhumane, whether or not this is the case. Euthanasia, for
example, should not be carried out in the presence of pupils for this reason.
Strict hygiene is needed at all times. Everyone, staff and pupils, should wash
their hands before, as well as after, handling animals or coming into contact with
their bedding or, if aquatic, the water in their tanks. Adequate facilities nearby,
eg, hot water, liquid soap, disposable towels etc, are essential.
This is covered in section 8.9 (Animal housing) and in CLEAPSS Guide L56.
Weekends and
holidays
1414
© CLEAPSS 2006
Caring for animals during holidays is a major problem at a time when many
schools do not pay technicians to work during holidays and when access can be
difficult. When planning the keeping of animals, holidays must be a major consideration. Animals can generally be left unattended over the weekend, if steps are
taken on Friday afternoon to inspect the animal collection and check that all is
well. Ensure that small mammals and birds will have sufficient food and water to
last them until Monday morning; increase the number of water bottles if necessary. For other animals, check that water supplies are adequate. For periods longer
than a normal weekend, for example at half term, it is essential that animals are
inspected, fed and watered during this time.
Animals, particularly small mammals, should not be taken out of the school during holidays unless absolutely essential. There is the possibility that mammals
may become infected after contact with other disease-carrying animals. It is also
much more difficult to ensure that animals are cared for properly off-site. It will
be important to provide all necessary training, information, equipment, food etc
for pupils or other people taking animals home. Ideally, mammals should go to
homes where no other domestic mammals are kept and be segregated for a week
on return. It is sensible that parental assent in writing is obtained before small
mammals and other animals are taken home. Information should be provided on
animal care, including where additional foodstuffs and veterinary attention can be
obtained if needed.
Treatment of
animals
The Animals (Scientific Procedures) Act 19861 prohibits, without a licence, any
procedure on live vertebrates that causes pain, suffering, distress or lasting harm.
Significant departures from good husbandry may have such adverse effects. It is
therefore forbidden to train a mammal to negotiate the route through a maze if
the animal is kept hungry to reinforce appropriate choices by receiving a reward of
food. Similarly, observing blood flow in the fins of a fish taken out of water and
placed on a microscope slide is also a very questionable procedure.
In physiological studies, decerebration including the pithing of frogs is prohibited
but decapitation (followed by destruction of the brain) is not. This is, however, a
traumatic operation which students may find very distressing; it should be
undertaken only if really needed for A-level studies, after careful consultation and
by someone confident that he/she can complete the task quickly and efficiently.
Similar considerations also apply to investigations involving the observation of
chicks at different stages of development. It is not permitted to open a fertile egg
after more than half of the incubation period has elapsed. It is thus possible to
study, for example, developing chicken embryos up to a maximum age of 10 days.
Such work is again, however, extremely disturbing for many pupils and should
only be carried out after much thought and consultation.
Invertebrates are not defined as animals under the above Act but should be
treated as humanely as other animals, though there is no legal obligation to do so.
Specimens used in practicals or collected on field trips should not be left lying in a
lab to starve or dehydrate. They should be properly housed & cared for and
returned to their habitat or killed, humanely, only as a last resort. Undue stress
should be avoided in experiments such as observing the effect of temperature on
the heartbeat of Daphnia; the rise in temperature should not be excessive.
Injecting Xenopus If the resulting tadpoles are cared for, the injecting of Xenopus to induce spawning
can be considered to be for the purposes of husbandry and is not prohibited under
the Animals (Scientific Procedures) Act. Use a sterile syringe and needle. Make up
the gonadotrophin hormone in sterile distilled water. Hold the animal in a cloth to
give a good grip and prevent it struggling; this reduces the chances of injury to
both the animal and the operator. It is preferable for one person to hold the toad
while another performs the injection. [For more details of the technique, see
Nuffield Biology Teachers’ Guide 12. Note that the use of an anaesthetic such as
1
For more details, see Roger Lock, Investigations with animals and the Animals (Scientific Procedures) Act 1986 School Science
Review 71 (255), December 1989, pp 74-75.
2
Revised Nuffield Biology Teachers’ Guide 1, Nuffield Foundation, Longman, 1974.
© CLEAPSS 2006
1415
MS-222 to sedate the animal is no longer permitted.] The injection should be done
in the presence of pupils only if they will find the operation acceptable and if it
can be done deftly.
14.1.5
Feeding and cleaning
Animals fed well on an appropriate diet are more likely to remain healthy. Mammals
and birds need a constant supply of food and water; other animals can be fed less
regularly, depending on individual needs. Special dietary requirements should be
checked carefully and observed; eg, guinea pigs require vitamin C in their diet and
also good-quality hay. Many invertebrate and vertebrate animals require a good
supply of calcium. Reptiles and amphibians being fed on invertebrates benefit from a
variety of foods rather than a constant supply of just one food type and the provision of
a vitamin supplement which is dusted on their food.
Feeding
Use suitable hoppers or containers within the animal housing to prevent unnecessary wastage through soiling of food and to ensure that food is not buried as an
animal burrows through soil, bedding or litter. Ensure there is adequate water
available in an appropriate container which will reduce spills but give a constant
supply.
Foodstuffs
Buying in bulk pelleted food for mammals (eg, from Lillico) can be economical but
it does deteriorate in storage and should be used within six months; (three months
with diets containing a vitamin C supplement). Label stock and use in rotation. If
seeds are given as animal food, ensure that they have not been treated with
fungicides. A good source of high quality seeds is John E Haith. Sources of live
invertebrate animal foods are given in Table 14.2.
All dry food should be kept in heavy-duty plastic bags, closed with ties, bulldog
clips etc to reduce risks of infestation by insects, mites, fungi etc. In turn, the bags
should be kept in a container with a lid, eg, a plastic dustbin or a large biscuit tin
depending on quantity, to prevent attack by mice and other vermin. Hay also
should be kept away from vermin; a dustbin with a lid is ideal. If hay has an
infestation of mites, this is often dealt with successfully by storing quantities in a
freezer.
Fresh green food should be collected from suitable plants or bought locally, rinsed
and shaken dry before being introduced to the animal. Root vegetables and fruit
should also be washed before use. If carnivores are to be kept, the provision of a
constant supply of food must be carefully considered.
Cleaning
Animal housing must be easy to clean, secure and chosen to meet the needs of the
species; so must the media used including litter, bedding, gravel etc. [See section 8.9
(Animal Housing) and Guide L56.] Cages and tanks must be kept clean, not only for
the well-being of the animals, but also for that of pupils and school staff. A regular
cleaning routine is needed, the frequency depending on the species. During cleaning,
the animals should be handled sensitively by someone who has been trained in the
correct handling procedures.
m Spare housing
New-born litters
m Disinfectants
1
For most species, it is useful to have spare housing or other secure containers
available to which animals can be transferred while the normal housing is being
cleaned.
It is best not to disturb a cage of mammals in which young have just been born;
there is a risk that the young animals will be rejected or even eaten by the
mother. If at all possible, delay cleaning until the litter is about one week old.
For cleaning out cages, use safe, effective disinfectants1 such as ampholytic surfactants (Griffin ASAB, Harris BAS) or VirKon. Disinfectants should be freshly
made up to the correct concentration. Do not store diluted disinfectants.
See section 15.12.3 (Chemical disinfection).
Soiled bedding
m Precautions
1416
© CLEAPSS 2006
Soiled bedding should be disposed of carefully, ideally by incineration if practicable. Otherwise, it should be sealed in plastic sacks and disposed of with normal
refuse.
When cleaning cages, handling soiled bedding etc, it is sensible to wear gloves. In
handling dry bedding materials, aim to avoid raising dust.
Aquatic animals
See section 14.3 (Aquaria) and Guide L56.
Amphibians and
reptiles
Uneaten food must be removed before it decomposes.
Invertebrates
The amount of cleaning required will depend on the species kept but it is often
enough merely to remove droppings and any mouldy food. For locusts and stick
insects, sweeping out the bottom of the cage is usually sufficient but look out for
stick insect eggs in the debris.
14.1.6
Hazards to handlers
If suitable animals are chosen, obtained from a reliable source, handled sensibly and
good hygiene observed, the risks of disease transmission and injury are non-existent
or very small. Animals should be kept away from the face and individuals with open
wounds or sores should not be allowed to handle animals. Anti-tetanus vaccinations
for regular animal handlers may be advisable.
m Physical injury
Physical damage caused by animals biting and scratching can be avoided by
regular, proper handling and the use, where necessary, of suitable protection such
as gauntlets. Cages, aquaria and vivaria should be regularly inspected for broken
wires, sharp edges etc. As well as being a danger to the animals, they present a
risk to the handler. Cuts from these sources might cause infections.
m Allergic reactions
These may be caused in sensitive individuals by fur, skin, scales or feather particles etc when animals are handled regularly; there is often an initial period of
sensitisation when no signs are observed. People who have become allergic to a
particular species should normally avoid further contact with it.
Staff can develop an allergy to locusts by contact with air-borne particles originating from their faeces and exoskeleton. Symptoms include rashes, smarting eyes,
runny noses and asthma-like tightness of breathing. This is only likely to be a
problem if locusts are kept in continuous culture, which is no longer recommended. Where locusts are kept, adequate ventilation is essential, steps should be
taken to minimise dust when cleaning out the animals and the wearing of a dust
mask1, if raising dust cannot be avoided, may be beneficial.
Handling the larvae of moths and butterflies which have hairy bodies, particularly the brown-tail moth, Euproctis chrysorrhea, may also be hazardous; allergic
reactions have often been reported. With Euproctis the reaction can be very
severe; note also that other stages of this insect’s life cycle are also hazardous.
Where schools keep bees, care is needed since some people become sensitised to
their venom and show a very severe reaction to a sting.
m Zoonoses
1
These are diseases transmissible to humans from animals. They include Weil’s
disease, salmonellosis, tetanus, psittacosis or ornithosis, ringworm and parasitic
worms. Risks are often exaggerated and can be eliminated or reduced to very low
levels; good hygiene should be observed at all times. Rural studies in which various animals are kept may present the greatest risks. Small laboratory mammals
from a reliable source should not present a hazard. Wild rodents captured during
field work must be handled with particular care.
See section 3.3.5 (Respiratory protection).
© CLEAPSS 2006
14.1.7
1417
Health of animals
It is sensible to identify a suitable veterinary practitioner before one is needed to treat
animals that are unwell. Isolate animals presenting symptoms of illness and, if
necessary, consult the vet. Affected animals may have to be killed humanely. See section 14.1.8 (Anaesthesia and euthanasia).
Signs of illness
in mammals,
birds, reptiles
and amphibians
Checks should be made of the following: missing or elongated teeth or claws; damaged tails; patchy fur or feathers or loss of colour in skin or scales; swellings or
growths; exudates from the eyes and nose; sores on the skin and feet; scabs on the
tail; abnormal shape and colour of faeces. Examine all body openings for unusual
conditions. Note the behaviour of the animal: watch for abnormal movement or
breathing, unusual amounts of sleep, poor appetite and an unusual reaction to
handling. An animal that is ill may also show abnormal social behaviour.
Illness in fish
The following signs should be watched for:
• fungus (white or opaque patches);
• mouth ‘fungus’;
• white spot (white pinhead-sized bubbles on the body and fins);
• cloudy eyes;
• swim bladder disease (swimming on the side or upside-down);
• dropsy (swollen body with scales protruding).
Remove all diseased fish from the tank as soon as they are identified and treat
fungus, white spot, mouth ‘fungus’ (caused by bacteria) and eye infections with
proprietary medicines from a pet shop or aquarist. (The water may also need
treating.) It is sensible to have available a separate small ‘hospital’ tank (which
may also need aeration as some remedies lower dissolved oxygen levels) for the
isolation and treatment of sick animals. Fish with swim bladder disease or dropsy
may be observed for a short time to see if they recover naturally; then a vet should
be consulted, if the fish is particularly valued or valuable, or the animal killed
humanely; see the next section. For more details, consult fish-keeping manuals1.
14.1.8
Anaesthesia and euthanasia
Anaesthesia
The Animals (Scientific Procedures) Act 1986 does not permit any vertebrate to be
anaesthetised in school investigations. This includes the customary practice of using
the anaesthetic MS-222 with fish and amphibians. For invertebrates, chilling in a
refrigerator for a short period is successful but animals recover quickly.
m Drosophila
Ethoxyethane (diethyl ether) (EXTREMELY FLAMMABLE; HARMFUL) is commonly
used and is still quite acceptable. Take care not to use too much; the cotton wool
should be damp but not dripping. No naked flames or other sources of ignition
must be in the room. There should also be good ventilation.
An alternative anaesthetic is used in a kit called FlyNap2. ‘Wands’ are dipped in
the anaesthetic [50% triethylamine (HIGHLY FLAMMABLE; CORROSIVE; HARMFUL), 25% ethanol (HIGHLY FLAMMABLE) and 25% ‘fragrance’ (to mask the smell
of the triethylamine!)] and inserted into the bottle or tube containing the fruit
flies. Even with the fragrance, the anaesthetic smells unpleasant! A review of the
use and effectiveness of FlyNap is given in CLEAPSS Bulletin 106.
1
For example, the Interpet Manual of Fish Health, Chris Andrews et al, Interpet Publishing, 2002, ISBN 1842860674 or Interpet
Guide to a Healthy Aquarium, Neville Carrington, Interpet Publishing, 1999, ISBN 1902389565.
2
This is available from Blades Biological and Philip Harris.
1418
© CLEAPSS 2006
It is possible to anaesthetise Drosophila without the use of chemicals: chill the
flies using a freezer block, as used in cool boxes. Details are given in a technician
tip in CLEAPSS Bulletin 107.
Euthanasia
Euthanasia of vertebrates should only be carried out by a responsible person. Consult
a veterinary practitioner if in any doubt. Vertebrates should not normally be killed in
the presence of pupils.
m The following substances can be employed in school laboratories to kill animals humanely. Their use is discussed below.
Carbon dioxide
1,1,1-trichloro-2-methylpropan-2-ol (chlorbutol)
Ethoxyethane (diethyl ether)
Ethanol
MS-2221
Magnesium chloride
Menthol
HARMFUL
HARMFUL
EXTREMELY FLAMMABLE
HIGHLY FLAMMABLE
HARMFUL
No serious hazard
HARMFUL, IRRITANT
Other substances have been suggested for euthanasia, particularly in older reference
texts. The risks of using several of these are substantially greater than the chemicals
listed above. Consequently the following substances are not recommended for school
use.
m
Trichloromethane (chloroform)
2,2,2-trichloroethanediol (chloral hydrate)
HARMFUL
TOXIC
Ethyl carbamate (urethane, ethyl urethane) MAY CAUSE CANCER
Potassium cyanide (in eg, killing bottles)
VERY TOXIC
Small mammals
and birds
It is recommended that small mammals to be killed are left in their cage wherever
practicable; this reduces stress caused by removing the animal to an unfamiliar
environment. The entire cage is placed inside a heavy gauge polythene bag and
carbon dioxide from a cylinder introduced very slowly into the cage. The gas will
displace the air which should be allowed to escape from the neck of the bag. Leave
the bag sealed for 20-30 minutes after movement ceases to ensure that death is
certain. The same procedure can also be used for birds. The use of ethoxyethane or
trichloromethane is not considered to be a humane method of killing. Mechanical
methods are humane if used by an expert, though they may appear to be barbaric.
Fish and
amphibians
Use a solution of MS-222 (amphibians: 0.6%; fish: 0.01%) to which the animal
should be exposed for a prolonged period. Alternatively, chlorbutol can be used
(amphibians: 0.2%; fish: 0.03%).
Insects and
small terrestrial
arthropods
A killing bottle may be filled to a depth of about 2 cm with well-bruised laurel
leaves (Prunus laurocerasus, not variegated varieties). It should be left stoppered
for 2-3 days before use and will remain effective for at least a month. Alternatively
ethyl ethanoate (acetate) may be used.
Earthworms
Immerse the earthworms in a solution of dilute ethanol (less than 30%).
Aquatic
invertebrates
10% ethanol, saturated magnesium chloride solution mixed with an equal volume
of the animal’s water, menthol crystals scattered on the water surface or carbon
dioxide in soda water can all be used.
1
The methanesulfonate of ethyl 3-aminobenzoate.
© CLEAPSS 2006
14.2
1419
Animals in the wild
Wild animals in England and Wales are protected by law, particularly by the Wildlife
and Countryside Act 1981. Elsewhere in the UK, the relevant legislation is the Nature
Conservation (Scotland) Act 2004 and the Wildlife (Northern Ireland) Order 1985. In
each part of the UK there is a separate body responsible for wildlife conservation:
English Nature, Countryside Council for Wales, Scottish Natural Heritage and the
Environmental Heritage Service (Northern Ireland).
The main legal restrictions affecting schools are usefully summarised in the DES
Administrative Memorandum 3/90 that has superseded the previous AM 1/89 (which
contained serious errors). Schools should note, however, that this revised document
still contains errors1 and does not include some relevant information. The booklets
Wildlife, the Law and You (Nature Conservancy Council 1982) and Protecting Britain’s Wildlife: a Brief Guide (Department of the Environment 1988) are now out of
date and out of print. For details of protected animals, use the web site of the Joint
Nature Conservation Committee www.jncc.gov.uk; click on ‘legislation’ and ‘Wildlife &
Countryside Act’.
Birds
All wild birds, their eggs and nests in use are protected in various degrees; birds
must not be killed or captured. For some species it is necessary to obtain a licence
from the relevant conservation body (see above) before approaching or photographing them while near their nests. Refer to the JNCC web site (above) and the
RSBP2 for further information on the legislation protecting birds.
Other animals
Over 40 species of other animals are similarly protected and must not be killed,
injured or taken, even for ecological marking studies. Damage to animal shelters
etc is also prohibited. The protected species include bats3, dormice, red squirrels
and common otters as well as some reptiles, amphibians, fish, butterflies, moths,
crickets, dragonflies, beetles, grasshoppers, spiders, snails and other invertebrates. Some mammals that are not protected must nevertheless not be killed or
taken. (See AM 3/90 and/or the JNCC web site, referred to above.) A licence from
English Nature etc, rarely granted, is needed before fully-protected animals can
be taken for educational purposes. English Nature etc will, however, grant licences to schools to capture shrews4 or for pond-dipping etc activities where great
crested newts are resident, provided that they are released unharmed after
capture and study.
The common frog, viviparous lizard, palmate and smooth newts, slow worm, grass
snake and common toad may, however, be collected and studied without a licence,
despite a widely-held belief that they are fully protected.
Trapping
animals
When surveying small mammals in an area, Longworth traps should have a
13 mm hole to allow shrews to escape5. (If shrews are to be trapped intentionally,
a licence is needed; see above.) The trap should contain suitable food and bedding
and great care must be taken when captured animals are removed to prevent inj-
1
In section 20, AM 3/90 should state that it is illegal to trap shrews intentionally without a licence. If shrews are trapped by
chance without a licence, this is not illegal. Also, if schools wish intentionally to capture and study the great crested newt, then a
licence is needed but can be obtained from English Nature etc. In section 21, a licence is not needed to uproot protected plants
other than those listed in Annex F of the AM; the permission of the land owner, however, is needed. In Annex C, it should be
noted that all species of British butterflies, not already listed, may not now be sold, either alive or dead, without a licence.
2
Materials produced by the Royal Society for the Protection of Birds include Bird Photography and the Law (26-039, free), Birds
and the Law (26-005, free), Birds and the Law - Some Questions Answered (26-004, free) and Wild Birds and the Law - a Plain
Guide to Bird Protection Today (26-013, priced publication).
3
English Nature issues the publication Focus on Bats; other materials are available from the Bat Conservation Trust. English
Nature also publishes various booklets about other protected animals that may be encountered in gardens and local areas.
4
General guidance on trapping shrews is provided by English Nature etc with the licence.
5
Without a means of escape, it is likely that shrews will die in the trap. Their small size (and therefore large surface area: volume
ratio) means that shrews must eat throughout the day and night to provide sufficient energy to maintain body temperature.
1420
© CLEAPSS 2006
ury to the handler or the animal. The Mammal Society has produced a helpful
booklet Live Trapping Small Mammals - a Practical Guide, which should be
consulted if possible before any fieldwork is contemplated. The accompanying publication, Discovering Mammals: Practical Projects for Young People, is also
valuable.
When invertebrates are caught in pitfall traps, or moth traps emitting u-v radiation are used, it is important that the captured animals are released unharmed
in the area where they were trapped after studies are complete. The use of sticky
and water traps for insects which involves killing the animals should be contemplated only if really necessary. Before studying insects, teachers should consult
the Code for Insect Collecting, produced by the Joint Committee for the Conservation of British Insects.
m Schools should be very cautious about bringing live wild mammals into school; even if
it is legal to accept a particular species, they may carry parasites and diseases that
can infect humans. They must be kept apart from mammals already in the school.
Dead animals and birds should not be accepted. Pupils should be told not to bring in
wild mammals but, if a pupil arrives with an injured bird or mammal; it would not be
humane to refuse to take in the animal and deal with its injuries. See section 14.1.2
(Animals from local habitats).
Care of wild
animals
In the interests of conservation, animals legally brought into schools, eg, after
field trips, should be properly and immediately cared for and returned as soon as
possible to a suitable habitat, preferably at the place where they were collected.
For many invertebrates, they will need humid conditions to ensure that they do
not desiccate. It must be realised that, for mammals in particular, bringing an
animal into school removes it from a carefully-balanced environment, from its own
territory and possibly from its young. If returned only a few yards from where it
was caught, it may be in another animal’s territory and have to fight for its life.
Animals that hibernate should be released well before the winter to allow them to
build up the fat stores essential for their survival.
If small mammals are trapped or collected for short-term studies, there should be
close supervision, careful handling while wearing protective gloves, strict hygiene
and, if brought into school, total isolation from the school’s stock of animals. Obviously, the mammals must be properly housed and fed; since they are not used to
captivity, particular care must be taken to reduce stress as much as possible. This
requires planning prior to trapping.
14.3
Aquaria
m NB
With aquaria, the combination of mains-operated electrical accessories
and impure water presents a particular hazard. Special care must be taken
with wiring and any local authority regulations must be obeyed. See section
14.3.4 (Accessories and electrical safety).
A tank of fish or other animals provides constant movement and interest in a room
but, more importantly, will provide opportunities for a variety of studies in the Science
National Curriculum. The animals in the aquarium can be used to introduce pupils to
aspects of the variety of life, classification, variation or the processes of life and to
encourage the development of observational, hypothesising, measuring and recording
skills. It is important, however, to ensure that an aquarium is used for educational
work and not merely ignored once it has been set up1.
1
A valuable reference book for establishing and maintaining aquaria is The Aquarium Technology Handbook by A Jenno, David
& Charles (1985) but this is now out of print. There are many popular books on aquaria that may be referred to; a particularly
good range of books is available from Interpet Publishing. The more detailed manuals or titles dealing with specific groups of
fish will be found to be most useful.
© CLEAPSS 2006
1421
There are certain procedures and considerations that are involved in the establishment of any type of aquarium; these are included in section 14.3.1 which discusses the
requirements of cold-water organisms. The additional demands of setting up tropical
fresh-water and marine aquaria are considered in sections 14.3.2 and 14.3.3. The use
of electrical accessories, safety considerations and maintenance of equipment are discussed in sections 14.3.4 and 14.3.5. See Table 14.1 in section 14.1 for references that
may provide useful information when contemplating keeping certain animals in the
aquarium. There is also further discussion on setting up aquaria in Guide L56.
14.3.1
Cold-water aquaria
A cold-water aquarium will usually require only a small amount of regular maintenance, though this will be influenced by the type and number of animals and plants
that it contains. Once most fresh-water aquatic animals are established, they usually
live for a long time and arrangements for their care during the holidays are often easy
to make. An aquarium of cold-water fish, such as goldfish, is often set up but a coldwater tank is also a suitable home for other animals. These include the entirely aquatic Xenopus clawed toad, the axolotol (an unusual salamander which becomes adult
though still a tadpole) and the tadpoles of common frogs or toads hatched from spawn.
Various other animals such as crayfish and a wide variety of other invertebrates can
also be housed.
If the aquarium is to house only a very limited number of small animals, it is possible
to set up a tank without the complications of air pumps, filtration and lighting. If,
however, the intention is to stock the tank with rather more animals than this, at
least filtration of wastes from the water will be needed. If starting on a small scale,
when making an initial purchase of an aquarium it is sensible to think ahead and consider how it might be used in the future. Aquarium accessories can be obtained later
but it will then be too late to change the size of the tank.
Setting up the aquarium
Types of tank
1
There are three types that are used.
a)
Tanks with the glass held in place with putty and supported in a frame of
angle-iron. These have been almost completely replaced by all-glass tanks
(see below). As old or second-hand tanks of this type often leak, it is best not
to use them as aquaria; instead keep them for housing plants, gerbils, stick
insects etc. If you do use one as an aquarium, all the internal joints should be
sealed with silicone rubber adhesive before water is put in. [Make sure that
you use a sealant suitable for aquaria; some types contain chemicals to kill off
fungus and these will harm the animals. Also, mend the tank in a wellventilated area; the sealant gives off a strong vapour.]
b)
All-glass tanks with the pieces of glass held together by silicone rubber adhesive; these should be professionally made1. This is the type that should normally be chosen for an aquarium but check to see that the edges of the glass are
not sharp if these are exposed. In many cases, tanks of this type are supplied
with a plastic or metal frame. This does not add to the strength of the tank
but is useful to protect the edges of the glass.
c)
Plastic tanks. These are inexpensive but they are usually too small and very
easily scratched. They are not recommended for use as ‘permanent’ aquaria
but are very useful as temporary housing for animals. Gentle polishing with a
soft cloth and brass polish can be used to remove some of the scratches. Then
the tank should be washed with detergent to remove the polish.
A local aquarist or pet shop will be able to supply suitable aquaria and sometimes more cheaply than large firms such as Philip
Harris or Griffin Education. Look in Yellow Pages under ‘Aquarium and Pond Supplies’ for details of nearby stockists. A
reputable manufacturer of high-quality aquaria is John Allan Aquariums; ask for details of local suppliers of John Allan products.
1422
© CLEAPSS 2006
Size of aquarium A minimum size for an aquarium is about 60 cm long, 38 cm tall and 30 cm deep
(24×15×12 inches). A height of 38 cm is best for viewing the organisms in the
aquarium. The other dimensions depend on the size and number of the animals
that will be housed, the space available for the tank and financial considerations.
Oxygen dissolves into an aquarium at the surface of the water, so it is important
to achieve the right balance between the size of the tank and the numbers of
animals which extract oxygen from the water. A rule-of-thumb suggests that
60 cm2 of water surface should always be allowed for each 1 cm of animal. More
animals can, however, be accommodated if the water is aerated.
Another factor should also be considered if, initially, an unilluminated aquarium
is to be set up which might need a lighting hood later. In order to take 60 or 90 cm
fluorescent tubes, hoods need to be about 10 cm longer. It is, therefore, sensible to
purchase, for example, a 70 or 100 cm tank instead of the more regular sizes.
Preparation of
the tank
Before the tank is filled, the back and possibly the sides should be painted or
covered with paper or card on the outside. This is to prevent light entering from
the sides and make the animals in the aquarium more visible; black, dark-green
or blue are colours often used. Emulsion paint is suitable and permanent; it could
be a nuisance, though, if the tank is used for something else later. If paper or card
is used, be prepared to replace it quite regularly when it peels off or becomes wet!
Finally, an all-glass tank must be supported on a soft base, for instance, polystyrene ceiling tiles, unless it has an integral base.
Position of
aquarium
A tank full of water is very heavy and will require a solid table or bench to stand
on. It should not be placed near a window where it will receive too much light
from the Sun and be liable to extremes of temperature. It is sometimes possible to
site the aquarium out of direct sunlight but still with enough light for the animals
to be viewed without having to install a lighting hood. If there will be plants in the
aquarium, artificial lighting is almost certainly essential. [See section 14.3.4
(Accessories and electrical safety).]
Another factor in siting the aquarium is to consider its accessibility for pupils.
They will need to be able to observe the animals clearly. If pupils are to help in
maintaining the aquarium, it needs to be at the right height. Alternatively, it
might be more important with some students to restrict access to the tank!
Gravel
The floor of the tank should be covered with 4-7 cm of aquarium gravel. This
should have a deeper layer at the back, sloping gently towards the front. This will
mean that pieces of debris will fall to the front of the tank from where they can be
removed more easily. The gravel is best bought from an aquarium shop and must
be thoroughly washed before putting it in the tank. If an under-gravel filter is to
be used, make sure that it is the right size for the tank and the airlift tube is fitted
before adding gravel on top. All parts of the filter must be covered at all times.
Half the gravel is often added first, then a layer of nylon mesh, followed by the
rest of the gravel. This ensures that digging animals cannot uncover the filter.
Rocks, wood
and cork bark
Rocks can be used to make the aquarium look more attractive and they may help
to keep the gravel sloping in the tank. Not suitable are pieces of broken concrete,
rocks with sharp edges and rocks that will slowly dissolve, such as chalk or soft
limestone. Choose pieces of granite or slate and smaller rounded pebbles; a local
aquarist shop should have stocks of suitable material. Pieces of well-washed
natural coal can look attractive, particularly if the back of the tank is not painted
black. Rocks can be positioned to hide things such as filters and their tubes.
Pebbles can be useful to anchor the roots of plants.
Wood and cork bark supplied by an aquarist provide variety and possible shelter
for some fish. They can, however, easily float, so they must be anchored in place
with rocks. Even then, they may come adrift; to avoid this the best remedy is to
stick the material to a piece of slate with aquarium sealant. The slate is then
buried beneath the gravel.
© CLEAPSS 2006
Water
1423
Ordinary tap water will usually be fine, but the chlorine in it can harm some
delicate animals. The chlorine is dissipated if water is left to stand at least
overnight. If the aquarium is fitted with filters or air pumps that stir the water,
these help the chlorine to escape. Delay putting animals into the tank until the
water has cleared. When topping up the tank later, always try to leave water to
stand first. Alternatively, dechlorinating tablets from an aquarist can be used.
If plants are to be added to the aquarium, do not overfill the tank at this stage;
allow about 10 cm space at the top for water displaced by your arms while planting. Add the rest of the water afterwards. To avoid disturbing gravel when adding
water, pour it onto a small saucer or some other object. A tip for removing any
scum which may collect on the surface of the water is to draw a piece of kitchen
paper towel over the surface. When an aquarium has become established, between
a fifth and a quarter of the water should be siphoned off and replaced at intervals
of around 3-4 weeks.
Covering the
tank
A tank requires a cover to reduce evaporation, to stop animals jumping out, to
keep out dust and flies and to deter the unwanted explorations of pupils. Tanks
are often sold with a hood but there should also be a cover inside the top of the
tank. Sheets of glass should be avoided if possible; plastic covers to fit various
sizes of tanks are available which can be cut to the exact size. Alternatively,
Perspex sheets might be used. Make sure that the transparent cover is always
clean, otherwise light entering the tank will be considerably reduced. There must
always be a cover between the water and any electric lighting, allowing holes for
tubing and access for feeding.
Lighting
A tank can often be placed where it will receive enough natural light for its
contents to be seen. Too much direct sunlight will, however, make the water too
warm and it may turn green due to algal growth. For this reason, tanks must not
be placed on a window ledge. It is sensible to think about artificial lighting when
first obtaining an aquarium as it is possible to buy a tank with a properlydesigned and fitted cover which houses the lights and encloses all electrical
connections. See section 14.3.4 (Accessories and electrical safety) for details of the
most appropriate means of providing artificial illumination. Some animals prefer
more-shaded conditions; provide for their needs by including shelters, eg, cork
bark, beneath which they can hide.
Maintaining water quality
For a successful aquarium, maintaining the quality of the water and, therefore, the
health of the inhabitants, is vital. This will normally involve the use of some form of
filtration, though plants also have a part to play. In most circumstances, when an
aquarium is well managed it should not be necessary to monitor quantitatively various parameters such as pH, nitrite or nitrate levels. Such tests may, however, have
an educational benefit when discussing the needs of animals and the biological events
that are involved in the breakdown of wastes; simple kits are available from aquarists
for these tests.
Aeration
An aerator is not in fact essential in a tank. A stream of bubbles from an air stone
may look attractive but it does not directly help to supply oxygen to animals in the
tank. Providing the aquarium does not contain too many animals, enough oxygen
will dissolve into the water through the surface. Aeration does, however, cause
water to circulate in the tank, and this can help to replenish some oxygen. This is
why an aerated tank can hold more fish than should normally be allowed.
Before using an air pump, refer to section 14.3.4. A blocked air stone can usually
be cleared by holding it under a running hot water tap or by immersing it in a
beaker of boiling water for a few minutes.
Filters
Filtration is useful because it removes small particles from the water and wastes
from the animals. A small tank can be successful without a filter but the water
will need to be changed very regularly to remove wastes; the water may also be
cloudy. A filter is therefore a good idea to reduce routine maintenance.
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© CLEAPSS 2006
There is a wide choice of filters, all with advantages and disadvantages. Some just
filter out particles, some remove organic wastes, while others do both. Apart from
considerations of cost, the type of filter used is often determined by a desire for the
unit to be hidden and not spoil the appearance of the aquarium.
An under-gravel filter is quite cheap and consists of a plastic base plate with an
attached tube. For a big tank, two filters may be needed. Once installed, the filter
is operated either by an air or water pump, is invisible and generally requires
little attention. It works by drawing water through the gravel; this will help to
remove particles and also cause animal wastes to be broken down by bacteria in
the gravel. As wastes such as ammonia are removed, nitrates are formed. If there
are plants in the aquarium, the nitrates will be taken up as fertiliser and encourage growth. Without plants in the tank, the nitrates may stimulate the growth of
unwelcome algae. Regular changing of about a fifth to a quarter of the water in
the tank will, however, keep the nitrate levels under control in these circumstances.
An under-gravel filter is placed in the tank with its uplift tube in a back corner. If
used with a water pump that fits on top of the uplift tube, the pump needs to be
completely immersed in the water; it is, therefore, important to keep the aquarium always topped up with water as it evaporates. This type of water pump
cannot be used with older types of under-gravel filter that have a narrow uplift
tube.
Box filters are also used. These work by drawing water through a medium such
as polymer wool that removes particles. Activated charcoal is also included to
remove chemical wastes. The filters either sit inside the tank or are fitted outside.
They do require regular cleaning out and can look unsightly. A similar criticism
can be made of filters made from plastic foam but these do have the advantage
that bacteria and microscopic animals colonise the medium and so they perform
biologically as well as mechanically.
Finally, there are power filters, for internal or external use, which remove
wastes by pumping water through a compartment filled with filter medium. They
provide the best means of filtration but can be quite expensive. Before fitting a
power filter, refer to section 14.3.4.
Plants
By utilising carbon dioxide and some organic wastes, as well as producing oxygen,
plants are helpful in maintaining water quality. They can, however, cause more
problems than they solve! See the next part of this section.
Snails
Water snails are not essential in the aquarium and many would argue that, if
there are plants, they are an undesirable addition. They may, however, remove
some of the algae from the glass or plants and they will clear up food missed by
the fish. There are several types of large or small snails with flattened or pointed
spiral shells and, on educational grounds, the inclusion of various snails in an
aquarium can be justified, since they will introduce pupils to another group of
invertebrates.
Cleaning and
other maintenance tasks
The amount of cleaning that has to be done will depend on whether an aquarium
has a filter unit and, if so, which type. With an under-gravel filter, and provided
the animals are not overfed, an aquarium should require little attention; an occasional raking of the gravel should be all that is required. If the water is clear and
has no smell, then conditions are likely to be satisfactory for the fish or other
animals.
With other types of filters, the filter medium will need to be washed or changed at
intervals. How frequently will depend on the number and type of animals kept
and the size of the tank. The glass on the front and sides of the tank may become
covered with algae, especially if the aquarium has artificial lighting, and should
be scraped off regularly.
Particularly if there are plants in the tank, there may be a gradual build up of
debris on the surface of the gravel. This is easily removed using a siphon tube,
with care taken not to suck up too much gravel at the same time.
© CLEAPSS 2006
1425
The only other regular tasks will be to top up the water level in the tank as it falls
as a result of evaporation and to change about 20-25% of the water every 3-4
weeks. An under-gravel filter increases the concentration of minerals in the water
and this may encourage algal growth. Regular changes of water will reduce the
build up of minerals and help to solve the problem. Plants in the tank will,
however, take up some of the minerals and reduce the need for water changes.
Remove the chlorine from the replacement water as described earlier.
Table 14.3 summarises the regular routine chores involved in looking after an aquarium.
Table 14.3
Care of aquaria: Routine maintenance tasks
TASK
ANIMALS
WATER
Each
Day
Check their number and health
✔
Feed appropriately.
✔
Each
Week
✔
✔
✔
Remove dead/floating leaves and sediment on leaves.
✔
Prune and reposition plants.
✔
✔
For under-gravel filters - rake gravel gently.
✔
For box and power filters - replace filter medium.
GENERAL
✔
✔
Check and replant any that have come adrift.
Look out for, and remove, filaments of green algae.
FILTERS
✔
✔
Change 20-25% with fresh water.
PLANTS
When
Needed
✔
✔
Top up level in the tank.
Each
Month
✔
Remove algae from front/sides of tank.
Siphon debris from surface of gravel.
Clean transparent cover at top of tank.
Check all accessories for normal operation.
Check wiring of electrical equipment.
✔
✔
✔
✔
✔
✔
✔
✔
✔
✔
Plants
It is much more difficult to keep plants alive and well than it is to keep fish or other
animals in a healthy condition. Artificial lighting is essential for most plants and
should be of sufficient intensity to stimulate growth. Do not feel that you must have
plants in your tank; an aquarium with unhealthy and dying plants inevitably conveys
very negative messages. An attractive display can be created using just rocks, wood
and cork bark but, without plants, regular changing of some of the water will be
needed to keep levels of minerals in check. If the aquarium is to have plants in it,
cleaning the tank more frequently, removing dead leaves, replacing the plants at
intervals and seeing prize specimens nibbled by fish, must all be expected. Plants may
be a continual trouble and expense but they can enhance the appearance of an
aquarium considerably, as well as helping to maintain water quality.
Choosing plants
1
A local aquarist shop will be able to recommend suitable plants that are currently
available1. Some species do not grow well in cold-water aquaria, so specify particular requirements. Choose tall plants with strap-like leaves for the back and
sides of the tank. Put small, bushy plants near the front of the tank and at the
base of rocks.
Reference may also be made to publications such as the Interpet Guide to Aquarium Plants, Interpet Publishing, 1999, ISBN
190238959X.
Encouraging
healthy growth
1426
© CLEAPSS 2006
It is sometimes recommended that the roots of plants should be slightly trimmed
to encourage new growth. When planting, do not bury the crown (the junction
between roots and leaves) in the gravel or the plant may rot. A helpful tip is to
bury the plant too deeply and then gently pull it out to the correct position.
Under-gravel filters are sometimes blamed for poor plant growth but there is no
conclusive evidence for this assertion. What is important is to ensure that the
layer of gravel in the aquarium is sufficiently deep.
The importance of minerals in the water must not be forgotten. Plants will obtain
some from the decomposition of organic wastes in an under-gravel filter. If,
however, an external filter, or none at all, is being used, the plants are likely to be
deprived of nutrients. This problem can be tackled by adding fertiliser to the
water; contact a local aquarist for advice. An alternative strategy that is sometimes recommended is to root some or all of the plants first in shallow pots of
potting compost or ‘plant plugs’ and then bury these in the gravel.
Floating plants
Some plants do not root in the gravel but float on the surface or cling to rocks and
wood. Too many floating plants should not be encouraged as they may cut down
much of the light needed by plants lower down. A plant called willow moss
(Fontinalis) is very attractive and particularly useful for fish that may breed. Eggs
can be laid in the moss which adults will not then be able to find and eat. This
plant is also very useful if larger animals such as clawed toads or axolotls are
kept; it cannot be uprooted or broken as the animals charge through the undergrowth! Willow moss is also very effective in absorbing nutrients from the animals’ wastes.
Artificial plants
An aquarium will be much easier to maintain if artificial plants are used. These
are available at aquarium shops but they are quite expensive, often look garishly
unattractive and are not to everyone’s taste. As algae grow on their surfaces, this
will, however, help to subdue their colour and, if they are copies of real plants,
make them appear more natural. The algae will also carry out some of the activities that make living plants an important part of a balanced aquarium.
Removing
filamentous
alga
Filamentous alga, sometimes called ‘blanket weed’, is definitely undesirable in an
aquarium. If this becomes established, it may spread and cover everything. Keep a
watchful eye for the first signs of its arrival and try to remove it at once. Use a
thin cane to ‘twirl’ up the filaments.
Animals
A cold-water aquarium can be used to keep more than just goldfish! The types of
animals to be kept will, of course, be determined by the educational objectives of the
curriculum being followed but some details for more popular animals are given below.
Refer also to Table 14.1 in section 14.1.1.
Fish
For suitable cold-water species, refer to section 14.1.1 (Choice of suitable species).
Stocking the
aquarium with
fish
With the rule of thumb suggested earlier of 60 cm2 of water surface per cm of fish,
a 60 cm tank will support 30 cm of fish. If four 5 cm fish are bought and placed in
a tank of this size, they will have plenty of room for growth. When purchasing
fish, always spend some time observing their general condition and behaviour; if
in any doubt about their health, do not buy! Before adding fish to a new aquarium,
it is sensible to let the tank settle down for a day or two first. Also the bag
containing the fish should be floated on the surface of the water in the aquarium
for an hour to allow temperatures to even out. If adding new fish to an established
aquarium, there is always the risk of introducing disease to healthy stock. If at all
possible, new fish should be quarantined for up to 2-3 weeks in a spare tank and
watched for signs of disease developing. When moving fish, use nets made from a
soft cloth rather than a coarse, hard mesh that can damage the animals’ skin or
scales.
© CLEAPSS 2006
1427
Amphibians
Suitable species are outlined in section 14.1.1, further information is provided below.
Xenopus
clawed toads
Once adult, these animals remain entirely aquatic throughout their lives, though
they come to the surface to breathe air. They are powerful swimmers with strong
back legs and a good cover on the tank is important to prevent escapes. Because
the toads do not extract oxygen from the water, the rules governing animal numbers and tank size do not apply. The adult toads should not, however, be kept with
other animal species in the same tank; Xenopus tadpoles must also be reared
separately.
Axolotols
These overgrown tadpoles retain their feathery external gills, never leave the
water and breed as tadpoles. It is just possible to keep two small axolotls together
in a 60 cm tank, but they may snap at each other if confined and would be better
in a larger tank.
Tadpoles of
common British
amphibians
Despite a commonly-held belief that frog or toad spawn must not be collected,
there are in fact no legal restrictions that stop schools studying the spawn of the
common species obtained locally, though conservation issues must be considered.
Only a small number of tadpoles should be kept; a clump of spawn about the size
of a tennis ball can contain as many as 350 eggs. To ensure that as many young
animals as possible become adults, frog or toad tadpoles should be reared as if
they are fish; filtration of their water is essential. For details of rearing tadpoles,
refer to CLEAPSS Guide L206.
To allow for metamorphosing tadpoles leaving the water, stock the aquarium with
pond weed reaching to the water’s surface; pieces of floating cork bark could also
be useful. The young frogs or toads are quite difficult to rear and so arrangements
should quickly be made to release them into the wild, preferably at their spawning
site, if this is known. Place the young animals in damp vegetation by the water’s
edge, and try to release them in the late evening so that they have time to hide
overnight from predators.
Feeding
Fish
These, like any other living creatures, require a balanced diet. This can be supplied by any of the proprietary fish foods that come in a variety of forms including
flakes, pellets, tablets and powders. Choose floating flakes or pellets for fish that
feed at the surface; tablets and granular foods sink more rapidly and will be taken
by bottom feeders.
Try to provide a variety of food types, including freeze-dried, frozen or live foods
such as water fleas, Tubifex, larvae and worms1 as an occasional treat. Food
should be given twice a day but provide only as much as can be eaten in three to
five minutes; the amount that can be piled on a drawing pin is a sensible quantity
to start with. Any filter should be turned off during feeding time so that the food
stays longer on the surface or floating in the water.
If the fish are fed regularly during the week they will be quite capable of lasting
the weekend without being fed as they are cold-blooded animals and can survive
prolonged periods without food. Feeding once or twice a week during the school
holidays with the normal quantity of food will be adequate. It must be noted that
overfeeding can kill (uneaten food pollutes the water and consumes oxygen);
underfeeding does far less damage.
Xenopus and
axolotls
1
Both adults are carnivorous but can be fed easily on maggots, mealworms, pieces
of earthworms or small slices of liver, heart or other red meat. Complete diets for
clawed toads may be available in the form of pellets and are very convenient.
Toads that are accustomed to fresh food may not, however, immediately take to
this artificial diet. Clawed toads are voracious feeders, detecting food by their
Local aquarists should be able to supply a variety of fresh and frozen foods. Refer also to the list of suppliers of live foods in
Table 14.2 of section 14.1.3.
1428
© CLEAPSS 2006
sense of smell. Axolotls may need to be trained to accept food, by dangling it in
front of their mouths. As with fish, do not overfeed or leave uneaten food in the
water to rot. Feed three times a week; the animals will come to no harm without
food over the weekend. During holidays, feeding could be reduced to once or twice
a week if necessary.
Tadpoles
Xenopus tadpoles are filter feeders and require liquid or powdered preparations,
as supplied for fish fry, which they sieve out of the water. Turn off any filtration
unit and supply sufficient food to be eaten in less than an hour, two or three times
a day. Initially, most frog and toad tadpoles are plant eaters but, when they start
to grow their hind legs, they become progressively more carnivorous. Tadpoles of
common frogs and toads can be fed on flake or pelleted foods, starting with a type
made from plant products and later switching to a food prepared for carnivorous
fish. Floating food pellets show more clearly how much tadpoles can eat within an
hour or so of being fed. Varying the diet for older tadpoles to include occasional
feeds of small pieces of chopped liver or heart is desirable. As always, the rule is
not to overfeed and not to leave uneaten food in the water.
The tadpoles of newts and salamanders require live food such as hatchling brine
shrimps, micro-, white or grindal worms, Tubifex, Daphnia or bloodworms. A local
aquarist should be able to supply some of these foods.
Disease
Disease is rare in a well-established aquarium but may be introduced with new
animals (if not quarantined), new plants or live food. Several fish diseases can usually
be treated successfully with proprietary medicines1 from a pet shop or aquarist.
Clawed toads and axolotls are relatively hardy animals and do not often show signs of
disease. See sections 14.1.7 and 14.1.8 for symptoms and treatment.
Weekends and holidays
An aquarium will require no attention over the weekend; lights should be turned off
(unless they are on a time switch) but air pumps and filters left working. During holidays, unless lighting is switched on and off with a time switch, replace the hood with
a transparent sheet so that daylight can shine through the top of the tank.
Feeding animals Feed once or twice a week with the normal daily quantity of food, as described
earlier. During the Christmas holiday when school heating may be greatly reduced, the animals will be less active and need less food anyway as a result of the
lower temperatures. If the animals are to be fed by someone else, provide a
written list of instructions on feeding together with the necessary foods. Make
certain that he or she knows that overfeeding is dangerous, so that there is no
temptation to give the animals just that little bit extra. It is also sensible to provide some basic information on the accessory equipment in case of an emergency.
14.3.2
Fresh-water tropical aquaria
The general advice given in section 14.3.1 is equally applicable to the establishment of
heated aquaria. See also information in CLEAPSS guide L56.
There is a much greater variety of colourful and interesting tropical fish than of
species living in cold water. Similarly, there are certain species of amphibians and
fresh water turtles or terrapins that require a warm environment. To provide for these
animals, their water must be warmed by a thermostatically-controlled electrical heater or heaters. A thermometer will be needed to check that all is well.
1
Reference texts can also be consulted, for example, the Interpet Guide to a Healthy Aquarium, Neville Carrington, Interpet
Publishing, 1999, ISBN 1902389565.
© CLEAPSS 2006
Using heaters
1429
For full details of their operation, refer to section 14.3.4 (Accessories and electrical
safety). The heater(s) should have an appropriate total wattage for the size of tank
and should be of sufficient power to maintain the required temperature (for
example, around 24 oC for most tropical fish) even if the room heating is switched
off in the winter (which may happen at weekends and during holidays). As a
general guide, allow 10 W for each 4.5 litres of tank volume. Aquarium heaters
are available in a range of ‘sizes’; choose the heater that most nearly matches your
requirements. For a typical aquarium, eg, 60 × 40 × 30 cm, a 150 W heater would
suffice in a normally-heated room but a 200 W heater might be needed if the room
temperature were to fall very low. To achieve an even temperature throughout the
water in a large tank, use a separate heater at each end; ensure that their combined wattage is correct for the size of tank.
A heater can be installed vertically or horizontally and, if a combined heater and
thermostat is used, ensure it is positioned so that the thermostat is not at a level
below the heater. It is usual to hide the heater behind rocks or plants but ensure
that, if held horizontally, it is not too near the gravel and that there is a good
circulation of water around it.
Covering and
lighting the
aquarium
Especially with heated aquaria, a cover is required as, otherwise, evaporation of
the warm water will put an extra load on the heater and cause the water level to
fall, perhaps uncovering the heater or thermostat and further reducing their efficiency or causing overheating and fatalities. The cover should be designed so that
the water condensing on its inner surface drips back into the aquarium and not
down the outside.
A transparent cover may allow enough sunlight into an aquarium to view the
inhabitants but, to maintain a plant population, artificial lighting will almost
certainly be essential; see section 14.3.4 for details. Where possible, the lighting
hood should be in addition to a cover that reduces evaporation.
Water quality
As with a cold-water aquarium, tap water, suitably treated to remove chlorine,
will be adequate for many purposes. For certain types of tropical fish or when the
encouragement of breeding is important, there needs to be more precise regulation
of factors such as pH and hardness; this may, for example, involve the use of
sphagnum moss peat to acidify the water. When such action is thought appropriate, reference should be made to specialist texts on tropical fish. For general
information, refer to Table 14.1 in section 14.1.1.
Stocking the
aquarium with
animals
Follow the general advice given in section 14.3.1, particularly with regard to the
number of fish obtained for a particular size of tank. Section 14.1.1 makes some
suggestions for suitable species of fish but there are many other types that can
also be considered. In a ‘community’ tank, it is important to choose fish that will
live peacefully with others and which do not have unusual requirements such as a
low water pH or elevated temperature. It is customary to select fish that will exploit different regions within the aquarium, ie, ‘top’, ‘middle’ and ‘bottom’ feeders.
Always include a significant number of fish that browse on algae and scavenge
particles from the gravel surface; these will help considerably in maintaining a
healthy aquarium. A good local aquarist should always be able to advise on the
most-appropriate species currently available and reference to any comprehensive
manual on tropical fish keeping will provide the necessary information on the
habits of particular species.
If amphibians or aquatic reptiles requiring heated water are to be kept, useful
information can be found in The Care of Reptiles and Amphibians in Captivity by
Chris Mattison; see the reference in Table 14.1.
14.3.3
Marine aquaria
The establishment of an unheated or tropical marine aquarium is not an exercise to be
undertaken lightly; there are many difficulties and the financial outlay in stocking an
aquarium with tropical species can be extraordinarily large. The rewards, however, if
successful, are great; some of the most fascinating invertebrate and vertebrate
1430
© CLEAPSS 2006
animals are marine. Essential factors in achieving success include a high level of commitment and the expenditure of a considerable amount of time. Holiday periods, when
expert attention may not be available, are a particular problem.
Cold-water
marine aquaria
Organisms collected in rock pools on the seashore often do not survive for long if
kept in a salt-water aquarium at typical room temperatures. The long-term success of such an aquarium will usually demand some form of cooling of the water.
How this is achieved practically either involves considerable expense or the use of
d-i-y installations which may fail to meet appropriate electrical safety requirements. In most situations, therefore, laboratory observations of marine organisms
will involve only short-term studies with the animals returned to their original
environment as quickly as possible.
Tropical marine
aquaria
It is much easier to heat up a tank of water than to cool it down, so the establishment of a tropical aquarium is more practicable. Before proceeding, ensure that
your bank balance indicates a good deal of surplus cash and obtain expert advice,
preferably from someone who has achieved success. A tropical marine aquarium
can, however, often look stunning only because the animals are replaced at very
regular intervals!
The input of as much oxygen as possible and the removal of toxic wastes are vital
for a healthy marine aquarium. The build up of ammonia, nitrite and nitrate
needs to be monitored regularly and urgent action taken to reduce dangerously
high levels. A biological and/or chemical system for their removal, with water
passing over media that will extract or breakdown the toxins, should be adopted if
possible. The maintenance of a clean, sterile environment by the use of a protein
skimmer, ozoniser and/or ultra-violet steriliser is often a key factor in success but
may not be feasible in a school situation.
m 14.3.4
Accessories and electrical safety
The problem
The most common accessories include aerator pumps, heaters and thermostats, lamps
with their controls and power filters. They present particular hazards because of the
proximity of mains supplies and impure water. When installing aquarium accessories,
it is essential that schools follow local authority regulations. For a general discussion
of electrical safety, see section 6 (Mains electricity).
Electrical accessories intended to operate outside an aquarium must be sited so that
they do not fall in and as far away from the tank as practicable to avoid splashes.
Avoiding the problem
The simplest way of minimising the electrical hazards is to manage without these
accessories. As was explained in section 14.3.1, it is possible to:
a) rely on the atmosphere to oxygenate the water;
b) operate the aquarium at room temperature;
c) use daylight to provide lighting (by siting the aquarium appropriately);
d) rely on plants, snails and other organisms to maintain the environment.
An aquarium without accessories is, however, rather restricting.
Aerator pumps
There are two types of pump which are commonly used with school aquaria to oxygenate the water and also to operate filters of various types. Either should be positioned
so that water will not splash on it or siphon from the tank into the pump if it stops
working. If possible, the pump should be mounted on a shelf or wall above the
aquarium water level. If the pump has to sit on the bench near the aquarium, a
reliable non-return valve should be included in the air line between pump and tank.
© CLEAPSS 2006
1431
Hy-Flo pumps
Although these are no longer made, schools may still be using them. A rotor drives
reciprocating pistons. They are quiet, efficient and reliable provided they are
maintained according to the manufacturers’ instructions.
Vibrating diaphragm pumps
These pumps1 are often noisy, although noise can be reduced by suitable mounting. They require little maintenance.
Leads
These can be two-core or three-core and can enter the pump through a grommet.
In some designs, the mains lead hole is also the air inlet and the grommet will not
fit tightly round the lead. The lead should be fitted with a plug with a 3 A fuse.
Heaters and thermostats
Aquarium heaters and immersion thermostats can be sold legally only if the electrical
parts are enclosed within two walls or a single wall with a high resistance to impact
damage. Some designs also incorporate earthed parts and a fuse for extra safety.
While modern designs are very safe and are unlikely to present a serious hazard to
the user even when handled incorrectly, it is still essential to switch off the supply
before removing heaters from aquaria to prevent them overheating.
Older pattern
heaters
These have a single wall of ordinary glass. While it is not illegal to use them, they
should be replaced as soon as signs of trouble become apparent.
External
thermostats
These are unsuitable for use in permanent school aquaria because the control is
too easily tampered with. They do, however, have applications in other studies.
Combined
heater /
thermostats
One or more of these are preferred for most school tropical aquaria because they
are generally tamperproof and, if a separate mains socket is available for each
accessory, the only connections to be made are those into the mains plug. They
should be pre-set to the required temperature (normally around 24 oC for tropical
fish). Although these combined units are long (250 to 300 mm), the newer types
can be mounted horizontally and are then suitable even for shallow aquaria or
vivaria. (NB Some older types are only for vertical use.)
m Separate heaters
and thermostats
Connections for
a simple, one
heater, one
thermostat
system
These are sometimes required, perhaps for the use of existing equipment. In this
case, great care must be taken to make the connections well insulated and tamperproof. The arrangement illustrated below will achieve this. Wires twisted
together and covered merely by patent insulators such as ‘Scruits’ or insulating
tape are not satisfactory.
Terminal block
BR
BR
Mains supply
BL
BL
BR
BL
BL = Blue
BR = Brown
Heater
Thermostat
= Earth conductor
yellow and green
N.B. Any earth conductor from the incoming mains cable
should be securely connected to the terminal on the inside of the box.
1
There are many different models, that perform in a similar way, available from various companies.
1432
© CLEAPSS 2006
NB Suitable die-cast boxes are available from R S Components. Conduit boxes are
usable if holes are drilled for the grommets instead of using the ‘knock-outs’.
Earth connections have not been shown but are essential. The yellow-green
conductor from the mains supply should be connected to a tag on the box so that it
is earthed and to the conductor in the terminal block marked ‘E’ if the heater and
thermostat have earth (yellow-green) leads which should also be connected to this
conductor. The box should be mounted on the wall above and behind the tank.
m Hoods fitted with lighting
A sheet of transparent plastic, which allows sufficient daylight into the tank, may be
all that is required to reduce evaporation to acceptable levels; see section 14.3.1. In
many school applications, however, a hood fitted with a lamp or lamps will be required. Local authority regulations sometimes restrict the type of lighting hood that
can be used by specifying a particular construction or supplier1.
To prevent water vapour from condensing on the electrical components in a hood,
there must always be at least one transparent barrier between the water and the
lights. In some cases, a sheet of glass or Perspex is built into the hood that encloses the
lights and control gear for fluorescent tubes. This type of installation is to be preferred. With less well-protected fittings, there must be a cover over the water at the
top of the aquarium. Where possible, in addition to a transparent barrier forming part
of the hood, it is best to have a cover at the top of the tank; this can be made of plastic
with holes cut to allow for tubing and access for feeding. Ensure that condensation
drips back into the tank rather than outside it. Also be careful never to allow the
lighting hood to slip and fall into the tank.
Lighting is best supplied by fluorescent tubes that are sold in various types to promote
plant growth, reduce algal growth and show off particular colours of fish.
Advantages and These are preferred because:
disadvantages of a) for a given light output, they produce less heat than tungsten filament lamps;
fluorescent tubes b) the colour of the light is the most appropriate for encouraging the growth of
c)
d)
plants and for displaying the colours of tropical fish;
they are more economical than tungsten lamps of equivalent light output;
bayonet lamp holders, as commonly used with filament lamps, are particularly susceptible to corrosion.
Unfortunately, the installation cost is higher because fluorescent tubes require
control gear (eg, choke and starter) which is not needed for filament lamps.
No aquarium requires lighting all the time. Ideally, the lighting should simulate that
of the natural environment of the fish and plants; an 8 or 10 hour ‘day’ with abrupt
‘dawn’ and ‘sunset’ is, however, usually satisfactory.
Time control of
lighting
Jenno (see 14.3) discusses simple ways of changing the light level more slowly but
these may not be practicable in schools. In spite of the abrupt changes, school
aquaria probably benefit from simple on-off control by a time clock. Experience
will establish whether problems occur if fish are kept that are more nervous and
sensitive to sudden changes in light intensity.
Any time clock and control gear for fluorescent tubes can be housed in an earthed
metal box fixed to a wall behind the aquarium. This has the advantage that the
hood does not have to support a heavy choke and that the installation is more
tamperproof.
1
There are several manufacturers of lighting hoods; John Allan Aquariums has been found to supply a good range with a high
quality of construction.
© CLEAPSS 2006
1433
Water pumps and power filters
Rather than use an air pump to drive a filter, it is often better to use a small water
pump with an under-gravel filter or, instead, a power filter1 that pumps the aquarium
water through a compartment filled with a filter medium. These power filters are
more efficient than air-powered filters, silent in operation and are available in models
for use either inside or outside the tank; they are highly recommended but can be
more expensive. For pumps and power filters that are designed for operation immersed in water, it will be important to ensure that losses due to evaporation are
topped up so that the pumps do not become exposed.
m Other safety points
Tanks
Tanks should be all-insulated and the water not earthed.
Earthing of all
metal hoods
If an all-metal hood must be used, then it should be earthed.
Other
accessories
Other electrical accessories, such as sterilisers and ozonisers, should be constructed on the principles discussed above. Any accessible metal should be earthed or
double insulated. Any holes in the casing should be checked to see whether or not
they could allow water onto electrically live parts and, if so, such ingress of water
must be prevented. It seems cheaper to choose robust non-metal casings rather
than to double insulate metal ones. In all cases it is important to comply with the
manufacturer’s instructions on the correct mode of use.
Connecting
boxes
Reference has been made several times to the hazards presented by poor electrical
connections. A single aquarium control box or ‘console’ is attractive if it is properly
constructed but it is expensive; simpler systems will be adequate in many cases.
The most obvious system has a separate, switched socket for each accessory, in a
position safe from splashes and preferably above water level in the tank. The row
of sockets should be connected to the mains through a switched and fused spur.
If separate heaters and thermostats are used, then a connecting box as shown on
page 1431 will be required as well.
Summary of recommendations
Planned
installation
a)
The tank should be all-insulated and the water not earthed.
b)
The aerator pump should be protected from flooding. If its cable has to be
replaced, any earthing must be maintained.
c)
The heating should be provided by combined heater/thermostats of a design
that has double glass walls or single toughened glass or plastic walls.
d)
The tank should have a hood fitted with one or more fluorescent lamps and
built to a high standard of electrical safety.
e)
Any accessory should:
i)
be mounted so that it cannot fall into the water;
ii)
be of double-insulated construction or adequately earthed;
iii)
1
have its own mains plug (with appropriate fuse) and a separate
switched socket.
Various manufacturers supply water pumps and power filters but ‘Eheim’ is a particularly good range; the supplier is John Allan
Aquariums.
Existing
installation
14.3.5
1434
© CLEAPSS 2006
f)
If the power is supplied through a protective device such as an earth-leakage
circuit breaker or isolating transformer, often installed by a local authority
electrician, the system should be reasonably safe if properly maintained. Ask
the local authority electrical maintenance department for advice if there is
any doubt about safety.
g)
Where there is no special supply, the system should conform to recommendations in Connecting boxes above. If the lead on a heater or thermostat is too
short to reach a safely-positioned connecting box or socket, the whole unit
should be replaced.
h)
All external wires should have two layers of good insulation around the
metallic conductors. If any wires have only one layer of plastic or rubber
insulation, with or without a fabric cover, they should be changed.
i)
A metal hood/lighting unit should be checked to ensure that it is properly
earthed. If bayonet lamp holders are in use, they should be checked for corrosion and, if of metal construction, for earthing. A sheet of glass or Perspex
between the water and the lighting unit is needed to minimise problems.
j)
Any thermostat or heater showing signs of age or wear (eg, cracked rubber
bung, internal condensation etc) should be replaced.
k)
The aerator pump should be re-sited if liable to flooding.
l)
The aquarium tank should be examined for corrosion if it has a metal frame,
for cracks if it is plastic and for weeping seals if it is made of glass.
Maintenance of accessories
m NB Disconnect any mains-operated equipment before it is opened for servicing. All equipment with earthed metal parts should be checked annually
with an earth-bond and insulation tester.
Rotary pumps
The instructions issued with these pumps indicate that they should be oiled daily for
the first few days of use and thereafter at weekly intervals, depending on the amount
of use. The Hy-Flo pumps in use in schools benefit from such regular attention. Neglected pumps must, however, be cleaned before more oil is applied; see the next section
for comments on the type of oil that is suitable.
Routine
maintenance
There are two ranges of Hy-Flo pump; the ‘Technical’ range, painted black and
with pistons 7/16 inch diameter and the ‘Super’ range, painted gold, with pistons
3/16 inch diameter.
Hy-Flo pump
from the
‘Technical’ range
Crank pin
Piston
Cylinder
Valve block
© CLEAPSS 2006
1435
NB. Several of the oiling points mentioned below are holes packed with fibre to
hold the oil and to act as a wick. Do not attempt to remove this fibre although it
looks as if it is dust trapped in the hole.
Dismantling
On each side of the cylinder there are two screws holding the valve block to the
main body of the pump. Remove these screws. Gently pull on the valve block; the
crank pin will slide through the head of the piston allowing the piston, cylinder
and valve block to be removed together. If the pump is one of the ‘Technical’
range, the piston can be taken right out; this is not possible with the ‘Super’ range
pumps.
Cleaning
On the opposite side of the valve block to the cylinder can be found a spring which
presses the valve block and the cylinder together. These should be held apart and
a degreasing agent dropped from a teat pipette into the tubes on the valve block.
(1,1,1-trichloroethane used to be recommended for this task; as an alternative, try
Evolve CH15, Lotoxane or Volasil 244.) When these passages are clear, the surfaces of the valve block and the cylinder should be cleaned in the same way. The
crank pin and the hole through the top of the piston are then cleaned together
with, in the case of a ‘Technical’ pump, the piston and the inside of the cylinder.
Oiling
Reassemble the parts, taking care in the case of a ‘Technical’ pump that the piston
is replaced with the oiling point towards the front for easy access.
The pump must then be oiled. The manufacturer of Hy-Flo pumps recommended
sewing-machine oil, Shell Vitra 27 or its equivalent. 3-in-1 oil is too thick and
should not be used.
Oil should be applied to the holes in each of the following places.
a)
b)
c)
The main body of the pump above the axle of the flywheel (on both sides).
The valve blocks.
The pistons (‘Technical’ range).
Oil should also be applied on:
d)
e)
the crank pins where they pass through the piston head;
the piston rods (‘Super’ range).
If too much oil is applied, the slight ticking noise heard when the pump is running
will become louder. Little and often should be the rule for oiling. The frequency of
cleaning depends on the amount of dust in the air where the pump is used.
General points
a)
b)
c)
Water must not be allowed to enter these pumps; take great care attaching
the air lines if the pump is a type with a vacuum connection. If it is not
possible to site the pump above the water level in the aquaria, add non-return
valves to the air lines.
Even the smallest Hy-Flo pump will supply air for several diffusers. If several
are not required, fit an extra piece of tubing with a screw clip to allow excess
air to escape.
The electrical components of the pumps need no maintenance as such
although the earthing and insulation should be checked at least once each
year. The coils, and hence the body of the pump, become quite warm in use
but this does not necessarily indicate faulty operation.
Vibrator pumps
These require very little maintenance but occasionally the diaphragm may need
replacement and the rubber flap valves may need cleaning. Noise can be a problem
caused by vibrations in the surface on which the pump is standing or in adjacent
equipment. This can sometimes be reduced by standing the pump on a pad of a soft
material such as foam or felt or by sticking foam strips under the feet of the pump.
Electrical faults
1436
© CLEAPSS 2006
The only electrical fault that is likely to occur is a broken connecting wire. If no
vibrations occur when the pump is plugged into a live socket, disconnect from the
supply, open the plug and inspect the fuse (if there is one) and the connections. If
no fault is located, check the connections inside the pump, looking for a broken
wire. If still no fault is found, it is worth checking the lead before consigning the
pump to the dustbin. Remember that a new lead must have two layers of insulation.
Vibrator pumps are usually of ‘double-insulated’ construction and do not have an
earth connection. It is difficult for a school to check the insulation on such a unit.
Replacing the
rubber diaphragm
in a vibrator
pump
The most common cause of breakdown of this type of pump is a perished rubber
diaphragm. In this instance, the vibrations can still be felt but little or no air is
pumped.
A replacement diaphragm or, alternatively, one or two pieces of thin rubber such
as bicycle inner tube will be needed. Disconnect the pump from the mains and
remove the cover. Inside will be seen a small electromagnet with a vibrator arm
attached, together with a small cylinder with a rubber diaphragm clamped over
the top; the centre of the rubber is attached to the vibrator arm. An exploded diagram of the vibrator arm and diaphragm of a typical pump is given below.
Nut
Fibre washer
Vibrator arm
Fibre washer
Spring
Nut
Metal washer
Rubber washer
Diaphragm
Rubber washer
Metal washer
Screw
Remove the clamp and the old bits of rubber and release the vibrator from the
electromagnet. Dismantle the vibrator, laying the parts out carefully in the correct order so that it is easy to reassemble. As appropriate, cut the rubber to a
suitable size (often two layers are needed) and pierce a small hole in the centre for
the screw. Reassemble the vibrator with the diaphragm or rubber clamped between two washers.
Look inside the cylinder: you will see a small piece of rubber sheet that acts as a
flap valve to cover the air inlet during part of the pumping cycle. Carefully wipe
out any dirt.
Replace the vibrator, clamp the rubber diaphragm to the cylinder and replace the
cover.
© CLEAPSS 2006
Replacing a
flap valve
1437
Rubber flap valves last much longer than diaphragms because there is very little
mechanical stress on them. If the pump is very old and a new diaphragm does not
make it work properly, it is worth trying to install a new flap valve. It is not
possible to give details here, however, because models vary considerably in the
way that the valve is fixed.
Heaters
If a heater fails, or its outer tube breaks, it is usually inadvisable for a school to
attempt a repair. If it is fairly new, it can probably be repaired by the manufacturer
but, if it has been in use for over 12 months, it is probably best to replace it.
With some designs, the user can remake the water-tight seal; the instructions for
this must be kept until needed. (If the seal relies on a plastic strap, this may be
available from R S Components as a ‘cable tie’.)
Thermostats
If a thermostat has to be opened to make an adjustment, the power must be switched
off first and the bung must be replaced in its original manner before use. A good
thermostat will give reliable service for several years but some thermostats can fail
quite dramatically if the contacts corrode and arcing occurs, welding them together so
that the thermostat keeps the heater permanently on.
Cleaning the contacts is not recommended because this can remove a coating that
is applied to minimise the problem and, in addition, even a small movement of the
bimetal strip can upset the operation of the device. Most thermostats have a neon
indicator that flickers on and off periodically when the thermostat is working
properly. Observation of this indicator and monitoring the tank temperature can
provide a warning that the thermostat has reached the end of its life.
Lighting
At least once each term, the lighting unit should be checked to determine that the
wiring is in good condition and that water is not condensing onto the electrical
fittings. If the hood is metal, the earthing point should also be checked each term for
corrosion. The earth continuity should be checked with a test set at least annually.
Other powered accessories
At least once each term, the unit should be checked to see that the wiring is in good
condition and that the earthing is adequate (if metal-cased) by means of a simple test.
Bearings may also require a light application of lubricating oil; sewing-machine oil
will probably be of the correct type.
Protective devices
Where an aquarium installation is supplied through an isolating transformer, earthleakage circuit breaker or any special ‘black box’, the school should check with the
local authority electrical engineer whether any internal component requires maintenance. In most cases, the unit will require no more attention than the rest of the school
wiring.
If the unit has a push-button with a label ‘TEST OFTEN’, it probably contains an
earth-leakage circuit breaker. The button should be pressed weekly or even daily
to check that it causes the supply to be switched off. The supply should be restored
by operation of a RESET or ON / OFF switch nearby.
14.4
1438
© CLEAPSS 2006
Body fluids and cells - human
There may be several occasions during the teaching of science or biology when teachers may decide that it is appropriate for students to tackle practical activities involving the study of human body fluids: blood, saliva, urine and sweat. Work on cells can
be enhanced by activities using human cheek cells. There is, however, the potential for
the transmission of disease caused by bacteria and/or viruses present in the fluids.
Some employers, particularly local authorities, may have restricted the study of some
or all human body fluids. Such prohibitions may have been made following advice
issued in the past by the DES1 (for schools in England and Wales) or DENI1 (for
Northern Ireland). However, more recent guidance from the DfEE2 is less restrictive.
Schools must follow the requirements of their employers but for any restrictions still
in force that were made many years ago, it is worth exploring whether the employer’s
policy might be changed, in the light of current advice, discussed below.
Teachers working in foundation, voluntary-aided or independent schools and incorporated colleges are less likely to experience restrictions on practical activities in science
made by their governing bodies (which are the employers in these establishments)
than teachers in local-authority community and voluntary-controlled schools. If
science departments do not have, in writing, ‘bans’ sent out or confirmed recently by
their employers, they should assume that there are no restrictions. Teachers often
believe that a ban exists, preventing them studying some or all body fluids /
cells, when in fact none has ever been issued. In fact, in a recent CLEAPSS
survey3, over 80% of responding local authorities reported that they did not ban or
discourage pupils taking their own blood samples, while over 90% stated that they had
issued no restrictions on taking cheek cells or the use of saliva.
As required by the COSHH and Management of Health & Safety at Work Regulations,
teachers must, before starting work on human body fluids or cells, assess the risk of
using such fluids/cells with a group of students and then adopt appropriate procedures
and precautions to ensure health & safety. Naturally, if a teacher cannot be reasonably certain that a particular class of students will behave themselves during work
with body fluids/cells, therefore possibly causing infections to be transmitted, such
activities would not be allowed to proceed. The information presented in this section of
the Handbook provides model risk assessments to be used in this process.
14.4.1
Blood
m When human blood, plasma or red cells from any source are handled, there is a slight
risk that viruses (or bacteria) may be transmitted. The most significant of these are
the human immunodeficiency virus, HIV, (the cause of AIDS) and hepatitis viruses B
and C. The risks of transmission of diseases that can affect humans from the blood of
mammals other than humans are considerably lower. Human blood-borne viruses are
only transmitted if blood from a carrier or infected person infects another person via,
say, a scratch in the skin. There is no significant risk if the correct sterile procedure is used but an employer may not be sure that this would always be followed. A
suitable sterile procedure to be used when obtaining and handling blood samples is
given later in this section.
1
2
3
For example, in AIDS, Some Questions and Answers (Department of Education and Science, 1987), in answer to the question “Is
blood sampling in class safe?”, the response was given: “No. Taking blood and cell samples for science demonstrations could
carry a risk; therefore this practice should be discontinued.”. Safety in Science Laboratories (Safety Series no 1, Department of
Education for Northern Ireland, 1989) includes the statement “No body fluids should be used in the laboratory including blood
from any source.”. It should be noted, however, that the DfES (or former DES) has never issued official advice warning against
activities using urine, saliva or sweat.
In Table 17.14 of Safety in Science Education (DfEE, HMSO, 1996, ISBN 011270915X).
Surely That’s Banned? A report for the Royal Society of Chemistry on chemicals and procedures thought to be banned from use
in schools, October 2005. The report and appendices can be accessed on the RSC web site: www.rsc.org/education/ and a copy
of the report has been sent to all schools.
© CLEAPSS 2006
1439
Why study blood? In the study of blood, it is important that it is shown to be a composite liquid con-
taining several different components including red cells, white cells and plasma.
Aspects of physiology, including the determination of blood groups, clotting time
and measurements of blood sugar levels, are also very valuable and suitable
subjects for practical studies1. Most students are fascinated by such work. They
are eager to look at blood smears under the microscope, investigate blood groups
and so on. However, there are difficulties in obtaining blood samples, even the
drop or two normally required for such activities.
Blood from different sources
The use of blood from other mammals (henceforth called ‘mammalian blood’) is less
hazardous than human blood; mammalian blood will be used if larger quantities are
required for the practical activity. Time-expired human blood, if samples can be
obtained, will be less hazardous than blood samples taken from students or staff. Both
these sources of blood can be used for several of the activities described in Investigations with blood below. However, such blood samples are somewhat limited in their
usefulness because they do not permit personalised studies of some aspects of physiology, eg, blood groups or glucose levels. For these, only finger-prick samples of fresh
human blood will be satisfactory.
Mammalian
blood
Like other mammalian tissue, this may be a potential source of infection and it
should be handled with reasonable care; the risks are much the same as those of
handling meat in the kitchen. Mammalian blood can usually be obtained from an
abattoir if one is conveniently located (see Yellow Pages) and can be purchased
from Blades Biological, Timstar and some other suppliers.
Occasionally, schools that have contacts with a vet may be able to obtain blood
samples taken from a mammal such as a horse or dog that is being treated. In
such cases, it is important to ascertain that the health of the animal is such that it
has not made the blood sample more hazardous to handle.
Note that the ‘blood’ which drips from meat such as liver bought from a butcher is
likely to be plasma contaminated with haemoglobin from burst red cells.
Anticoagulants
Mammalian blood must be treated to prevent coagulation as soon as possible after
collection. Purchased blood is either defibrinated, removing the protein that causes the blood to clot, or an anticoagulant, usually oxalate, has been added. If blood
is to be collected from an abattoir, take a bottle already containing anticoagulant2.
(Note: it is prudent to label the bottle “contains anticoagulant”; it has been reported that the contents of the bottle may be rinsed out in the abattoir before the
blood is added!)
Treated mammalian blood can be stored in a refrigerator for 2-3 weeks and disposed of as for time-expired human blood; see below.
Time-expired
human blood
Sometimes time-expired blood can be obtained from a local blood bank. A contact
in a hospital may be able to help. Because this blood will have been screened, it
will be free of the HIV virus but, as the screening for hepatitis is not 100% effective, sterile procedures should be adopted (see below). (Precautions 2, 3, 4, 10, 11,
12, 13 and 14 of the suggested sterile procedure will apply if time-expired blood or
red cells are used.) After use, autoclave the blood samples before pouring down a
drain or toilet, followed by plenty of water.
1
It is not easy to find details of practical activities with blood in books that are currently in print. A useful reference for various
investigations with blood is Revised Nuffield Advanced Biology, Practical Guide 1: Gas Exchange & Transport in Plants and
Animals, Chapter 4, and the accompanying Teachers’ Guide I, pages 95-101 (Longman, 1985). If these materials are not available, CLEAPSS should be able to provide assistance.
2
Di-sodium hydrogen citrate is the preferred anticoagulant. A suitable solution contains 1.7-2% plus 2.5% dextrose (glucose):
120 ml of this prevents the coagulation of 420 ml of blood. Alternatively, use 10 g of sodium citrate in 100 ml of water for 1 litre
of blood. Another anticoagulant can be prepared by mixing 3 parts of ammonium ethanedioate (oxalate) with 2 parts of
potassium ethanedioate (oxalate); use 5 ml for every 100 ml blood.
Fresh human
blood
1440
© CLEAPSS 2006
If the employer has not banned taking blood samples from students and/or staff,
teachers may decide, after a full risk assessment, that it is safe to take fingerprick samples of human blood under sterile conditions, to be used for exciting
practical activities with some of their students. After all, blood is frequently shed,
in far from sterile conditions, during school sports activities without any outcry!
Teachers should ensure that students fully understand the precautions that must
be taken and the consequences of not taking them. This is valuable for their
general education as well as essential for blood sampling to be safe and healthy.
HIV and hepatitis Any students or staff who know that they are HIV-positive or have tested positive
for hepatitis B &/or C viruses should not give blood samples. Procedures should
permit affected students to be excluded from, or allowed to opt out of, the sampling activity without having to admit publicly that they have tested positive for
these viruses. Confidentiality should, of course, be preserved at all times; teachers
need skill in managing such situations. As many as 2-3% of students in some
areas may be hepatitis B &/or C positive but their identity in a group will often
not be known to teaching staff. It should be made clear to students that they
should not take part if they have good reason to believe they may pose a particular
risk to others. At the same time, however, the teacher should allow students to
decline to take part without in any way drawing attention to any possibility of
infection; see below.
Student
participation
There must be no pressure on a student to provide a blood sample. Teachers
should make it clear by their attitude that it is perfectly normal for some students
not to want to provide a sample and not to want to take part in the practical work.
If this is done well, it is likely that such students will gradually become involved
in the work. Students should be allowed to change their minds either way.
Parental
permission
Unless specified by an employer’s policy, there is no legal requirement to obtain
the permission of parents or guardians for their children to provide blood samples.
However, it would certainly be prudent to do so. A suitable letter that could be
used as the basis for communications with parents is included in the ‘Customisable documents’ section of the CLEAPSS Science Publications CD-ROM; code
DLH14-4.
Who takes the
blood samples?
It is recommended that the teacher should supervise the activity closely but that
students take their own samples of blood. In this way, there can be no accusations
made later of assault by the teacher. However, students who are quite willing and
eager to provide a sample may find it difficult to prick their own finger with a
lancet. This is particularly likely if a standard lancet is used which requires the
student to summon the courage to stab him/herself (and the tip of the lancet could
penetrate the skin to a depth of a couple of millimetres).
The use of an automatic finger-pricking device1 is therefore strongly recommended. The teacher is responsible for assembling the instrument with a fresh lancet
and then hands it to the student who simply presses the device onto the skin and
triggers the lancet. The instrument is handed back to the teacher who then disposes of the lancet safely (see the sterile procedure for blood sampling below).
If, for some reason, the teacher does take a blood sample from a student, it is
recommended that teachers ask students to sign to indicate that they have given
permission for this to occur. A suitable consent form is included in the ‘Customisable documents’ section of the CLEAPSS CD-ROM as part of the document
DLH14-4.
1
Various automatic lancing devices have been developed to help diabetics sample their blood as painlessly as possible. Several
companies sell suitable devices for school use including those from Owen Mumford (Unistix 3 and Autolet Impression) and
Accu-Chek (Softclix). The Softclix Pro Finger Pricker is designed for professional multiple use and ejects the lancet after each
sample is taken. Local pharmacists may be a suitable source for purchase and there are also several internet suppliers including
www.mypharmacy.co.uk, www.expresschemist.co.uk, www.spservices.co.uk and www.westons.com. The mail order division of
Owen Mumford is www.medicalshop.co.uk.
Lancets for automatic lancing devices are manufactured in a range of gauge sizes; for example 21 gauge lancets have thicker
needles for producing maximum blood flow, while 28 gauge lancets are finer for greater comfort. Note that automatic lancets are
often sold as part of a kit including digital glucose monitors.
© CLEAPSS 2006
1441
Suitable sites for Blood should be taken with a sterile lancet from the side of a finger, near the nail,
using a new lancet for each person. It is not recommended that blood is taken
taking blood
from a finger tip because of the greater thickness of the skin at this point and the
samples
risk of subsequent infection. The ear lobe has sometimes been suggested as an
alternative site but this is also not recommended because of the danger if a
student jerks his or her head as the sample is taken and the difficulty of transferring drops of blood for investigation.
The best position is 5-10 mm from the lower corner of the nail (see diagram). It is
easier to insert the lancet if the finger has been crooked at the top joint.
To help ensure that sufficient blood will flow from the punctured skin, the hand
should be warm (so encouraging blood flow to the skin). It is sometimes helpful to
force blood to the extremities by vigorously shaking the hand or rapidly moving
the arm in a circle around the shoulder joint (take care to ensure that the arm
cannot hit anything or anyone!).
Care with lancets Teachers must supervise the issue, use and subsequent disposal of the lancets
(using a ‘sharps’ container) extremely carefully. Relatively inexpensive small
sharps containers are available from school science suppliers, eg, from Scientific &
Chemicals Supplies and Philip Harris; their use is preferred over d-i-y sharps
containers. If necessary, however, equivalent sharps containers could be constructed; see the information in section 8.1.1 (Solid waste) on page 809.
Schools generally already have arrangements for the disposal of clinical waste
from first-aid rooms etc. It is likely that the sharps container could be disposed of
via this route at no extra cost.
Spills of blood
Spills of blood, however small and from any source, should be wiped up at once
with a cloth soaked in freshly-prepared sodium chlorate(I) (hypochlorite) disinfectant containing 10 000 ppm available chlorine; see section 14.13.
Taking and using human blood samples safely
A sterile procedure must be adhered to for all work with human blood. If the procedure1 in the box overleaf is followed rigorously, risks of disease transmission are
eliminated. When a human blood sample has been taken for investigations, it should
be used only by the person providing the sample. In this way, risks of cross-infection
are again eliminated.
m Investigations with blood
Demonstrating
effect of gases
on blood
1
The effect of mixing different gases (air, oxygen, carbon dioxide, carbon monoxide)
on the colour of blood is instructive. Using mammalian or time-expired human
blood, gases can be passed through the blood in an enclosed apparatus (eg, wash
bottle) and then bubbled through water to avoid the release of any aerosols2 into
the atmosphere. An alternative method is to draw 2 ml of mammalian or timeexpired blood into a 20 ml syringe through a length of tubing. The syringe is
swirled to obtain a thin film of blood on the inside of the barrel and gases are then
drawn into the syringe, most conveniently from a cylinder for oxygen and carbon
dioxide. Car exhaust fumes or cigarette smoke could be a source of carbon
monoxide; take great care in collecting exhaust fumes! Strong plastic bags could
be used to collect the gas samples and these attached to the syringe, the nozzle of
which is then capped or sealed off with a clip on the tubing.
This has been approved by a leading virologist.
1441A
© CLEAPSS 2006
Sterile procedure for taking and using human blood samples safely
Before the lesson
1. Slides or any other glassware that might come into contact with the site from which a blood sample is
taken should be sterilised in an autoclave or oven; see section 15.12.2 (Heat sterilisation).
2. A suitable disinfectant, able to kill viruses, should be freshly prepared. The recommended disinfectant is
a solution of sodium chlorate(I) (sodium hypochlorite) containing 10 000 ppm available chlorine [IRRITANT]. [This can be obtained by preparing a 10% dilution of a laboratory solution of sodium chlorate(I)
containing not less than 10% (100 000 ppm) available chlorine [CORROSIVE]. Note that domestic hypochlorite (bleach) solutions have already been diluted, often by an unspecified amount. It is difficult to
make up accurate dilutions using such sources of the chemical.] VirKon is less suitable for use with
blood because it has to be used as a powder rather than a solution.
During the lesson
3. Because of the risk of contamination through broken skin, the participation in this practical work of
anyone with any open wound, particularly on or near the face or hands, should be strictly limited;
depending on the nature and position of the wound, the student may need to be excluded from the work
altogether.
4. Students and teachers must thoroughly wash hands using soap and water. Those providing blood
samples should pay particular attention to washing the site chosen for the sampling. Dry hands using
only disposable towels or by other hygienic means.
5. Using a cotton wool swab, wipe the chosen puncture site with 70% alcohol [70% v/v, propan-2-ol
(isopropanol) or ethanol] and allow it to dry.
6. Immediately prior to its use, take a new sterile, disposable lancet, fit it into the lancing device, if one is
being used, and detach the cap over the lancet tip. Handle it carefully and do not allow the sharp tip to
touch anything.
7. Puncture the skin of the chosen site using the lancet and immediately place the lancet into a ‘sharps’
container (see discussion on Care with lancets above), small enough to fit in an autoclave if a d-i-y
container is being used. Lancets must be used once only.
8. Collect the blood by letting a drop or two fall into a sterile tube or onto a sterile slide or sterile rod (see
1). There must be no contact between the area of the pinprick and any apparatus unless the apparatus
has been sterilised.
9. Apply a sterile gauze dressing to the puncture site and press gently until bleeding has stopped. Once
blood flow has stopped, place the dressing in the container used for the lancets or an autoclavable
disposable bag.
10. Any blood spilt on the bench etc must be wiped up at once using the freshly-prepared disinfectant (see
2). Hold the swab with forceps or wear nitrile, rubber or plastic gloves.
11. The greatest care must be taken to avoid contamination of the skin with blood from another person. If
this should occur, however, the contaminated area must be washed thoroughly with soap and water.
12. When students have finished with the slides and any other contaminated glassware that will be reused,
these should be placed in a discard jar of the disinfectant referred to in 2 but diluted to produce a
solution containing 25 000 ppm available chlorine. Note that sodium chlorate(I) is rapidly inactivated by
the presence of organic matter, including blood. Sharp items for disposal should be placed in the
‘sharps’ container with the lancets. Non-sharp items, (eg, blood-grouping cards) should be placed in the
disposal bag with the swabs and dressings.
13. At the end of the practical, wash hands again using soap and water and dry thoroughly using disposable
towels or other hygienic means.
After the lesson
14. The disposal bag with the contaminated swabs etc should be closed, not sealed, and autoclaved,
together with the slides and other contaminated glassware from the discard jar and a d-i-y ‘sharps’
container. After autoclaving, the disposal bag should be sealed and, together with the ‘sharps’ container,
placed in a black plastic bag and placed into normal refuse. A purchased sharps container should be
sealed and disposed of with other clinical waste that is collected from the school. Autoclaved slides, etc
can be washed for re-use, (particular care must be taken in handling any coverslips, which are a
common cause of cuts). Alternatively, if glassware has not been contaminated by too much blood, it
could remain in the discard jar of disinfectant overnight before being washed in the normal way. Gloves
should be used to protect the skin from the disinfectant.
© CLEAPSS 2006
1441B
Carriage of
oxygen
It is possible, though not always easy to obtain accurate figures, to measure the
amount of oxygen that is carried by haemoglobin in blood samples. The technique
involves taking a measured sample of blood (eg, defibrinated or anticoagulanttreated) that has been oxygenated and adding potassium hexacyanoferrate(III)
solution; this causes the haemoglobin to release all its oxygen. The oxygen is collected and its volume measured, either using a J tube or some form of respirometer. Details of the procedure are given in the references quoted in the footnote1.
Centrifuging
blood
This will show the relative proportions of cells and plasma; centrifuge at 20002300 g for 30 minutes. Use only plastic centrifuge tubes that are designed to be
capped. Uncapped tubes or d-i-y attempts to cover uncapped designs should be
avoided as there is a risk of infective aerosol2 formation if blood is released in the
centrifuge when tubes break or loose-fitting caps come off. Separation of blood
components may be achieved by allowing a sample to stand for several days in a
stoppered tube.
Catalase activity Most tissues use catalase to destroy the dangerous hydrogen peroxide that is pro-
duced as a waste product during metabolism, and blood is no exception. It is,
however, inadvisable to use blood as a source of the enzyme in catalase studies if
they are carried out in a way that permits the considerable frothing which will
occur to release aerosols into the air. Open vessels must not be used. Liver bought
from a butcher will be a more manageable source of catalase.
Osmosis and
red blood cells
Although red cells can withstand a high water potential, the importance of maintaining a relatively constant concentration of blood in the body can be shown by
osmotic studies. One method is to use a drop of blood on a microscope slide, add a
coverslip and then place drops of distilled water and various concentrations of
saline solution at the edge of the coverslip so that mixing with the blood occurs.
There has been reported success in using blood extracted from a herring obtained
from a fishmonger for studies of haemolysis and crenation in red blood cells. Blood
can be extracted from a major blood vessel using a syringe and hypodermic needle
and may be diluted with 1% saline if larger volumes are needed for easier handling. If, on microscopical examination, the red cells differ in size or shape from
normal, the concentration of the saline may need slight adjustment.
Blood smears
The cellular components of blood can, of course, be observed using prepared slides
from a laboratory supplier. These stained preparations are most convenient but
lack the impact and immediacy of students making for themselves a smear of
fresh or citrated blood and then staining it.
The technique of making a blood smear on a microscope slide shown overleaf is
described in older texts, for example, in the Nuffield Biology materials already
referred to. A small drop of blood is placed at one end of a slide. A second slide is
held so that its edge stands diagonally across the lower slide at an angle of 60°
and it just touches the front of the drop of blood. The upper slide is then pushed
quickly along the lower slide so that the blood is dragged into a thin film. Make
sure that the blood does not reach the edges of the lower slide.
1
If aerosols are formed, a fine, invisible ‘mist’ of liquid droplets is released into the air; this can easily be inhaled and may be
carrying infective microorganisms.
2
The Nuffield Biology reference in the footnote on page 1439 describes the J-tube technique. Two respirometer techniques are
described in the following articles: Payne P F C, Estimation of oxygen content of blood, School Science Review, 58 (203), 1976,
p253 and Guilbert G, The carriage of oxygen by blood, Journal of Biological Education, 16 (1), 1982, p19. The respirometers
discussed use 3-way taps; see section 10.9.5 for details of where to buy these.
1441C
© CLEAPSS 2006
The slide is waved in the air to dry the smear. To see white cells and platelets
clearly, the preparation must be stained. Leishman’s stain is traditionally used
but some prefer Giemsa’s stain. After staining1, the preparation is examined
under the microscope; the best area of the smear to view the cells should be a little
way back from the end of the smear.
Blood grouping
Blades Biological, the US company Carolina (via its agent in the UK, Blades
Biological), Philip Harris, Scientific & Chemical Supplies and Timstar all sell kits
that use synthetic blood to simulate the expected results when typing blood
groups. Carolina also sells kits using aseptic samples of red cells of known blood
group that are then mixed with anti-sera. These kits of course do not allow
students’ own blood groups to be identified.
Students find work identifying their own blood groups quite fascinating and are
usually willing to provide a finger-prick sample of their blood for this or other
purposes. It should be emphasised to students, however, that blood grouping performed in school cannot be relied upon to be accurate for medical purposes.
Antisera A and B allow the ABO blood groups to be investigated. Anti-D serum
allows the Rhesus blood group to be determined. To carry out a blood-grouping
test, a drop of fresh blood can be placed on a microscope slide and a drop of the
appropriate antiserum added. If the red cells agglutinate (clump together), this
indicates a positive result. Thus cells of blood group A and B agglutinate with
antisera anti-A and anti-B respectively, to identify blood groups A, B and AB.
Blood group O causes no agglutination with either antiserum. Rhesus-positive
blood agglutinates with anti-D serum. The relative frequency of each blood type
differs from one population to another. In the UK, the proportions for the ABO
groups are as follows: O = 46%, A = 42%, B = 9%, and AB = 3%. 85% of the
population is rhesus-positive.
Antisera can be purchased from the US company Carolina (via its UK agent
Blades Biological), in 4 ml / 50 ml dropper bottles or dried onto Eldon cards. For
these cards, a drop of water is mixed into the dried antiserum to reconstitute it.
The cards are sold individually (and rather uneconomically) or in sets of 25 or 30.
There are companies in the UK which supply antisera for the National Transfusion Service and some of these2 will sell antisera in small quantities to schools.
Clotting
mechanism
Clotting time can easily be investigated by drawing up fresh blood into a finely
drawn-out capillary, perhaps best made by strongly heating a length of glass
tubing and pulling the two ends apart. At timed intervals, the tip of the bloodfilled capillary is broken off using forceps and the time noted when strands of
clotted blood are observed. Wear eye protection for this activity and ensure that
all pieces of capillary are placed immediately into a sharps container for subsequent autoclaving/disposal.
1
Place 5 drops of Leishman’s stain on the blood smear and leave for 1 minute. Add 5 drops of distilled water buffered to pH 6.66.8 and leave for 5 minutes. Wash the slide in 50 ml buffered distilled water to which a few drops of Leishman’s stain have been
added until the film becomes a rosy-pink colour. Press lightly with filter paper to remove the water and wave in the air to dry.
[To prepare 100 ml of buffered water, mix 56 ml potassium dihydrogen phosphate solution (9.078 g per litre) with 44 ml of a
solution of disodium hydrogen phosphate.12 water (11.876 g per litre).]
2
Alpha Laboratories (orders can be placed online at www.alphalabs.co.uk), Biotest (UK) and Lorne Laboratories. Most supply
10 ml quantities (30 ml from Alpha Labs) of anti-A, anti-B and anti-D sera. Contact CLEAPSS for details of the catalogue
numbers of antisera and the various costs from each of the suppliers including Carolina via Blades Biological.
© CLEAPSS 2006
1441D
Nuffield Biology suggests the use of a Dale & Laidlaw tube - is a length of narrowbore tubing with a small ball bearing inside. A tube is warmed to 37 °C and tilted
repeatedly. The time for the ball bearing to become immobile represents the
clotting time. Dale & Laidlaw tubes cannot, however, easily be purchased and
technicians are unlikely to have a readily-available supply of small ball bearings
with which to construct their own tubes.
Blood sugar
levels
The product Dextrostix was developed to provide semi-quantitative measurements
of the amount of glucose in a drop of blood. The colour that developed on the end
of the Dextrostix strip was compared with a colour chart.
Driven by the need for diabetics to monitor their blood sugar levels, there are now
several types of inexpensive glucose meters on the market. Each uses its own
brand of test strip onto which a drop of blood is placed. The colour that develops,
and therefore the amount of glucose that is present, is measured by inserting the
strip into the meter which gives a digital display. These meters can be purchased
from a local pharmacist or from the companies that sell lancing devices referred to
earlier.
With these meters there is ample scope for investigations of glucose levels in
students’ blood and whether the sugar level is well controlled after, for example,
consumption of a sugar-laden soft drink or a bout of exercise.
Blood cholesterol Relatively-inexpensive test kits are also available to monitor blood cholesterol. No
attempt should be made to make a medical diagnosis and it is not usual for people
levels
aged less than 20 years to be monitored for cholesterol levels. Nevertheless, there
may be value in bringing such kits to the attention of students and there is certainly scope for investigations of the way in which cholesterol levels vary from
person to person and whether they are affected by diet and exercise.
14.4.2
Cheek cells
As discussed in the introduction to section 14.4, the DES originally advised1 against
the taking of cells from the inside of cheeks. This prompted some local authorities to
ban the taking of cheek cells. However, more-recent advice from the DfEE2 overturned the earlier guidance, stating that cheek cells may be sampled, if a safe
technique is used. Most local authorities now permit cheek-cell sampling. Students
should sample their own cheek cells, only examine their own samples and be
responsible for the disposal of their own materials. If, following a risk assessment of the behaviour of students who will provide cheek-cell samples, the technique
described below can be adhered to, there will be no risk of the transmission of disease.
Safe sampling of 1. Take a cotton bud from a newly-opened pack.
cheek cells
2. Move the cotton bud over the inside of the cheek on one side of the mouth and
along the outer lower side of the gum.
3. Smear the cotton bud over a small area of a clean microscope slide.
4. Place the used cotton bud immediately into a small volume of disinfectant3 in
a suitable container, (eg, 5 cm3 in a 10 cm3 specimen tube) or into a bag or
beaker for subsequent autoclaving.
5. Add 0.1-1% methylene blue stain from a dropper pipette onto the smear and
cover with a cover slip.
1
AIDS: Some Questions and Answers, DES, WOED, 1987.
2
In Table 17.14 of Safety in Science Education (Department for Education & Employment, HMSO, 1996).
3
Use sodium chlorate(I) (hypochlorite) solution (containing at least 1000 ppm available chlorine; see CLEAPSS Recipe
Card 62) or 1% VirKon solution. See also section 15.12.3.
1441E
© CLEAPSS 2006
6. Observe the smear under the low-power magnification of a microscope. When
the cells are in focus, increase the power of the objective to achieve maximum
magnification and resolution.
The cytoplasm will be stained pale blue and the nucleus will be stained a
darker blue.
7. After the cells have been observed, immerse the slide and cover slip in a beaker of disinfectant3 or place them into a beaker for autoclaving.
8. After at least 15 minutes disinfection, the used cotton buds should be transferred, while wearing gloves, into a polythene bag that is sealed and then
disposed of into normal refuse. Alternatively, autoclave before disposal.
9. After at least 15 minutes disinfection, slides (and possibly cover slips) should
be washed thoroughly, dried and re-used according to normal practice.
Alternatively, the slides and cover slips are autoclaved before washing up.
Alternatives to
cheek cells
For the few schools that have recently been sent, or had confirmed, in writing a
local authority ban on students sampling cheek cells, there are some alternatives.
Using a trachea of a pig or a lamb from an abattoir / butcher, epithelial cells can
be scraped with a spatula and stained as above; these are often easier to see than
human cheek cells. Liver cells can be sampled by scraping the surface of a portion
of fresh liver bought from a butcher. With this technique1, it is claimed that cells
with more than one nucleus are observed. If Sellotape is applied to a well-washed
wrist, removed and stuck onto a microscope slide, cells with nuclei may be visible.
14.4.3
Saliva
The DES, DfEE or DfES has never recommended that the use of saliva in practical
activities should cease though a few employers have made restrictions on the use of
human body fluids. Where schools have been obliged to use alternatives to saliva for
amylase investigations, there have been reports of science staff becoming sensitised to,
for example, diastase when insufficient care has been taken to avoid inhaling dust
from a powdered enzyme preparation. In such cases, the careful use of saliva would,
paradoxically, have been less hazardous.
Because saliva can spread infections such as colds and sore throats, proper hygiene
must be observed. In fact, the use of saliva provides a good opportunity for teaching
hygiene. Students should use only their own saliva samples and be responsible for rinsing their own equipment. They should be provided with small beakers
or disposable cups into which they are asked to spit their saliva samples.
The use of saliva has other major advantages over prepared amylase solutions.
• It saves money.
• It frees technicians from spending time making up amylase solutions and testing
them to make sure they work effectively.
• Students are more likely to be interested in studying their own mouth secretions
than a chemical solution.
• The catalytic effect of sodium chloride on the speed of action of salivary amylase is
often discovered accidentally when students’ samples, taken after eating crisps or
salted peanuts, perform unexpectedly rapidly.
• Salivary amylase is generally a more reliable enzyme to study than purchased
amylases. Most of these contain sugars, so monitoring the production of sugars
when starch is digested becomes a pointless exercise.
1
See Burton I J, A simple technique for preparing liver cells for microscopical examination, Journal of Biological Education,
33 (2), 1999, p113.
© CLEAPSS 2006
1441F
• In studies of the influence of temperature on amylase activity, the optimum is
more easily identified with saliva as the source than with some purchased
amylases that have an unusually high optimum which students do not predict.
Salivary amylase denatures when it is boiled; this may not occur with amylase
from other sources.
• Some students do not produce amylase in their saliva; this can be used as an
example of variation.
• Lipase is also secreted in saliva, together with lysozyme that destroys some
bacteria, providing further opportunities for investigatory work.
Cleaning of
glassware etc
Even though students should be asked to rinse out glassware etc after practical
work, items contaminated with saliva should always be placed directly into a
vessel already containing an excess of freshly-prepared disinfectant, such as hypochlorite1 and left for 30 minutes before it is washed with hot water and detergent.
This should help to reassure technicians that the use of saliva in class practicals
will not endanger their health.
Beware of
aerosols
During tests on saliva samples within the laboratory, care should be taken to
avoid the formation of aerosols. These are microscopic, airborne droplets of fluid,
created when the surface of a liquid is disturbed in some way. If the saliva
contains infectious microorganisms, these will be carried by the aerosol droplets
and could spread around the laboratory.
Other precautions After work with saliva, it is prudent to wipe benches with disinfectant and make
sure that students wash their hands.
Mouthpieces,
clinical thermometers etc
14.4.4
These should be used only once and then placed in disinfectant for the appropriate
time to ensure disinfection. Milton is recommended for items that will be placed in
the mouth but note that this requires 30 minutes disinfection time. The use of
ethanol is quicker (5 minutes) but leaves an unpleasant taste. Disinfected items
should be rinsed before being used again. See section 15.12.3.
Urine
A typical reason for wishing to study urine is in the investigation of sodium chloride
excretion2. Urine is normally sterile but precautions similar to those when using
saliva should always be used.
Each student is responsible for providing his or her own sample of urine and should
not handle or test other samples. Students should be provided with appropriate containers in which urine samples can be collected and transported. The containers should
be made of plastic or strengthened glass with an effective means of closure so that
there is no risk of breakage or a spill during transit.
Aerosols
As with saliva, care should be taken to avoid the formation of aerosols when
handling urine samples.
Disposal and
cleaning
After tests are completed, students should hygienically dispose of all samples
containing urine, flushing them to waste using a toilet. They should then rinse
contaminated glassware. All items of apparatus that have come into contact with
the urine samples should be subsequently sterilised by autoclaving or, as for
equipment used with saliva, immersed for 30 minutes in a solution of sodium
chlorate(I) or 10 minutes in a 1% solution of VirKon disinfectant. Benches should
also be wiped with disinfectant and students should wash their hands.
1
Use sodium chlorate(I) (hypochlorite) solution (containing at least 1000 ppm available chlorine; see CLEAPSS Recipe
Card 62) or 1% VirKon solution. See also section 15.12.3.
2
For example, as described in Revised Nuffield Advanced Biology, Practical Guide 3: Cells, tissues and organisms in relation to
water, chapter 10 and Teachers’ Guide I.
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© CLEAPSS 2006
It is anticipated that investigations of urine samples for chloride ion content will
normally occur in Years 12 and 13 but studies of other urine contents might be tackled
by more junior pupils, if an assessment of risks indicates that work can be conducted
safely with all necessary precautions adopted.
Where the objective of the investigation is merely to show the presence, absence or
relative amounts of substances, it is preferable to use a simulation of urine samples.
This also avoids any problems if tests are carried out on actual samples of urine and,
from the results obtained, students believe that they may be suffering from some
illness. Tests performed in schools may not be sufficiently reliable and should not be
used for medical diagnoses. The use of Bayer diagnostic reagent strips1: Clinistix (for
glucose), Albustix (for protein) or Uristix (for both) is convenient and representative of
simple medical diagnostic checks.
Table 14.4 gives suggested compositions of body fluid analogues.
Table 14.4
Blood plasma
(renal artery)
Blood plasma
(renal vein)
Urine
Urine
(diabetic)
50 mg
trace
2g
2g
Glucose
100 mg
100 mg
-
500 mg
Albumen
7g
7g
-
-
Chloride
300 mg
50 mg
600 mg
600 mg
Urea
Water
Colour2
14.4.5
Body fluid analogues
100 ml
v. pale yellow
100 ml
v. pale yellow
100 ml
deep yellow
100 ml
deep yellow
Sweat
This body fluid is perhaps less likely to be a particular focus for practical activities
than those already discussed. Suggested investigations of the secretion of sweat
involve the application to the skin of pieces of cobalt chloride or thiocyanate paper.
CLEAPSS advice is given on the reverse of Hazcard 25; it is recommended that the
skin should be washed after the papers are removed. One curriculum project has
suggested making measurements of the amount of sweat secreted by attaching preweighed tampons under the armpits during a bout of heavy exercise! Such activities
obviously require good hygiene precautions and disposal, as discussed above.
14.5
Breathing investigations
m Pupils
allowed to take part should be selected carefully; if any unusual breathing
occurs, it might be harmful to anyone with a bronchial condition such as asthma, with
a heart condition or who suffers from epilepsy. Gentle exercise would not be expected
to lead to ‘unusual’ breathing for most pupils but there may be problems with certain
individuals. Asthmatics in school should be known to science staff and may need to be
advised to use a bronchodilator before exercising.
There must be no pressure by the teacher on pupils unwilling to take part. Teachers
must also beware of the subtle influences of peer groups on hesitant pupils; there are
often strong pressures on them to conform and not to be seen to be different from their
friends. The teacher could be the subject for the investigation, but this is often not a
1
These can be purchased from, for example, Philip Harris, Timstar, local pharmacist shops or on-line pharmacists. For economy,
it is sensible to cut the strips in half, longitudinally.
2
Yellow food colouring, a few coffee granules or a little Marmite (which is reputed to help create a more convincing urine
aroma!) can be used.
© CLEAPSS 2006
1441H
good idea as it may interfere with classroom control or the ability to explain or comment on the activity. The teacher could ask for volunteers and from them discreetly
select ‘good athletes’, checking that they are not suffering from colds or any other
minor ailments. This is particularly important for spirometer investigations and
measurements of lung volumes.
Any exercise carried out prior to observations of changes in breathing should not be
excessive and arranged so that it is completed safely; competition between pupils
should be discouraged. Running up and down stairs is inherently much more dangerous than exercising in some other ways. If exercise involves stepping on and off a
raised platform, this must be stable and solidly constructed. Exercise can be made
more quantitative if it involves lifting a known mass (eg, a bag containing 1 kg of
sand) vertically through a known distance, eg, 1 metre.
Pupils should normally be seated during an investigation and carefully observed.
Anyone fainting or complaining of feeling faint should be laid down with legs raised
and sat in the fresh air when fully conscious.
See also section 11.8 (Pupils as subjects of experiments).
14.5.1
Spirometers
If misused, a spirometer1 could cause harm; a teacher should be present all the time a
pupil is using a spirometer. For simple measurements of lung volumes, no carbon
dioxide absorbent is needed.
Possible hazards with oxygen and carbon dioxide absorbent
m Oxygen
As oxygen forms explosive compounds with many oils and greases, such lubricants
should not be put on the connections of oxygen cylinders, on the regulators or on
the tubing connections to spirometers. Water or soap solution could be used to
ease connections between the apparatus and corrugated tubes; glycerol, sometimes used as a lubricant for rubber tubing, should not be used in this instance.
Purging a spirometer with oxygen before use can mean that the air near the
spirometer is enriched with oxygen and there could be an increased fire risk if
ventilation were poor and space confined. There should be no flames nearby.
Medical oxygen
1
The British Oxygen Company has stated that schools should use medical oxygen
cylinders for investigations of breathing. The risks in using ordinary oxygen are,
however, insignificant. The oxygen put into the cylinders is the same; the difference is that ordinary cylinders might have been contaminated when out on hire.
The level of contamination possible is hardly likely to be a hazard to anyone briefly using a spirometer.
This expensive piece of equipment consists of a chamber that is filled with air or oxygen; breathing this gas in and out causes the
chamber to move up and down. The Clifton spirometer is the only model currently listed by school suppliers and costs from £600
(2006 prices). The old SRI spirometer that many schools will already possess and previously sold by Philip Harris, is arguably a
much better model. It is still available from Harvard Apparatus as the Student Spirometer (50-1676) but costs £800. (Harris still
sells spares for this model but they are also available from Harvard.)
The movement of the chamber can be used to record and measure breathing rates and lung volumes including tidal volume,
inspiratory/expiratory reserve volumes, residual volume and vital capacity. If exhaled air passes through a carbon dioxide
absorbent, measurements of oxygen consumption, linked to the effects of exercise, are also possible. In addition, the mechanism
in the body that regulates breathing rate can be investigated. Unless the equipment is to be used only for a few breaths, when
measuring lung volumes, a spirometer must be filled with oxygen.
The use of a spirometer is described in a few references including Revised Nuffield Advanced Biology, Practical Guide 1: Gas
exchange and transport in plants and animals, chapter 2 & Teachers’ Guide I and Roberts, King & Reiss, Practical Biology for
Advanced Level, Nelson, 1994.
Note that a device called a ‘pocket spirometer’ is much simpler, can only measure lung capacity and does not require the levels
of supervision demanded with a normal spirometer. Various spirometer sensors are now sold to accompany dataloggers and these
permit the analysis of lung volumes and breathing rate.
1441I
© CLEAPSS 2006
A size F cylinder is probably the most convenient to use. The valves and regulators now supplied are the same for medical and industrial cylinders. Schools with
the chromed regulators formerly supplied for medical oxygen can, of course, still
use them. Some schools have been successful in obtaining medical oxygen from a
local chemist. See section 9.9 for details of sources, relative costs (though these
should be checked with local suppliers) and the handling & storage of gas
cylinders.
m Soda lime
Soda lime or ‘Carbosorb’ are CORROSIVE but much safer to use than sodium
hydroxide; even so, eye protection must be worn when handling them. For spirometers, the larger particle sizes, such as 5 to 10 mesh, are usually recommended.
(As a rough guide, 8 to 14 mesh has lumps similar in size to demerara sugar,
whereas 5 to 10 mesh has lumps the size of aquarium gravel.)
It would obviously be unwise to inhale any dust in the soda lime. Ensure that the
valve in the tubing connected to the spirometer is positioned correctly so that air
is always exhaled through the soda lime, rather than allowing inhaled air to be
drawn through it. As well as this precaution, we would recommend that when a
new bottle of soda lime arrives, it is taken outside on a windy day and poured
from beaker to beaker so that any dust is blown away. A small layer of polymer
wool (as used in aquarium filters) could also be placed at the inflow and outflow of
the carbon dioxide absorbent chamber.
Because it is important to know when soda lime is exhausted, we recommend the
use of soda lime which has been treated with an indicator dye that changes colour
when no more carbon dioxide can be absorbed. For example, ‘Carbosorb’ (from
VWR) changes from pale brown to greyish white when exhausted and ‘Indicarb’
changes from white to violet. We suggest that indicating soda lime should be
stored in a fairly dark place to prevent any fading of the colour. If there is any
doubt about the condition of the soda lime, a small sample should be treated with
carbon dioxide (chemically generated or from a cylinder) to see if there is any
further colour change.
m Maximum times for investigations
Any investigations involving rebreathed air require particular vigilance on the part of
the teacher.
Spirometer filled
with oxygen:
carbon dioxide
absorbed
This is the most usual way to use a spirometer. Maximum time: 5 minutes.
Spirometer filled
with air:
carbon dioxide
not absorbed
Maximum time: 1 minute.
The only contaminant in the oxygen will be the nitrogen left in the lungs after the
subject has breathed out just before connection to the spirometer chamber. Without carbon dioxide accumulation, the normal regulation of breathing does not
occur and oxygen lack will produce no visible signs of distress. There should, however, be no danger of the subject losing consciousness through oxygen shortage if
the experiment is stopped before half the oxygen has been used or if the investigation lasts no longer than 5 minutes. Useful results can usually be obtained in a
shorter time, say 2-3 minutes.
Only about a fifth of the available air is oxygen and this will become even lower as
the investigation proceeds. Without absorbent in the spirometer, the concentration of carbon dioxide in the air rises and induces a feeling of suffocation. The
subject soon feels short of breath, breathes faster and deeper and will invariably
remove the mouthpiece by him or herself before there is any danger of losing
consciousness due to lack of oxygen. The teacher should time the investigation,
observe the changes in breathing pattern and stop the investigation as soon as a
sufficient recording has been made, or after a minute, whichever is sooner. If the
spirometer is filled with oxygen rather than air, there can be no danger of a lack of
oxygen but the subject on the spirometer will still show the symptoms of stimulated breathing.
© CLEAPSS 2006
Spirometer filled
with air:
carbon dioxide
absorbed
1441J
Maximum time: 1 minute.
The subject feels no ill effects as the investigation proceeds and loss of consciousness is likely to be the first indication of lack of oxygen; the absence of carbon
dioxide prevents a change in the breathing pattern. With the subject at rest, there
should be little danger if the experiment is run for the same time as before: a
minute or less. It is, however, vital that the teacher times the activity carefully
and monitors the subject; observing him or her writing while using the spirometer
enables a constant check to be made for the onset of oxygen shortage affecting
mental alertness. For investigations of breathing after exercise and with most
studies of oxygen consumption which need a more extended period of recording,
the spirometer must be filled with oxygen and not air.
Other points
Resistance to
breathing
If the tubes used to connect different parts of a spirometer are narrow, resistance
to breathing can be felt. The wider the tubes used, however, the more ‘dead space’
there will be. This is important only where there is a single tube that carries both
inhaled and exhaled air. Modern spirometers, where the mouthpiece fits directly
onto a T-piece containing the valves, do not have an appreciable dead space. Little
resistance should be felt with 35 mm diameter tubes though with 21 mm tubes it
is possible that some pupils, particularly those of slight build, may experience
resistance to breathing. When the influence of exercise on oxygen consumption is
being investigated, spirometer readings should be made immediately after, rather
than during, the exercise period. Most resistance to breathing will be felt when
ventilation rate and amplitude are elevated, and performing strenuous exercise
while attached to a spirometer can easily become stressful.
If sudden resistance is felt, it is due to one of the valves sticking. These should be
replaced if the problem persists.
m Hygiene
When a spirometer is in use, water vapour from the exhaled air condenses and
collects in the tubes (the same problem that woodwind and brass players experience). This condensation can be reduced by allowing the spirometer and its tubes
to reach room temperature beforehand. Sometimes saliva can enter the tubes, but
it is less likely to happen if subjects keep their heads erect.
Obviously the mouthpiece must be disinfected after each person has used it; for
this a freshly-prepared solution of Milton is preferred followed by a rinse in water.
Milton does not leave an unpleasant taste but disinfection will take 30 minutes.
If this length of time is impossible, ethanol is quicker (5 minutes) but even a
thorough rinsing in water may not remove the unpleasant taste. [See section
15.12.3 (Chemical disinfection)]. Because of problems of disinfection during the
lesson, it is clearly beneficial to have more than one mouthpiece or use disposable
mouthpieces, if these are available.
At the end of each lesson, the T-piece should be disinfected while the corrugated
tubes connecting the T-piece to the spirometer need to be disinfected and hung up
to drain. For this task ethanol is preferable, because it will evaporate more readily
from the tubing crevices.
Recording
results: using a
kymograph, chart
recorder or
datalogger
1
Spirometer recordings have traditionally been made with a kymograph1 and then
photocopied for individual students if necessary. Spirometers have provision for
mounting a pen on the float. The greatest problems are often experienced with ink
pens having fine writing points that easily become blocked; always ensure that
the pen is thoroughly washed out after use and that the thin wire supplied with
the pen is replaced. With the kymograph model most often found in schools (which
has numbered speed settings), speed 2 is normally employed for investigations of
Although the use of a kymograph with a spirometer is arguably still the most convenient means of making direct recordings,
schools that find the funds to invest in a new spirometer are most unlikely to be able to afford a new kymograph as well. Of the
school suppliers, only Griffin Education currently lists a kymograph which also has a stimulator that is unnecessary for use with
a spirometer and doubles the cost. The manufacturer, Harvard Apparatus, can supply a kymograph without stimulator (50-7350),
a drum (50-5560) and kymograph paper (50-5362) but the total cost is still over £900 (2006 prices)!
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© CLEAPSS 2006
oxygen consumption and the effects of carbon dioxide accumulation, while speed 3
is used for recording lung volumes. [See also section 15.1 (Kymographs).]
A movement sensor attached to a chart recorder (if the department has one) can
also be employed for recording spirometer traces but using a sensor and datalogger “with care and patience” (as described in one supplier’s catalogue!) is the
technique most likely to be required. Depending on which brand of datalogger is
used, there will be a matching movement sensor available.
14.5.2
Alternatives to spirometers
Often because of the expense of a spirometer or perhaps the difficulties of making
recordings with it, less-sophisticated methods of measuring lung volumes are used.
Such activities easily permit correlations to be made between lung volumes and
students’ age, height, mass etc.
Breathing out
into a bell jar
A student’s vital capacity can be measured by a forced exhalation into a large bell
jar immersed in water in a bucket or a deep sink. The bell jar is previously calibrated by pouring in measured volumes of water and a scale in litres is marked
indelibly on the outside. If a suitable bell jar with an opening at the top is not
available, a d-i-y version can be constructed by cutting off the base of a large (at
least 5 litre) plastic container as shown overleaf.
A length of very wide-bore tubing is then attached to the bell jar/plastic container.
The corrugated tubing as used with spirometers is best. Normal-diameter rubber
tubing must not be used because it is extremely difficult to force out a lung full of
air through a narrow aperture.
The bell jar or container is placed in the water and its position adjusted so that
the water level is at the zero mark. A student, wearing a nose clip, breathes in
deeply and then exhales to the full extent possible through the tubing. The bell
jar/container is lifted out of the water as the air is exhaled. The end of the tubing
is then either pinched closed or a bung inserted and the water level inside the
container is made level with that outside before the air volume is recorded.
Beware of students who are athletes or play wind instruments; their lung capacities may be much greater than the volume of the bell jar etc; anticipate water being
spilt everywhere!
Lung-volume
bags
School science equipment suppliers all currently list kits containing different
numbers of special long bags that usually have a capacity of 6 litres and have a
scale marked on them. Some of these kits also contain a valve system1 that
enables investigations to be extended to include the effect of exercise on ventilation volumes. The bags are wrapped around mouthpieces and secured with elastic
bands. For schools on a budget, it is possible to manufacture these bags from layflat polythene tubing2.
1
For example, in the Breath volume kit, A20655, from Philip Harris.
2
Lay-flat tubing can be obtained from Transatlantic Plastics. Details of construction are in an article: Barker, John A, Measuring
lung capacity, Journal of Biological Education, 1983, 17 (4), p286.
© CLEAPSS 2006
1441L
In the past, larger-volume bags, including Douglas bags, were readily available
(sometimes included as part of so-called ‘metabolic-rate apparatus’) to enable the
air breathed out over an extended period to be collected and used to investigate
the relationship between exercise and the amount of air breathed. Such bags will
still be available in some schools (or could even be improvised) and can be put to
good use1. Harvard Apparatus sells Douglas bags but these are expensive.
‘Spirometer’
sensors
Various datalogging manufacturers now sell a ‘spirometer’ sensor that enables
lung volumes to be plotted and is often the equivalent of a ‘pocket spirometer’ (see
footnote earlier).
Peak-flow meters These are used to monitor the breathing of asthmatics and give an indication of
how constricted are the bronchioles. They can be used in science to measure this
aspect of lung function and link this to factors such as age, lung capacity, fitness
etc. Measurements are made by taking a deep breath and forcing out air from the
lungs as quickly as possible. It is customary to repeat the measurements at least
3 times and record the mean value.
Hygiene issues
14.5.3
As with spirometers, it is important to disinfect mouthpieces after use; see earlier
for discussion on disinfectants and disinfection times. Spare mouthpieces can be
purchased for peak-flow meters and lung-volume bags; some types may be supplied with disposable mouthpieces. For the tubing attached to bell jars etc, it is
sensible to attach a separate mouthpiece (perhaps a short length of a wider piece
of tubing), rather than simply inserting the end of the tubing in the mouth. This
will facilitate re-use of the equipment. It is very difficult to disinfect and dry out
the inside of lung-volume bags and since air is not normally breathed in from the
bags, it is arguably unnecessary to do so.
Breathing rate measurements
Many investigations of breathing rate simply involve, by observation, a direct count of
the number of breaths taken per minute and the influence of exercise etc. However,
there are some items of equipment that can be used to enable data to be recorded
electronically and then displayed and manipulated.
Stethographs and A stethograph consists of a short length of corrugated tubing attached to a belt or
chain that is wrapped round the chest. The movement of the chest during breathbreathing belts
ing causes pressure changes in the corrugated tubing which, when the stethograph is attached to a tambour2, a manometer tube and electronic manometer or a
pressure sensor, enables recordings to be fed into a datalogger to provide a
qualitative analysis of breathing.
A device which functions in the same way as a stethograph is a breathing rate belt
sold by various datalogger manufacturers. This is inflated with a hand pump and
pressure changes caused by chest movements are fed to a pressure sensor and
datalogger.
‘Spirometer’
sensors
As well as allowing lung volumes to be studied, datalogger ‘spirometer’ sensors
should also permit the display and analysis of breathing rates. The Philip Harris
breathing rate sensor, which detects temperature changes in exhaled air, is not
designed to measure lung volumes.
1
A good discussion of how to use the large breathing bags in metabolic-rate apparatus, with links to sports and fitness, is given in
section 3.12 of Nuffield Secondary Science 3, Biology of Man, Longman, 1971. It may be old but it’s good stuff! Schools that do
not have access to this reference should contact CLEAPSS.
2
Tambours are no longer sold by school science equipment suppliers but schools may already have them. Pressure changes in the
stethograph cause a rubber sheet in the tambour to move up and down which in turn moves an attached lever. If a pen is attached,
recordings can be made onto a kymograph. Alternatively, the tambour arm could be attached to a movement sensor for datalogging. (However, for such work, it would be simpler to attach a stethograph directly to a pressure sensor.)
14.5.4
1441M
© CLEAPSS 2006
Manometers - pressure gauges
As part of the introduction of pupils to the idea of pressure, teachers often ask them to
blow into manometers. The same apparatus can be used with the aim of investigating
the maximum pressure that the lungs can exert. Whatever the purpose, a trap should
be fitted to prevent the manometer fluid from being ingested.
Over exertion
Pupils who over-exert themselves blowing into manometers etc can faint. See the
introductory section of 14.5.
Disinfection of
mouthpieces
The mouthpiece should be changed after each pupil. For disinfection, see earlier
discussions.
14.6
Disposal
For the disposal of chemicals, see section 7.5 (Chemicals, Disposal); for radioactive
substances, see section 12 (Mainly physics).
m Many local authorities have rules for disposal that should be followed. Ideally, most
discarded non-plant material should be incinerated. For specified items (see below),
steam sterilisation in an autoclave or pressure cooker before disposal will be the norm;
see section 15.12 (Sterilisation). In the process of disposal, it is important to avoid the
creation of aerosols and dusts.
Microbiological
cultures, agar
plates, contaminated pipettes,
syringes etc
While schools should not deliberately study pathogens, there is always a danger,
whenever microorganisms are cultured, that stray pathogenic species may be
present. Destroy all cultures unopened whenever possible. [See also section 15.2
(Microbiology).]
Liquid cultures,
blood
The caps of vessels containing liquid cultures or blood should be loosened slightly
before autoclaving. After autoclaving, the contents should be flushed away as
liquid waste. Use a toilet or drain followed by plenty of water.
If incineration1 is not possible, used agar plates, all cultures including yeast
suspensions, yoghurt and mouldy bread, fruit & vegetables and other contaminated items should be autoclaved before disposal using an autoclavable disposal
bag which is only two-thirds filled and loosely closed, not sealed. Chemical disinfection is no longer considered to be an acceptable method of treatment, even in
emergencies. After autoclaving, the bag should be sealed and placed in the refuse.
Autoclave bags that carry the biohazard warning symbol, although no longer a
hazard after sterilisation, may cause alarm in people who may handle and sort
items in the refuse. Therefore it is preferable that such bags should be placed
inside an opaque plastic bag before disposal.
Swabs used for blood obtained from any source [see section 14.4.1 (Blood)] and
Items contaminated with blood other contaminated items, eg, following accidental cuts, should be autoclaved
before incineration or disposal with the refuse.
Disposal of
formalin
If there is a need to dispose of quantities of formalin, dilute the solution 100 times
before running it to waste down a toilet or drain.
Dead animal
material
A vertebrate animal which has died, or been killed, because of a disease should be
placed immediately in a thick plastic bag and incinerated. If incineration cannot
be arranged for the disposal of an infected mammal or bird, it would be advisable
to contact the local authority disposal department or a vet.
1
Incineration of agar plates, ie, cultures in plastic Petri dishes, poses a particular problem because the dishes are made from
polystyrene. Incineration breaks this down and may release the styrene monomer (phenylethene) which is harmful by inhalation
and irritating to eyes and skin. Styrene vapour should not be inhaled. A purpose-built incineration unit with a very tall flue must
therefore be used and it is most unlikely that schools will have such a facility.
© CLEAPSS 2006
1441N
The carcasses of healthy animals, the remains of preserved specimens and materials obtained from a butcher or abattoir are often wrapped in newspaper, placed
in a thick, opaque plastic bag and put in the refuse.
Cage litter and
droppings
These should be wrapped in an opaque plastic bag. If incineration is not possible,
the waste should be disposed of along with normal refuse.
‘Sharps’ ie,
syringe needles,
scalpel blades,
etc
These should ideally be placed into a purpose-made ‘sharps’ container that cannot
be opened once sealed; see the discussion under Care with lancets in section 14.4.1
(Blood). If a commercial sharps container is not available, another container can
be used, eg, a tin with a lid; which should be firmly attached before the container
is disposed of in normal refuse. In this case, however, hypodermic needles should
first be carefully broken by crushing the plastic mount with pliers.
14.7
Dissection
14.7.1
Whole-animal dissection
Compared with practices in biology that were common in the past, dissection of whole
animals in schools is no longer such a widespread activity. In some areas, animalrights campaigners may have pressed for the abolition of such dissection. Some pupils
and staff may find whole-animal dissection repugnant and teachers do not always
agree on its value. Most would accept that unwilling pupils should not be forced to
dissect or watch a dissection of, say, a small mammal.
Nevertheless, there are teachers who wish to include the dissection of whole animals
in their teaching and they should be permitted to do so. It would be prudent, however,
for such teachers to be able to justify their decision to perform dissections, having
thought through the educational objectives1 for the activity.
It has been recommended by the Royal Society that students below the sixth form
should not carry out a small-mammal dissection themselves but demonstration dissections, performed by teachers, can be shown to students of any age. If small mammals
kept in school are to be used for dissection, it is essential that pupils never come into
contact with these animals while they are still alive.
In almost all cases the use of fresh or recently-defrosted specimens for dissections,
rather than preserved material, is to be preferred. Frozen rats and mice (and some
other animals) are sold by suppliers such as Blades Biological and Timstar. Frozen
small mammals are also sold, more cheaply, by pet shops as food for snakes. Such
frozen specimens could be used for dissection purposes too. Of course, good hygiene etc
is needed; see section 14.7.4.
Dissection
boards
1
Dissection boards, made from soft wood so that awls can be inserted, are available
from suppliers such as Scientific & Chemical and Timstar. Philip Harris has
developed a board using MDF that has a removable and replaceable cork insert.
These boards typically have raised edges to retain fluids.
An alternative is to use old enamel pie dishes or heavy-gauge aluminium dishes
with sloping sides and fill these with molten black wax (eg, from Philip Harris;
A72291 or A72308). Some schools have used large white ceramic wall tiles successfully (but obviously these do not permit animals or organs to be pinned down).
Performing dissections on layers of newspaper is an inexpensive option; experiment with the appropriate number of layers to prevent fluids soaking through to
the bench. Newspaper is particularly useful for eye dissections; students accidentally find out about the properties of the flexible lens when it is removed from the
eye, they squeeze it and they read the newsprint through it!
Refer to documents on the CLEAPSS CD-ROM: PS3, Keeping and using animals and plants: Towards a science department
policy and PS3A, A joint ASE/IoB/UFAW statement: The use of animals and plants in school science may be helpful in clarifying
the educational objectives for performing dissections.
14.7.2
1441O
© CLEAPSS 2009
Dissection of material from butchers etc
There is generally less opposition or reluctance when items for human consumption
from butchers, abattoirs or fishmongers are dissected. Although pupils often feel much
less strongly about the dissection of organs from these sources, it is still important
that they should have the opportunity of declining to become involved in the work.
Following the emergence of bovine spongiform encephalopathy (BSE) and variant CJD
in humans (a form of Creutzfeldt-Jakob disease thought to be caused by eating meat
products from BSE-infected cattle), as well as outbreaks of foot & mouth disease, there
has been obvious apprehension about dissecting organs from butchered animals.
However, as a result of government-initiated controls to protect the health of both
humans and farmed animals, such fears are now groundless and any restrictions
imposed by employers are now based on outdated concerns and outdated guidance.
However, good hygiene etc is needed when handling animal organs; see section 14.7.4.
Dissecting eyes
Why have eyes The infective agents that cause BSE in cattle and scrapie in sheep & goats are
been considered found in the tissue of the central nervous system and, since the eye is intimately
connected to the brain via the optic nerve, handling eyes might be seen as hazardhazardous?
Specified Risk
Material
Availability of
eyes
ous. Note that the agents causing spongiform encephalopathies have not yet been
found in other farmed animals including pigs, deer, horses, llamas and ostriches.
In 1997, new legislation was implemented1 which revoked the earlier orders for
cattle, sheep and goats and introduced the term ‘specified risk material’ (SRM) to
describe those parts of these animals that might present a risk to human health
and regulated its removal and supply. This has, in turn, been subsumed within
the Animal By-Products Regulations 2005 (and previously the Animal ByProducts Regulations 2003) and subsequent amendments. The most recent of
these being in May 2009 (DEFRA note on the Animals By-Products Regulations
2005, 15 May 20092). These regulations deal with the use and disposal of all
animal parts, and include a degree of exemption for “the use of animal by-products
for diagnostic, educational or research purposes”. The consequence of this is now
that, for educational purposes, eyes may be removed from:
• cattle killed before the animal reached the age of 12 months;
• sheep and goats killed before the animals reached the age of 12 months
(or with at least one erupted incisor!).
This may mean that butchers and abattoirs are prepared to supply eyes from
young cattle, lambs and kids, although it is possible that some will continue to
restrict the supply of eyes from cattle, sheep and goats of any age.
The eyes of pigs, horses, deer, llamas and ostriches may also be available. Eyes
from deer, horses or llamas are good alternatives because of their larger size. The
suitability of pigs’ eyes should be considered carefully if the school has pupils with
religious/cultural objections to handling material from pigs. Ostrich eyes would
provide material for interesting comparisons.
However, supplies of eyes from any slaughtered animal are sometimes difficult to
obtain; removing eyes is time-consuming and butchers/abattoirs may be unwilling
to do it and sell eyes. In this case, the alternative of dissecting fish eyes is worth
exploring. Fish heads, eg, salmon, can often be purchased relatively cheaply.
Dissecting and using other organs/tissues
The types of organs most often dissected (hearts, lungs and kidneys from any animal
as well as pig’s trotters and chicken’s feet/legs) are not classified as specified risk
material and therefore should be available and can be used in schools. Note that the
tops of hearts will typically have been cut off by the butcher, rendering them useless
for a school dissection. You need to cultivate your butcher so that intact hearts are
supplied or, alternatively, order several plucks - the heart and lungs together. From
these, the hearts can be removed with long lengths of blood vessels (useful for attach1
Statutory Instrument No. 2965: The Specified Risk Material Regulations 1997.
2
www.defra.gov.uk/animalh/by-prods/approvals/10.pdf.
© CLEAPSS 2007
1441P
ing to water taps to investigate the one-way flow through the heart). The lungs can
then be used for dissection and inflation investigations.
Unfortunately, organs will often have been cut into at the abattoir (as part of the inspection process), so choose the least-hacked about specimens for a demonstration.
Inflating lungs
Naturally we do not recommend that you directly exhale into the trachea! It is
also unwise to insert into the trachea a length of rubber tubing and then put this
into the mouth to exhale air into the lungs. Because of the elastic recoil of the lung
tissue, you might inhale a blast of rather unpleasant air! Use instead, for
example, bellows or a bicycle/foot pump attached to the trachea to inflate the
lungs. Air will almost certainly escape from cut surfaces of the lung, so it is best to
place the lungs inside a large, transparent, plastic bag to stop any aerosols,
contaminated with possible pathogens, from escaping into the laboratory air.
Brain dissections Schools occasionally wish to study a fresh brain; this will be classified as SRM
from cattle, sheep and goats. Therefore use a pig’s brain for a demonstration. An
entire pig’s head could be ordered but it would be much more sensible to enquire
whether the butcher will remove the brain for you!
Mossmax
If a local butcher/abattoir is unwilling to supply organs for dissection, the company Mossmax might be used. This is also a useful source if something unusual is
required, eg, an entire intestine or reproductive system. Mossmax will supply any
specified organs that can legally be removed and these are vacuum packed and
delivered directly to the school by a carrier. However, such a service is not cheap.
Cleaning bones
Use a knife to remove as much flesh as possible. Place the bones in a saucepan of
water to which sodium carbonate is added and simmer until the remaining flesh
can easily be removed, using an old brush. Return to the pan for more simmering
until the bones are cleaned. Immerse in bleach to whiten the bones if necessary.
14.7.3
Preserved material for dissection
In some circumstances, preserved specimens, eg, dogfish and rats, may be preferred.
From laboratory suppliers, these may have been preserved using a safer chemical
than methanal (formalin), though this will still have been used as a fixing agent. Some
suppliers offer a choice of preservation method for the specimens they sell.
Formalin (4%) may still be used if a school preserves its own specimens for dissection
or keeps specimens only for observation in sealed jars; see also section 15.8 (Preserved
materials). Even if specimens for dissection are in formalin-free preservative1, which is
preferable, they should be handled carefully.
m Formalin:
Methanal is TOXIC and CORROSIVE although the risk to dissectors from inhaling
formalin vapour is insignificant if specimens are washed before handling (preferably by previous soaking in water for at least an hour) and again after abdominal
cavities etc are opened. Much more serious is the risk of splashes to the eye, even
if the splash is from a 4-5% solution (HARMFUL and IRRITANT), the usual concentration in formalin. The eye must be immediately washed in running water for
10 minutes. Skin contact should be avoided, particularly before the specimen is
washed, and any splashes rinsed off. While a compound of methanal and hydrogen chloride is known to be a potent carcinogen, it is not believed to be formed if
low concentrations of the two vapours mix.
m 70% alcohol
Ethanol or propanol are HIGHLY FLAMMABLE. Specimens preserved in them
should be washed before use. If a splash subsequently enters the eye, the eye
should still be washed thoroughly.
a solution of
methanal
(formaldehyde)
solutions
Formalin-free
preservatives
1
Specimens preserved in these chemicals should be rinsed before use. Body cavities
should be washed out as soon as they are opened. If a splash subsequently enters
the eye, the eye should still be washed thoroughly.
For example, propylene phenoxetol from Blades Biological or Opresol from Philip Harris.
1441Q
© CLEAPSS 2006
Preservation of fresh material
Preservation by
freezing
A freezer is successful for preserving specimens for long periods but they will
quickly deteriorate if regularly removed, defrosted and later refrozen. Such repeated actions are not recommended.
Preservation in
preservative
Specimens immersed in formalin-free preservatives will last reasonably well but
not as well as commercial specimens which will have been ‘fixed’ first.
m 14.7.4
General precautions with dissections
Good hygiene
Ensuring hands are washed within the laboratory before leaving the room will
prevent the transmission of any infections from microbes (eg, food-poisoning
bacteria) present on the specimens/organs being dissected. It is not necessary to
wear gloves when performing dissections; they are not worn when preparing meat
in a kitchen at home.
Eye protection
This should be worn whenever there is a risk to the eyes, eg, when changing
scalpel blades, cutting bone or cartilage etc or when the dissection material has
been preserved.
Disposal of waste See section 14.6 (Disposal).
material & sharps
Cleaning equipment and work
surfaces
Dissecting instruments are best autoclaved after use, because most disinfectants
attack metal instruments. Contaminated equipment such as dissecting boards
should be cleaned with hot water & detergent and soaked for at least 10 minutes
in a freshly-prepared 1% solution of VirKon disinfectant. For working surfaces,
1% VirKon solution is again the most appropriate disinfectant to use; see also section 15.12 (Sterilisation). Performing dissections on layers of newspaper may
protect bench surfaces sufficiently that they do not need to be disinfected.
Cuts or
scratches
Any received during dissection work might lead to infection. Wash the wound in
cold running water, allow minor wounds to bleed freely and refer to a first aider.
Storage
Do not store material for dissection in refrigerators used for food. If flammable
preservatives are used, a refrigerator modified for storing flammable liquids safely
must be used.
14.8
Enzymes
A study of enzymes is an integral part of all biological work and it also plays a central
role in biotechnology. There is a difference in emphasis between the two. In biology,
enzymes used in the synthesis of new material are very important, as well as the
respiratory enzymes involved in the controlled release of energy within the cell. In
biotechnology, these intracellular enzymes are probably less important than the
enzymes that can be readily extracted and produce their effects in reaction vessels. In
both cases the hydrolases (ie, those involved in the breakdown of substances) are
likely to be the enzymes primarily studied practically but, in biological work, it is
important that other types of enzymes (eg, phosphorylase in the synthesis of starch)
are also studied. Students can otherwise be mislead into thinking that all enzymes are
involved in the digestion of food.
Keeping enzyme Enzyme experiments in the laboratory usually involve three stages.
a) The preparation of the reactants and enzymes so that they will react together
investigations
in the desired way and at a suitable rate. This usually means dissolving reactsimple
ants and adjusting concentrations and may involve buffering the solutions.
b) The reaction in which the enzyme catalyses the formation of the products from
the substrate.
c) Techniques required to display or measure the products.
© CLEAPSS 2006
1441R
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pagination. The text resumes from page 1442.
1442
© CLEAPSS 1991
Stage b) is the part which should be of primary interest but it is not uncommon to
find that the third stage involves the most manipulation and becomes the major
part of the exercise in the pupil’s eyes. In work with junior or weaker pupils, it is
important that the method used for displaying the products is kept as simple as
possible. For instance, in catalase experiments the oxygen evolved may be trapped
as foam, if detergent is added to the mixture. Similarly, in testing for the breakdown of starch (and therefore the appearance of sugars), it is often easier to use
Clinistix1 than for pupils to carry out a test with Benedict’s solution.
The use of kits
When technical assistance is limited, it is important that the preparation of
reactants should be kept as simple as possible. Kits, available from the major
suppliers2, can be of value here, since they provide all the reactants, possibly
already measured out, and give clear instructions for their use. Their drawback, of
course, is the relatively high cost of most kits. The kits normally include all the
chemicals required but assume that standard laboratory equipment is available.
Thermostatically-controlled water baths are commonly required for enzyme
experiments. Some teachers have commented that the most valuable aspect of
enzyme kits is in collecting together ideas for investigatory work which can then
be used at other times.
When dry chemicals are made into solutions, they will usually have a limited shelf
life and different solutions from the same kit will remain active for different
times. In particular, enzymes in solution have a short active period and instructions in the kits on storage of enzymes must be followed carefully.
The reagents used in the kits can all be replaced from the suppliers. In many
cases, the replacements will contain larger quantities of material with a saving in
unit price but a loss of convenience. In some cases complete sets of replacement
chemicals are available to recharge the kits. These sets may be available for
different class sizes.
R40a, Kits for Biology & Biotechnology: Enzymes, discusses the range of enzyme
kits available from commercial suppliers.
Immobilised
enzymes
Immobilised enzymes3 play an important part in biotechnology; the enzyme can be
recovered and reused, or employed in a continuous process. These enzyme-carrier
systems may have a longer active life than enzymes in suspension or solution. The
ease of recovery means that they can be used in more than one experiment and by
more than one set of pupils. For instance, an advanced class may prepare and use
pellets of alginate/yeast; when recovered these may be used by more junior pupils,
thus effectively reducing costs.
Sources of
enzymes
Apart from obtaining enzymes through normal laboratory suppliers, schools can
now select from a range of 10 enzymes, chosen for their use in science teaching,
available from the National Centre for Biotechnology Education.
m Safety
All enzymes are proteins and, as such, may produce allergic reactions. It is good
practice to handle any enzyme as a potential allergen and to minimise skin contact or
the possibility of inhalation. The powerful lipolytic and proteolytic enzymes (eg,
lipase, trypsin, pepsin etc) are potentially hazardous in solid form or in concentrated
1
The presence of certain reducing sugars other than glucose can be detected with Clinistix, though the reaction time to produce a
result is longer.
2
The widest range of kits is available from Philip Harris Biological. Several of the kits previously marketed by Griffin & George
are now available from Educational and Scientific Products.
3
Information on the use of immobilised enzymes can be found in kits from Harris Biological and Griffin and also in the following
valuable publications: P Wymer, Practical Microbiology and Biotechnology for Schools, Macdonald Educational / Simon and
Schuster, 1987, ISBN 0750100885; J Dunkerton & R Lock, Biotechnology - a Resource Book for Teachers, ASE, 1989, ISBN
0863571115.
© CLEAPSS 1991
1443
solution. Wear gloves and eye protection when making up solutions. Dilute solutions
are unlikely to offer any significant risk.
Amylase
Saliva can supply amylase at no cost; (the addition of a small amount of sodium
chloride speeds up the enzyme’s activity). The collection of saliva may be considered to be distasteful and the saliva will carry large numbers of harmless bacteria
from the mouth and possibly bacteria and viruses which are potentially pathogenic. Pupils should collect their own saliva samples. Glassware etc contaminated
with saliva should be put immediately after use into a vessel containing a freshlyprepared disinfectant and then washed up in the normal way with hot water and
detergent. See section 14.4 (Blood, cheek cells and saliva).
Catalase
Shop-bought liver or commercial catalase should be used as a source of catalase;
blood should not be used in an open vessel. Again, see section 14.4 (Blood, cheek
cells and saliva).
Demonstrating
starch digestion
with Aspergillus
Doubts have been expressed about the advisability of using any species of
Aspergillus in schools. Aspergillus oryzae produces a particularly active amylase
and has been recommended in studies illustrating the production of extracellular
enzymes by fungi. Although not listed among the microorganisms regarded as
‘safe’ in Microbiology: an HMI Guide1 this species of Aspergillus is not, as far as is
known, implicated as a pathogen.
14.9
Fermenters
A fermenter is a vessel in which biochemical reactions involving living cells or isolated
enzymes are carried out; careful control of the conditions allows the reactions to proceed at optimal rates to convert raw materials into new products. A fermenter is
invariably fitted with various ‘ports’, holes through which probes or other items are
inserted, while contaminants are excluded from the vessel. The probes measure parameters such as pH or temperature; equipment for stirring, heating or controlling
aspects of the reaction is often also used. A fermenter may be a simple d-i-y construction, a commercial unit in which the monitoring and control of the reaction is carried
out by means of a built-in ‘black box’ or a commercial unit which allows or requires the
operator to interact with the equipment and explore the technology behind the system.
It should be noted that a fermenter can be used for a variety of investigations, not just
for studies of fermentation, and also provides opportunities for supporting work in
information technology with its potential for computer control and datalogging. In
many situations, a fermenter will contain a large culture of microorganisms which
poses a potential hazard. By taking sensible precautions, however, and placing reasonable limits on the work done, any risks involved in using fermenters should be
reduced to an insignificant level.
The CLEAPSS guide R39, Fermenters in Schools, discusses fermenter systems that
are commercially available and includes details of models that are relatively inexpensive as well as more sophisticated units.
National Centre
for Biotechnology
Education
(NCBE)
1
Information on the use of fermenters is available from the NCBE [see section 1
(Addresses)] for establishments that pay the annual subscription to join the
‘Schools Biotechnology Club’. The Newsletter regularly provides information and
ideas for work with fermenters and the NCBE has also developed a simple
fermenter, that can be autoclaved in a pressure cooker, for sale to schools and
colleges. A separate guide to the use of fermenters is to be produced in the spring
of 1991.
DES, Microbiology: an HMI Guide for schools and further education, HMSO, (1990 Reprint with amendments), ISBN
0112705782.
14.9.1
1444
© CLEAPSS 1991
Safety
m Microbiological safety1
Anaerobic fermentation2 of organisms other than yeasts is not recommended for work
in schools.
The DES booklet Microbiology: an HMI guide3 does not refer directly to the use of fermenters but provides authoritative and relevant advice. Guidelines for biotechnology,
including work with fermenters, are given in chapter 5b of Topics in Safety4.
The increased
scale of work
with fermenters
Three levels of microbiological work have been defined and are described in section 15.2.3 (Microbiology). Most school work with fermenters will be at level 2,
with some level 3 work conducted by the teacher or sixth form students. These
levels of work were defined, however, for cultures grown in Petri dishes.
When much larger cultures are grown in a fermenter, it may be true that the
danger is no greater, if everything goes as expected, but the potential risks are
increased should anything go wrong. Thus, if an opportunistic organism should
contaminate a plate (due to poor technique or perhaps just bad luck) it is easily
destroyed by autoclaving without even opening the culture. Sterilising the culture
in a fermenter always poses greater problems because of the larger volumes
involved and, should the fermenter have become contaminated, then the risks are
increased.
Disposal and
spills with large
fermenters
Spills are potentially more hazardous with fermenters than in ‘standard’ microbiological work because of the large volumes of liquids involved; aerosol5
production is a particular risk if a fermenter full of culture is dropped. With some
designs of fermenter or where only a pressure cooker is available, the vessel may
need to be opened for disposal of its contents and this may expose the operator to
greater hazards. Where particular reactions have been investigated, large
volumes of potentially hazardous substances such as enzymes may have been
produced, with consequent problems of disposal or if spills occur. Refer to section
15.2 for details of safety precautions and procedures for dealing with spills.
Sterilisation
Ideally, the entire fermenter vessel complete with medium, all probes and ports etc
should be sterilised by autoclaving before inoculation with the organism that is to be
grown and after the work is completed. Fitting some fermenter vessels plus their appendages into a pressure cooker, which many schools use for sterilisation, is, however,
unlikely to be possible; an autoclave or a tall pressure cooker6 will be required. Different models of fermenter vary greatly in size; an initial, wise choice of fermenter may
solve some of the difficulties of heat sterilisation. Also, if pH is to be monitored, a pH
probe capable of withstanding autoclaving is expensive. It should be noted that the
plastic that is used to seal the ports of some fermenters does not allow the entire unit
to be autoclaved intact; if in doubt, check first with the supplier. There is some
1
For full details, refer to section 15.2 (Microbiology).
2
Some organisms will grow only in the presence of oxygen (aerobic conditions), others only in its absence (anaerobic). Some
organisms are ‘facultative’ and can grow in either situation but with different results. For example, Saccharomyces cerevisiae
(brewer’s yeast) in anaerobic conditions will produce alcohol. With oxygen available, however, it produces more yeast cells to
allow further alcohol production when oxygen is later excluded; yeast shows little growth in the absence of air.
3
DES, Microbiology: an HMI Guide for schools and further education, HMSO, (1990 Reprint with amendments), ISBN
0112705782.
4
Topics in Safety, ASE,1988, ISBN 0863571042.
5
6
A tall steam steriliser which is large enough to take a small fermenter is available from the National Centre for School Biotechnology.
© CLEAPSS 1991
1445
evidence that, for fermenter vessels constructed from plastic materials, a longer time
is needed during sterilisation for heat energy to penetrate; in these circumstances, the
usual period of sterilisation in an autoclave should be extended by 5 or 10 minutes.
Chemical
sterilisation
It is possible to work safely using organisms listed in the following section, Good
practice and procedures, with the fermenter vessel sterilised initially only by
chemical methods1. Scrupulous cleanliness is essential, however, as chemical
sterilising agents are easily swamped by excess organic matter. If the apparatus is
clean to start with and a medium that is unlikely to support pathogens is used,
then sterilisation with chlorate(I) solution will suffice. Solutions of suitable concentration must be freshly-prepared just before use. Air vents should be plugged
with sterile non-absorbent cotton wool before sterilisation. A filter pump is likely
to be useful for the removal of sterilising chemicals through the port that will
later be used for the introduction of medium.
Sterilising media After chemical sterilisation of the fermenter, medium is then introduced asept-
ically1, having itself been sterilised by autoclaving. It is feasible to sterilise media
in a domestic pressure cooker assuming quantities are not too large for the vessel.
Up to one litre of medium can be sterilised properly in a large capacity domestic
pressure cooker if it is given half an hour at 103 kPa (15 lbf in-2). (Note that this is
longer than recommended for normal sterilisation. If an automatic autoclave is
being used, it is not possible to extend the holding time. In these circumstances,
large volumes of medium should be sterilised in smaller batches of no more than
4-500 ml), preferably in medical flat containers. The effectiveness of autoclaving
can be checked by suspending a Browne’s tube2 within the medium on a length of
thread.
Sterilisation
after work is
completed
When investigations with the fermenter are finished, the used culture should be
sterilised before disposal. This should preferably involve the autoclaving of the
entire fermenter plus contents without opening the vessel. Where this is not possible, concentrated clear phenolic disinfectant should be added to the culture in the
fermenter; this will help to minimise the hazards when the fermenter is opened.
Cultures should then be removed into one or more vessels of an appropriate size
so that these can be autoclaved. Particular care must be taken during this process
to avoid spills and aerosol3 formation. Accessory items such as probes, together
with any equipment used in removing the cultures which itself becomes contaminated, should also be sterilised. Once empty, the fermenter itself is sterilised, by
chemical means if this is the only option.
Good practice and procedures
The use of good sterile (aseptic) technique4 is very important for work with fermenters. Contamination of the culture must be avoided.
m Safe growing
conditions
This is the most important principle for safe working in schools. Anyone with
experience in home brewing knows that reasonable attention to cleanliness and
sterilisation is important but this process is normally safe because pathogenic
organisms (ie those that can infect people) will not, in general, grow in the high
levels of sugar or (later in the process) alcohol involved.
School fermenters can be used with yeasts in sugar solutions. Bacteria which are
suitable for use in school fermenters are those which require unusual conditions,
unacceptable to most pathogens. Such conditions can include: low pH (using an
1
For full details, refer to section 15.12.3 (Chemical disinfection).
2
For full details, refer to section 15.12.2 (Heat sterilisation).
3
4
Refer to section 15.2 (Microbiology) for full details of the steps involved in sterile technique.
1446
© CLEAPSS 1991
acidic growth medium), temperatures other than 37 oC, high salt concentrations
or the use of growth media deficient in important nutrients, eg carbohydrate.
Using a high
concentration of
‘starter’ culture
m ‘Safe’ organisms
Table 14.5
Microorganisms
suitable for
fermenters
If sterile growth medium is given a large inoculation of the desired organism, eg,
at least 20% of the volume of the medium, there should be less chance of
contaminating pathogens becoming established in competition with the main
culture. The use of inocula of this size may, however, preclude studies of, for
example, the lag phase of growth.
Some organisms thought to present minimum risk for school microbiology in the
past are now no longer recommended, as our knowledge of their biology has extended. It is therefore very important that organisms identified as unacceptable in
the HMI Guide are not used. It is also possible that microorganisms listed as
being suitable for microbiological work on a culture plate scale are unsuitable for
work in the larger volumes involved in fermenters because there is the risk of
high exposure to humans. Some organisms that are generally considered acceptable for use in fermenters are given in Table 14.5. (Note that some of these are
not listed in the HMI Guide.)
Acetobacter aceti
A vinegar organism.
Azotobacter vinelandii
Grows in nitrogen-free media.
Chlorella sp.
Algae.
Fusarium graminearum
The organism used in producing mycoprotein.
Lactobacillus sp.
L. bulgaricus is a yoghurt organism.
Methylophylus methylotrophus
The organism used to produce ‘Pruteen’.
Photobacterium phosphoreum
A marine, chemiluminescent organism.
Rhizopus oligosporus
Makes ‘tempe’.
Saccharomyces cerevisiae
Brewer’s yeast.
Schizosaccharomyces pombe
A large yeast; easier to count.
Streptococcus thermophilis
A yoghurt organism.
Vibrio (Beneckea) natriegens
Grows in a high salt medium.
There are other organisms that could be used but the list in the table gives plenty
of scope for interesting work.
Electrical Safety
m The large volumes of liquid in a fermenter pose potential risks when the apparatus is
used with mains-powered electrical equipment but there should be few problems with
electrical safety as long as items, such as the pH meter, are obtained from a reputable
supplier.
Sensible
precautions
A fermenter assembly with probes and controls giving a forest of wires can easily
appear frightening but most wires will be at a low and safe voltage. Care should
be taken to keep mains and other leads tidy and it is clearly wise to site electrical
equipment as far as possible from the fermenter vessel and wet working areas.
Some models of fermenter use an aquarium heater and particular care must be
taken to ensure that only safe units are used appropriately.
m Any d-i-y electrical equipment used with fermenters should be designed to work at or
below 25 V.
© CLEAPSS 1991
1447
Production of gases
m Depending on the reactions conducted in a fermenter, there may be the production of
large volumes of gas such as carbon dioxide or methane. A fermenter must, therefore,
be adequately vented to prevent the build up of pressure. To stop the entry of
external microorganisms and prevent the release of aerosols, such vented ports are
usually plugged in some way with non-absorbent cotton wool or fitted with a
microporous filter cartridge. If a gas is produced which is flammable, eg ‘bio-gas’
generation from the fermentation of silage, particular care must be taken to avoid any
naked flames in the vicinity of the fermenter. The use of animal manure as an
inoculum in such work should not, however, be contemplated because of the unacceptable risks of introducing pathogens.
14.9.2
Practical considerations
Choosing a fermenter
There are several issues that must be considered before investing in a fermenter. Cost
is often of overriding importance and may limit choice to one or two cheaper systems
or a d-i-y approach using plastic drinks bottles. Other factors must, however, also be
considered before investing.
Work with the
fermenter
m Safety
For what work will the fermenter be used? Will it be suitable for use by students
as well as for teacher demonstration? Will it be needed for simple investigations
requiring straight forward operation as well as for more sophisticated
experiments in which additional ports for probes etc are essential? Will any
complexity of the hardware and control systems deter investigations by some
students (or teachers!) or is it more important that others can benefit from the
opportunity to use the facilities provided by the technology? Is it essential that the
fermenter system should provide some form of automatic control over parameters
such as pH or temperature?
Factors to be considered here include the volumes of culture used and the greater
risks involved in the event of accidents when using large quantities. The ease
with which the fermenter can be sterilised is also important; there may be problems if it is too large to fit into a large pressure cooker so that cultures have to be
siphoned out before disposal by sterilisation. Also consider possible electrical hazards involved in using the fermenter, for example with a heater.
Accessory equip- Will the fermenter require the use of additional equipment, other than an autoclave or pressure cooker? For example, air pump, thermostatically-controlled
ment needed
water bath, magnetic stirrer, sensors, computer or datalogger.
Running costs
and preparation
time
Some models of fermenter need much greater volumes of media or cultures. These
add appreciably to the cost of using the fermenter and particularly to the time
spent in preparing the media, sterilising and disposing of cultures after investigations are complete.
Working with a fermenter
The major requirement for fermenter work is a suitable vessel with several ports that
will allow probes etc access to the culture or for the addition and removal of culture
and chemicals. A fermenter for school investigations might require as many as seven
such ports and they must be designed so that a good seal against external contamination is achieved. Aspects of fermenter operation and uses for these ports are discussed below.
1448
© CLEAPSS 1991
Aerobic or
anaerobic
conditions
Maintaining aerobic conditions in a fermenter usually requires air to be blown in
continuously with stirring to avoid local oxygen-deficient areas. Anaerobic
conditions for yeast fermentations are easily created because the yeast will
quickly use up the oxygen initially in the system and will then prevent the entry
of further oxygen by the carbon dioxide it generates. Anaerobic fermentation of
some other organisms can be trickier to set up, however, if the organism is killed
by oxygen; such fermentation is not recommended for school work. (Work with
yeasts is probably the only case where anaerobic conditions are acceptable at
school level.)
Batch or
continuous
processing
In industrial production, a continuous process is preferred whenever possible but
much use of fermenters depends upon batch processing. In this, the culture is set
up and allowed to grow for a period, the fermenter emptied and the contents processed to extract the product. Continuous use of a fermenter, where fresh growth
medium is fed in slowly and spent medium removed at the same rate, is attractive
commercially but can have extra problems of control and avoidance of contamination. Such fermenters are sometimes referred to as chemostats1.
Investigations involving both modes of fermenter use are valuable in schools as
important principles, such as factors governing rates of population growth, are
involved and need to be understood. In particular, continuous culture allows the
study of population ecology, homeostasis, enzyme kinetics and the concept of limiting nutrients. Batch processing is, however, the mode of operation that will be
studied most often with the fermenters generally available.
Sensors for use
with fermenters
Most schools will have a pH meter suitable for use with a fermenter assuming
chemical sterilisation is used. An output suitable for feeding to a computer interface, datalogger (eg VELA) is likely to be necessary. A temperature sensor (or electronic thermometer) may also be required and may already be available in the
science department. In some circumstances, an oxygen probe might also be of use.
For more information about sensors, see section 17.3 (Measuring parameters by
electronic methods).
Agitation
It is often more convenient to use a magnetic stirrer which avoids the need for a
port for this purpose. Where agitation is very important, a more powerful stirrer
acting through a port may be necessary. Sufficient agitation of the culture may,
however, be provided by aeration, if used.
Aeration
This may be important for aerobic cultures and requires the use of two ports, one
for air to be blown in and the other for the excess to escape. Both ports must prevent the introduction of contaminants. Sterilised non-absorbent cotton wool filters
are often used. Commercial sterilisable filters are also available. Antifoaming
agents may be necessary with some cultures/media or if the volume of the fermenter is small. The use of antifoaming agents that can be autoclaved is often recommended.
Heating or
cooling
It will be more common in school use for the fermenter to be maintained at a higher temperature than ambient. To achieve this, either a heater is introduced into
the vessel or the fermenter is heated in a water bath. The use of a water bath
helps in keeping a fermenter system less complex. Cooling, if necessary, could be
achieved by the use of a coil of tubing carrying cold water.
The chemicals most likely to be needed include antifoaming agent or acid/alkali
Addition of
chemicals during for the control of pH. If small amounts of relatively concentrated solution are
added, the most convenient method, there is normally no need for concern about
operation
the introduction of contaminants.
Controlling and
displaying
reactions in the
fermenter
1
Control technology is of fundamental importance in the use of a sophisticated fermenter. To maintain the growth of many organisms, it is necessary to compensate
for changes in pH, resulting from their metabolic activity, by additions of acid or
alkali. Temperature may also need close control. The simpler and cheaper fermenter units do not provide control systems. Some models of fermenter, however,
Details of setting up a chemostat can be obtained from the National Centre for Biotechnology Education [see section 1
(Addresses)] for establishments that subscribe to its services.
© CLEAPSS 1991
1449
have built-in control units or use software to adjust pH, temperature or oxygen
levels. Another approach is for a computer to monitor conditions in the vessel and
then prompt the manual control of parameters by the operator. A computer and/or
datalogger plus appropriate software are needed to monitor and display conditions
in the fermenter in order that the reactions taking place can be studied in detail.
CLEAPSS Guides L153, Interfacing Laboratory Equipment to Computers, and
R170, Stand-alone Dataloggers, discuss the many possible units and software that
could be employed with a fermenter. (With a BBC microcomputer, an analogue
interface for accepting the inputs from a pH sensor etc is already incorporated.
An interface will be required, however, for control outputs to the fermenter
system. A connecting box providing 4 mm connections into the BBC computer’s
analogue port is valuable.)
If suitable software for control work is not available, software for this purpose
could be written locally. Suitable routines to provide basic control functions are
supplied with some interfaces and could be adapted.
Whereas controlling a low voltage heater via a relay output from an interface is
relatively straightforward, a suitable system for adding acid or alkali to control
pH is difficult to assemble at low cost. It may be possible to buy a cheap liquid
control valve at electrical surplus shops but it is essential that it can withstand
corrosive liquids. Car windscreen washer pumps could be used but tend to deliver
too much liquid even if pulsed on for a very short period. The method employed in
the control unit of the Philip Harris fermenter, involving a valve operated by an
electromagnet is an elegant solution to the problem.
Guarding
against spills
There is always a risk of the fermenter leaking or being knocked over, causing a
spill. It is therefore a sensible precaution to stand the fermenter in a large, impervious container which will retain any gross spillage. There is a particular risk of
a leak or small spill when withdrawing samples from a fermenter; it is often recommended that a pad of tissues, moistened with clear phenolic disinfectant,
should be placed underneath the sampling position to soak up any drops of culture
that might escape.
Sampling
cultures in a
fermenter
Depending on the reaction being studied, there may be the requirement to withdraw samples from a fermenter, for example, to monitor cell growth or to measure
a variable such as density (in the production of alcohol with yeast fermentation).
Some models of fermenter allow cell growth to be monitored in-situ by measuring
optical density. Where a sample has to be withdrawn, it is clearly important that
the operation is safe; avoid contamination of the environment by the culture and
guard against aerosol1 formation.
Fermenters: the d-i-y approach
Where financial considerations preclude the use of even the cheapest commercial
fermenter system, or the direct involvement of pupils in the technology of designing
and making a working unit is desired, the use of plastic, ‘fizzy drink’ bottles can be
very rewarding. These have several advantages over glass vessels: they are cheap,
disposable, unlikely to break, available in a range of sizes and relatively easily allow
the necessary ports to be created. The types of reaction that can be studied in such a
vessel, which is not easy to sterilise or ensure totally leak-proof, are limited to some
extent by safety considerations. Thus the growth of only culinary microorganisms,
such as brewer’s yeast or yoghurt bacteria, would normally be contemplated.
Making ports in
the fermenter
vessel
1
At least two ports will normally be needed: one for an air inlet and one to enable
samples of the reactants to be sampled. An additional port for a pH electrode
might be needed. The neck of the bottle can be used as a port for a thermometer or
temperature sensor probe, this function being combined with that of an air vent
1450
© CLEAPSS 1991
A d-i-y
fermenter
Cotton wool plug
pH electrode
Thermometer or
temperature probe
Air inlet from
aquarium pump
Cotton wool filter
in air line
Outlet for sampling with
a syringe & 3-way tap
Aquarium airstone
diffuser
by inserting the thermometer etc through a cotton wool plug. Ports in the side of
the bottle can be made using a cork borer, of appropriate size, which has been
gently heated in a flame. Carry out this operation in a fume cupboard in case the
hot metal causes fumes to be given off from the melting plastic. It is normally
necessary to use a cork borer which is smaller than the hole required, in order to
obtain a tight fit when a bung is fitted in the port. Alternatively, a soldering iron
fitted with an appropriate size bit can be used.
Positioning and
sealing ports
Some thought should be given to the position of the various ports in the fermenter.
The aeration port will naturally be at the bottom of the vessel. The port through
which samples are taken should not be too low or it may become blocked by sediment or settled cells. A pH electrode port will normally be towards the top of the
bottle. Having fitted bungs etc into the ports, these can be sealed into position
using silicone sealant, as used in aquaria, or with a hot glue gun.
Aeration and
mixing
Air pumped in from an aquarium pump via a diffuser stone will usually cause
sufficient agitation to keep the fermenter contents well-mixed. There may be
problems with the airstone becoming blocked if yeast and yoghurt suspensions are
cultured. An alternative is to use a Pasteur pipette to produce a fine stream of air
bubbles. Ensure that the aquarium pump is positioned higher than the fermenter
and/or a one-way valve fitted in the air line, to prevent siphoning of the culture if
the pump is turned off or fails. It is sometimes recommended that a loose plug of
sterile non-absorbent cotton wool is used as a filter in the air line.
Taking samples
of the fermenter
contents
Samples of the reactants can be drawn off using an appropriate size of plastic
syringe fitted to tubing attached to the sampling port. There may be concern that
the sample taken is not representative of the complete culture because too much
of it is from the ‘dead space’ in the tubing attached to the port. A simple solution
is to fit a 3-way tap into the sample line and attach two syringes. The first is used
to draw off an initial sample which will include the culture in the tubing. The tap
is turned and a further sample is drawn into the second syringe. The initial
sample can then be ejected back into the fermenter.
Stability
The fermenter with attached tubing, pH electrode, temperature sensor etc is
easily knocked over, so it will be important to make the vessel secure using retort
stands and clamps. As discussed earlier, the entire assembly should be placed in
a large tray to catch any spills in the event of a leak.
© CLEAPSS 1991
Sterilisation
14.10
1451
The size of the fermenter vessel and its plastic construction preclude steam sterilisation in an autoclave. The fermenter will therefore have to be chemically sterilised, as described earlier.
Genetic engineering
Genetic engineering involves the manipulation and transfer of nucleic acid carrying
inheritable genes into an organism where they are not normally found. These processes can be complex and require detailed and accurate work, particularly when the
donated DNA has to be built up in the laboratory from several sources.
This very exciting field can produce tremendous benefits in medicine, industry and
agriculture but it is also potentially dangerous and all but the simplest demonstrations are technologically outside the scope of schools. The simple investigations that
can be carried out in school lack scientific rigour and it may be argued that it is scientifically more acceptable to use well-prepared visual aids than these practical
activities.
Weaknesses in
commercial kits
available
m Safety
In commercial kits1, pupils have to be given most of the essential information;
they find out relatively little for themselves by experiment. For instance, in one
investigation, students are led to believe that the observed results have demonstrated the transfer of genes carried by plasmids from one organism to another.
There has, in fact, been no direct demonstration of this, other explanations of the
observed results being possible. The instruction booklets supplied are of necessity
very condensed and they make unsupported assumptions or refer to important
applications which are not always explained. The activities in most of the kits
require a high degree of sterile technique.
Investigations of plasmid transfer, when no new nucleic acid molecules are involved and no transmission of new characteristics to another generation occurs,
may be carried out in schools and the kits available fall into this category. It
should be emphasised that plasmid transfer demonstrates a process which does
occur naturally.
It is, however, poor scientific training if students are not made aware of the hazards of genetic engineering in general; the production of organisms carrying new
combinations of genes is potentially very dangerous and considerable restrictions
are placed on research laboratories.
Plasmid transfer work in schools does carry the risk of introducing known plant
pathogens into soil where they were not previously found. This is of course a risk
to plants and not pupils. Some of the techniques mentioned in the kit instruction
booklets are potentially hazardous if the instructions are not strictly adhered to.
14.11
Greenhouses
Greenhouses can be a valuable source of material for school science and used for a
variety of investigations. They must, however, be adequately resourced or else they
are useless. One way of cutting down on the labour they undoubtedly entail is to have
some degree of automation. To be effective throughout the year, some form of heating
is required, if only during frosts, but few schools can now afford this, except for the
smallest greenhouse. A further problem can be vandalism: unless there is a site where
this is unlikely to occur, it is unwise to erect a greenhouse. If adequate resources or a
suitable site are unlikely to be available, schools should consider growing plants in
laboratories: see section 15.5 (Plants and seeds).
1
The Plant Tumour Kit is available from Philip Harris Biological; the Genetic Transfer Kit, Introduction to Natural Genetic
Engineering Kit and the DNA Extraction Kit, all previously marketed by Griffin & George, are now available from Educational
and Scientific Products.
1452
© CLEAPSS 1991
Siting
As there should be sufficient sunlight, especially in winter, it is important to
consider shading by trees as well as by boundary walls and buildings, both
existing and projected, when the sun is low. Some protection from wind is an
advantage, however. The site should be sufficiently near the school for mains
water and electricity to be supplied without excessive cost and as far as possible
from games pitches and boundary fences to reduce damage from missiles. For
urban schools, a site on the roof could be considered but local by-laws might
prohibit this and wind is likely to be a problem.
Base
A solid base, such as in a school courtyard, limits the plant growing area to the
staging; it is much better to site it on open ground. Two spade-depths of good top
soil should be provided with paving slabs for pathways etc.
Orientation
If possible, the ridge of a separate greenhouse should be N - S. A lean-to should be
against a wall which faces south, at least approximately.
Size
If courses require pupils to do practical work in the greenhouse, it must be large
enough to take a full class and for investigations in progress to be left: a
commercial-sized greenhouse will be needed. Most schools may only require a
small house for the supply and maintenance of material for the biology laboratory;
even so, houses 3 m wide and up to about 6 m long are recommended. Cost of
heating must be considered, however.
Shape
The Dutch-light type house is preferable with glass down to the ground so that
plants can be at ground level, cutting down on costs of staging. If the ground
around the greenhouse is to be cultivated, the house should be supported by at
least one course of bricks to protect the glass from hoes etc. Sliding doors and the
absence of door sills are an advantage, especially if some pupils are handicapped.
Materials
Aluminium-framed houses are preferable to wooden-framed constructions for two
reasons. First, the glass supports on the roof are narrower in an aluminium house,
especially around the vents, allowing in more light. Secondly, less maintenance is
required, a considerable saving with larger houses.
Glass is still considered to be the best transparent material unless persistent
damage from stones is likely. Plastics have lower densities and their use would
allow greenhouses to be constructed with a reduced thickness of frame, so offering
less obstruction to light. However, they expand more than glass for a given
temperature rise and so are liable to problems with seals around the panes.
Although in small greenhouses the panes are probably too small for their
expansion to be a serious problem, this should be considered before fitting plastic
panels in a greenhouse designed for glass. Transparent acrylic sheet has been
used by some schools quite successfully but light transmission falls after a few
years because of scratching by grit; also, it is expensive. One manufacturer, Serac
Ltd, has sold greenhouses which use a double-skinned plastic material for a few
years; they appear very successful.
Polythene sheeting cannot be recommended for a school greenhouse because it is
so easily damaged. Some commercial growers use it but expect to throw it away
after a year or so.
Erection
Most firms offer an erection service and their experienced workmen are likely to
erect a greenhouse much more quickly and satisfactorily than amateurs or even
normal builders.
Services
If possible, all services, heating, extra lighting, water and ventilation, should be
automatic, so that demands on staff are minimised and holidays or staff absences do
not result in damage to plants. The controls for these services should be near the
entrance to the greenhouse.
© CLEAPSS 1991
Water supply
1453
The main stopcock should be near the door. Outlets for automatic watering
systems are needed but a tap is also essential for hand watering etc. Guttering on
the greenhouse is often neglected as a source of water; aluminium-framed houses
often have an integral narrow channel which can be used. Any system fitted must,
however, be as narrow as possible to minimise obstruction of light.
Electricity supply All electrical installations must be waterproof and outlets should be provided all
round the greenhouse. Electricity is not the cheapest way of heating a greenhouse
but it is the only method which is easy to automate.
Ventilation
Many greenhouses have an inadequate number of vents in the ridge; their area
should be at least one-sixth of the floor area. Some side vents should also be
provided together with, in larger greenhouses, fan ventilation controlled by rodtype thermostats, to produce enough circulation of air to reduce the growth of
moulds.
Heating
While electrical systems are ideal, occasional frost protection can be provided with
paraffin or LPG heaters. In some schools, a lean-to greenhouse could be heated
with radiators supplied by the school system. While the heating system would
probably shut down on most nights, there is likely to be a frost control which
would provide limited circulation when the temperature approached zero. It is
possible to insulate all or part of a greenhouse by fixing bubble film inside the
glass and hanging curtains of it.
Suppliers
Local garden centres and d-i-y stores sell greenhouses; the Yellow Pages will give
names of local suppliers. National suppliers at the time of writing include Alitex,
Cambridge Glasshouse Co, Clear Span, Pratten, Serac and Unique Dutch Light
Co.
14.11.1 Automatic watering systems
There are three types of automatic watering systems which might be considered
suitable for school greenhouses: capillary beds, trickle systems and mist systems.
Capillary beds
Media and
containers
The most common medium for a capillary bed is a sand/gravel mixture. Sand
alone can be used but, because of the smaller particle size and, therefore, greater
capillary rise, care must be taken to ensure that the bed does not become too
soggy, killing plants by overwatering. Similar care is required in using special
matting made of artificial fibres which is also suitable. Its advantages are its
lightness compared with sand/gravel and that it can be washed.
Sand and sand/gravel mixtures are heavy even when dry, so very substantial
staging is needed to support such a capillary bed. Containers made of fibreglass
are very suitable but are relatively expensive. Wooden (or metal) containers lined
with several thicknesses of heavy-duty polythene sheets are much cheaper but
care must be taken to ensure that movement of the medium, when flower pots are
settled into it, does not make holes in the polythene. Corrugated sheeting can also
be used for the base of a container.
If artificial fibre matting is used, only a flat horizontal surface covered with
polythene sheeting is needed on which the matting, cut slightly smaller than the
polythene sheet, is placed. The matting at some place should dip below or overhang the bench into the water supply. Capillary action will lift water about 5 cm
and spread it about 2 m from the supply point.
Maintaining the
water level
The most common method is to position a small cistern with a ball valve at the
required height. With the appropriate valves this can be used on low pressure or
mains pressure.
1454
© CLEAPSS 1991
Trickle systems
These consist of a system of pipes and plastic nozzles through which water drips.
Water is supplied either directly from the mains through a pressure regulator or
indirectly by slowly filling a small tank which then empties by syphon action.
Water supply
In the direct system, the amount of water a plant receives is governed by the
duration the mains tap is on. Schools would need to automate this, either with a
programmable water valve (battery-powered) or with an automatic water controller which responds to a sensor placed among the plants. The latter system
requires mains power and installation by a competent person.
In the indirect system, the amount of water that a plant receives is governed
mainly by the rate at which the tank fills and empties. The commerciallyavailable tanks that we have tested have been unsatisfactory because it was
impossible to maintain a steady flow rate: it became slower and slower and finally
stopped, perhaps due to deposits from hard water.
Piping
Two systems of piping are available. One uses flexible plastic tube approximately
8 mm internal diameter (id) with the nozzles fixed into it at regular intervals.
Another system has a rigid plastic pipe approx 20 mm id, with narrow bore
flexible plastic tubes, each with a nozzle attached, joined to the larger pipe, at
regular intervals. If the second system is used, the rigid pipe can be mounted say
a foot above the staging and it is then much easier to move individual flower pots
without upsetting the position of the nozzles for other pots.
Mist systems
These are the most complicated and expensive systems. They are most commonly used
for propagating cuttings, often in conjunction with soil-heating cables.
Mist head and
supply
A mist head consists of a very narrow nozzle through which water is projected at
mains pressure and a small flat plate which deflects the water horizontally
through nearly 360o. Mist heads can be mounted with water supplied from above
or below. If the supply is from below, the pipe can run beneath the bench or can be
embedded in the medium with 50 cm-long standpipes at about 1 m intervals with
the heads on top.
Control gear
Either the programmable water valve or the automatic water controller, mentioned in Trickle systems above can be used.
Choice of system
The most useful automatic watering system for a school, where plants are
unattended at weekends, half terms and holidays, is undoubtedly one or more
capillary beds. If sufficient help is available it would probably be cheaper to make
your own containers but fibre-glass containers are not so expensive that their use
in schools is ruled out. A fibre-matting system seems very satisfactory if a suitable
level surface is available.
Trickle systems are particularly useful when the soil inside the greenhouse has
been planted. The flexible pipe system is most suitable.
Suppliers
Local garden centres and some d-i-y stores sell watering systems; the Yellow
Pages will give names of local greenhouse specialists. At the time of writing,
national suppliers include Cameron Irrigation, Thermoforce and Two Wests.
© CLEAPSS 1991
1455
Simple header
tank for a trickle
watering system
Water supply
5 litre plastic
container
6 mm bore tube
pushed into the
10 mm tube
10 mm bore tube
To the trickle line
With a 12 mm diameter cork borer, make a clean-edged hole about 5 cm from the
top of one of the narrower sides of a 5 litre plastic container. Insert a 10 cm
length of 6 mm bore PVC tubing into about 60 cm of 10 mm bore PVC tubing
using some silicone grease and a piece of dowel to ease it into place about 15 cm
from one end. This acts as a constriction which helps the syphon start. Push the
end of the longer tube through the hole in the container, so that the restricted
part is half through the hole and the open end near the bottom; if its end is cut at
an angle, the water will syphon even if the tube touches the base of the container.
The trickle line is connected to the other end of the tube.
The tank should be placed at least 75 cm above the plant bench and the first
60 cm of the tube from the tank must be vertical.
14.11.2 Automatic ventilation
Greenhouses need to be carefully ventilated if plants are to thrive and if fungal
infections are to be minimised. Where ventilation is most critical is in preventing
overheating; a short period of sunshine can, unless the ventilation is increased, raise
the temperature of a greenhouse dramatically and rapidly harm the plants. If
ventilators are kept permanently open, however, the average temperature and humidity of the air in the greenhouse are likely to be too low for plants to thrive.
The purpose of an automatic ventilation system is to allow a greenhouse to become as
warm as is desired but without overheating. No one in a school has time to watch
continually the temperature of a greenhouse during the school week and no one is
available during weekends and holidays. Thus an automatic system can be useful and
justify its cost through plants made available for biology teaching or even for sale.
Here a thermostat controls a motor, causing it to rotate in the appropriate direcElectricallytion whenever the temperature is too high or low. The rotation opens or shuts
powered
ventilator opening hinged vents through a geared drive leading to a rack and pinion on each vent. It
should be possible to open some vents manually in case of power failure. At the
time of writing, details of systems of this kind can be obtained from Electroflora
and most greenhouse manufacturers.
Fan systems
1456
© CLEAPSS 1991
A thermostat switches on an extractor fan when the temperature inside the
greenhouse reaches a pre-selected value and switches it off when the temperature
has fallen sufficiently; cooler air is drawn in through ventilators, perhaps louvres
situated in the opposite wall to the fan or vents controlled by mechanical
ventilator-opening devices. The siting of the thermostat is critical; it must be
totally shielded from sunlight and not fixed to the aluminium frame or the glass.
Details of systems of this kind can be obtained from Cambridge Glasshouse Co
and Two Wests.
The expansion of wax in a cylinder moves a piston against a spring. The piston is
Mechanical
ventilator opening coupled through a simple system of rods to a hinged vent or to louvres which open
when the piston moves outwards. When the temperature falls, the wax contracts
and the spring moves the piston back, shutting the vent. This system is available
from garden centres, d-i-y stores, Thermoforce and Two Wests.
Comparison of
different
ventilation
systems
The most suitable system for greenhouses above 45 m2 (500 sq ft) is electricallypowered ventilator opening. Creedy1 suggests that, for houses below this size, a
230 mm (9") or 300 mm (12") extractor fan controlled by a 600 mm (24") rod-type
thermostat is suitable. However, for small houses, ie below 22 m2, mechanical
systems are likely to be satisfactory and cheaper.
The size at which a fan system becomes more economical to install depends on a
number of factors, one being whether electric power would have to be led to the
greenhouse for the purpose. One mechanical opener should be fitted to each
openable window but eight or so would have to be installed before their cost
reached the purchase and installation costs of a fan system. Mechanical openers
are easy to install, cost nothing to operate and have almost no maintenance costs.
They are the most practicable system for small greenhouses.
Number of
openers needed
14.12
The total area of the roof vents should be equal to one-sixth to one quarter of the
floor area of the greenhouse. This standard is unlikely to be met in smaller houses
and it may be found necessary to fit an opener to all windows; much depends on
the extent to which a greenhouse is sheltered from sun and wind. In smaller
houses, side vents kept permanently open in sunny weather may be essential for
adequate ventilation as, in conjunction with the roof vents, they produce a
chimney effect. It would be wise to experiment with openers fitted to some of the
windows before purchasing enough for all.
Habitat creation
School grounds provide the potential for a great deal of investigative work in science
and environmental studies. Such potential is obviously easier to exploit where the
school already has grounds which are ‘green’ and extensive. Much however, can also
be achieved in schools on built-up urban sites. Barren areas can be transformed or
existing sites can be improved. Schemes for developing new habitats need not be
elaborate; much can be achieved, for example, merely with washing up bowls or
troughs of soil or water to study colonisation and succession or with the erection of
bird tables or nesting boxes. Invertebrates can be encouraged by creating piles of
stones, logs or old timber. The construction of a pond, butterfly garden or wildlife area
will, however, provide more scope for investigative work.
Fortunately, there is a wealth of material to support schools in creating new habitat
areas and a selection of titles is given in Table 14.5. Comprehensive lists of reference
materials2 have been compiled by the Council for Environmental Education.
1
J Creedy, A Laboratory Manual for Schools and Colleges, Heinemann - London, 1977,ISBN 0435571303, p47.
2
See the CEE Resource Sheets: Helping the Environment and Practical Nature Conservation.
© CLEAPSS 1991
Table 14.5
1457
Reference material for habitat creation projects
Title
Conservation in school grounds
Promoting nature in cities and towns
Author
Nick Forster
Malcolm Emery
Date
1987
1986
Publisher
BTCV
Croom Helm
How to make a wildlife garden
Chris Baines
1985
Pond design guide for schools
Butterflies: A practical guide to using
school grounds
Ecology in the National Curriculum: A practical
guide to using school grounds
Learning through landscapes:
The final report
Learning through landscapes: Using school
grounds as an educational resource
The garden bird book
Creating & maintaining a garden to attract
butterflies
The pond book
Bird studies using school grounds
Graham Flatt
John Feltwell
1989
1990
P R Booth
1990
Eileen Adams
1990
Kirsty Young
1990
D Glue
John Killingbeck
1984
1985
Urban Spaces Scheme
Royal Society for the
Protection of Birds
Peter Cawdell
Peter Sibley
1986
-
Elm Tree Books/
Hamish Hamilton
Hampshire Books
Learning Through
Landscapes
Learning Through
Landscapes
Learning Through
Landscapes
Learning Through
Landscapes
Macmillan
National Assoc for
Env Education
PNL Press
RSPB
1987
1989
School Garden Co
School Garden Co
Starting a butterfly garden
Starting a wildlife pond
ISBN
0946752087
Out of print
but available
from TRUE1
0241118700
1870651227
1872865003
187286802X
1872865011
1872865046
0333367650
0907808123
185116801X
1851168028
14.12.1 Planning the development of a new habitat
The first task is to consider carefully the educational objectives that you hope to
achieve by the creation of a new habitat. These will determine the scale and complexity of the work to be done. If your intention is to establish more than just a few
mini-habitats, careful planning will be needed. Using school grounds as an educational resource from Learning Through Landscapes includes case studies and a
detailed ‘action plan’; the final report of the project is aimed more at managers, head
teachers and authority planners. Promoting nature in cities and towns also gives
extensive advice, particularly about the development of habitats on urban area sites
where planning permission may be required.
Choosing
a site and
planning its
development
A detailed survey will be needed. Factors to be considered include shading by
trees and buildings, drainage, shelter from winds, access to the site and to water
supplies, vandalism and ease of maintenance once the habitat is established.
Safety must be a particular concern, especially when constructing a pond. If pupils
will be using the habitat regularly in their investigations, it is important that you
plan for their presence so that it is not destroyed or damaged by trampling or
excessive disturbance.
Consultation
Consult widely at this stage to obtain the views of all interested parties and the
advice of experts within the LEA or elsewhere. Permission may be required before
plans can be implemented.
1
Hardback or paperback copies are available while stocks last from the Trust for Urban Ecology.
Costing and
finance
1458
© CLEAPSS 1991
A detailed costing will be required before bids for financial assistance can be
made. Money may be raised within the school, from parents or the local community, or from national bodies that award grants for environmental projects.
Applications for grants must often be made up to a year in advance.
Promoting nature in cities and towns includes helpful advice on fund-raising and
applying for grants. Major sources of grants include the Nature Conservancy
Council and the Trusthouse Forte Community Chest (organised in collaboration
with the Conservation Foundation). Details of grant-awarding bodies can be found
in the Directory of Grant-Making Trusts (consult local libraries), the publication
Environmental Grants from the Directory of Social Change and in the list
compiled by the British Trust for Conservation Volunteers1.
14.12.2 Creating habitats
References in Table 14.5 should be consulted for detailed information. Promoting
nature in cities and towns is wide-ranging and discusses all types of habitats; it also
includes many further references to other sources and organisations. Apart from
obtaining advice on how to build the new pond, garden etc, consideration must also be
given to the practical aspects of carrying out the construction work and who will do it.
The British Trust for Conservation Volunteers can advise on the skills required and
the organisation of teams of volunteers.
Studying birds
School grounds can easily be improved to attract more birds and facilitate their
study. The RSPB, among others, can provide guidance on the construction of bird
tables and nest boxes and on other strategies.
Constructing a
pond
Refer to Table 14.5 for several useful references, including Emery; the book by
Sibley is particularly accessible and helpful. The publication Waterways and
wetlands in the ‘Conservation Handbooks’ series from BTCV will also be useful
where detailed guidance is needed. Materials for the construction of the pond
include clay, pre-formed glass fibre structures, concrete and flexible plastic or
butyl rubber liners. For many ponds, the use of a flexible liner is likely to be the
most appropriate method to adopt; all other methods have particular drawbacks
or problems associated with them. Butyl rubber is probably the best lining
material, though the most expensive.
Ponds
and safety
In designing a pond, particular attention must be paid to safety matters; its
location, size, depth, edging and even its method of construction will all be
influenced by the need to ensure that the pond can be used safely. A pond is best
sited so that it is obvious but not near a regularly-used pathway. Restricting
access to the pond in some way will help to make it safer; this need not involve
the construction of a high fence or wall.
A large pond (up to 12 m2) allows the development of a shallow marsh region
together with an area of open, deeper water. It is often recommended that any
marsh area and up to two thirds of the rest of the pond should be flat-bottomed
and only 25 cm deep. The remaining area of the pond should be terraced into two
regions of 50 and 75 cm depth with this deeper water sited so that there is less
risk of pupils falling in. The edges of the pond should be clearly marked and
preferably raised to reduce the risk of accidents. A fuller discussion of safety
issues has been made by Bunyan2.
1
Schools requiring a copy of the BTCV information should contact CLEAPSS.
2
P Bunyan, Safety and the school pond, School Science Review, 69 (247), 1987, p286.
© CLEAPSS 1991
1459
Stocking
a pond
Once the pond has been constructed, plants can be introduced, using appropriate
species chosen, according to the structure of the pond, from recommended plants
listed in the books by Sibley, Emery etc. The water in the pond can be left to
become naturally colonised by animals, or invertebrates can be deliberately
introduced from an established pond. If a variety of animal life in the pond is
needed, do not introduce fish such as goldfish or koi; these will consume most, if
not all, of the invertebrates present. The pond may be colonised naturally by
amphibians but spawn of common species1 collected elsewhere can be introduced.
Mini-ponds
If the construction of a full-size pond is impracticable and small, shallow troughs
or washing-up bowls are too limiting, the use of cattle drinking water troughs2
may be appropriate. These are large, sturdy, plastic tanks, with a capacity
between 600 and 1800 litres, which provide the additional depth and volume that
are lacking in smaller mini-ponds.
Wildlife gardens
and meadows
Again refer to Table 14.5 for suitable sources of information. Hedging3 and
Woodlands4, in the ‘Conservation Handbooks’ series from BTCV, are particularly
detailed. The precise structure of your wildlife area will have been determined
earlier at the planning stage when objectives were considered. It may include the
construction of a pond, the introduction of plants to encourage butterflies or other
animals, the management of species-rich meadowland or leaving areas to ‘run
wild’. The creation of a ‘natural’ community of plants and animals will often
require deliberate intervention, for example, by reducing soil fertility or sowing
the seeds of certain species5. Expert advice is, however, readily available.
14.12.3 Management of a new habitat
Once the new habitat has been developed, work does not stop; there will be a demand
for its regular maintenance and management; the initial creation of the habitat will
only have begun the process of change and maturation. Promoting nature in cities and
towns is particularly extensive in its discussion of the long-term management of
habitats. Ponds will at least require the removal of fallen leaves in the autumn, while
grassland areas will need regular cutting and selective weeding. It is important that
these regular commitments are considered at the initial planning stages, so that the
first wave of enthusiasm and commitment is not wasted by allowing the site to
deteriorate.
14.13
Hygiene
High standards of hygiene are essential in any science laboratory and should be
practised whenever appropriate by all staff and pupils so that they become routine.
This is not just to protect against the inadvertent transfer of microorganisms and
chemicals but also to provide pupils with the opportunity to understand the importance of good hygiene as part of the school’s general programme of health education. In
encouraging the correct attitudes and approaches, it is vital, therefore, that good
practice is observed at all times rather than just when the topic of hygiene occurs in a
teaching programme.
Good hygiene must also extend to all preparation rooms and, even in the busiest
science department, there should be regular occasions when cluttered sinks and
1
Refer to section 14.2 (Animals in the wild).
2
Drinking troughs in various sizes are available from Paxton and described in their agricultural products catalogue.
3
Alan Brooks & Elizabeth Agate, Hedging, BTCV Publications, 1975, ISBN 0946752028.
4
Alan Brooks & Gay Voller, Woodlands, BTCV Publications, 1980, ISBN 0950164372.
5
Details of the suppliers of seeds for planting wild flowers or ‘native’ species are included in relevant titles listed in Table 14.5
and also in the CEE Resource Sheet Practical Nature Conservation.
1460
© CLEAPSS 1991
benches are completely cleaned and disinfected. Consuming food and drinks in a prep
room at break and lunch time should never be allowed; see section 3.5 (Personal
safety: Eating and drinking).
14.13.1 Good practice
Washing
facilities
Adequate means of washing hands with soap and warm water must be provided in
both prep rooms and laboratories. It is especially important that all pupils in a
class can properly wash their hands in the laboratory, rather than being required
to use the facilities in the toilets which may be some distance away, after
completing any practical work involving the handling of animals, plants,
microorganisms, soil, pond water and chemicals. Particular attention must be
paid to the provision of appropriate means of drying hands after washing; a single
communal towel is quite inadequate and may leave hands more contaminated
than they were before drying!
Cleaning sinks
and surfaces
In most instances, for example after using various chemicals, it will be sufficient
to wipe down a bench surface with a clean, damp dishcloth. After microbiological
work or at other times when there is a risk of contamination with microorganisms,
the bench top should be disinfected1. It follows, therefore, that all laboratories and
prep rooms should have an adequate supply of dishcloths that are regularly rinsed
(preferably by students after using them), kept reasonably clean and replaced
when no longer fit for use. Cloths used to disinfect surfaces should either be
disposable or themselves disinfected, preferably by autoclaving. Sinks should be
routinely cleaned and disinfected.
Eye protection
and microscope
eyepieces
There is no risk of transferring head lice on items used for eye protection. Where
a student is suffering from conjunctivitis or some other contagious eye infections,
there may, however, be a small risk of disease transmission after wearing eye
protection and more particularly after microscope work from contaminated eye
pieces or rubber eye protectors. Under these circumstances, disinfection of the
equipment would be a wise precaution. [See sections 3.2.3 (Eye protection) and
15.12.3 (Chemical disinfection).]
Disposal of
biological
material
Section 14.6 (Disposal) discusses the hygienic disposal of cage litter, dead animals
and other material used in biology.
Fieldwork
Appropriate hygienic precautions must be observed when coming into contact with
soil, pond water etc and after handling animals caught in traps. See section 17.1
(Fieldwork) for details of particular hazards.
Handling animals Pupils and staff should always wash their hands before and after handling any
animals or cleaning out aquaria and cages. This will be particularly important if
and cleaning
animals, such as terrapins, which may pose a greater risk of the transmission of
cages
microorganisms, are kept in the school. Cages should be disinfected when they are
cleaned out and the soiled litter disposed of hygienically.
Investigations
involving tasting,
eating and
drinking
1
Normally no eating, drinking or smoking will be allowed in laboratories. Where
practical activities require that foods will be eaten or tasted, for example, after
making bread or yoghurt or in mapping areas of the tongue sensitive to different
tastes, these are best conducted in a home economics room. Where this is not
possible and the activities are to continue, particular care must be taken to ensure
that very high standards of hygiene are observed.
See sections 15.2 (Microbiology) and 15.12 (Sterilisation) for details of cleaning procedures and the selection of appropriate
disinfectants.
© CLEAPSS 2006
Investigations
with pupils
1461
In some investigations, there may be a risk of disease transmission from one
person to another when, for example, spirometers, nose clips, mouthpieces, dental
mirrors or clinical thermometers are used. All such equipment must be adequately
disinfected before reuse1. When an investigation involves the safe use of human
body cells or fluid, such as cheek cells2 or saliva3, the chances of cross-infection are
eliminated if each pupil uses only his or her own materials which are placed in
disinfectant immediately after use. In addition, each pupil should be responsible
for washing up his or her own glassware.
In studies of dental hygiene, where pupils are required to brush their teeth, they
should bring in from home their personal tooth brushes; under no circumstances
should these be shared between pupils. Care must also be taken with the source of
any drinking water used in washing out mouths etc; only use a tap which provides
water straight from the mains supply, in case water in the reservoir tank has
become contaminated.
Using chemicals Pupils and staff should always wipe down the bench top after chemicals of any
kind have been used, even if no noticeable spill was caused. This is to guard
against work surfaces becoming contaminated with very small quantities of
chemicals that are dropped but escape attention. If such regular cleaning is not
done, there is a significant risk of pupils, for example, rubbing their eyes, having
inadvertently picked up some of the chemical on their fingers. When a spill occurs,
this must naturally be dealt with as soon as possible, taking appropriate
precautions depending on the nature of the chemical spilt.
14.13.2 Human blood
Section 14.4.1 discusses various aspects of the use of blood. In the event of injuries or
nose bleeds in which blood is spilt or items become contaminated with blood, a sterile
procedure should be adopted when clearing up or disposing of blood-soaked materials.
This procedure can also be used or adapted when handling time-expired blood, if
obtainable, from a blood bank or animal blood from an abattoir.
Cleaning up
blood spills
a)
A solution of sodium chlorate(I) [hypochlorite] disinfectant4 containing at
least 10 000 ppm chlorine should be freshly prepared.
b)
Wearing disposable gloves, wet some paper towels or a clean cloth with the
disinfectant and place over the area contaminated with blood. Then pour over
the towels or cloth more disinfectant so that the contaminated area is
completely soaked with sodium chlorate(I). Leave in contact for at least 15
minutes and preferably for 30 minutes.
c)
Still wearing gloves, remove the towels or cloth and place them under running
water to rinse away the sodium chlorate(I). The disinfected area is then
washed down to remove surplus disinfectant.
d)
After clearing up, towels, cloths and gloves should be disposed of hygienically
and the hands washed thoroughly with soap and water.
e)
Items contaminated with blood should be placed in a solution containing
sodium chlorate(I) as above and left for up to 30 minutes or, preferably,
placed in a heat-resistant plastic bag and sterilised in an autoclave or
pressure cooker5. These can then be disposed of or washed up in the normal
way.
1
See sections 14.5 (Breathing investigations) and 15.12 (Sterilisation).
2
See section 14.4.2 (Cheek cells).
3
See section 14.4.3 (Saliva).
4
For more details, refer to section 15.12.3 (Chemical disinfection).
5
For details of steam sterilisation, refer to section 15.12.2 (Heat sterilisation).
14.14
1462
© CLEAPSS 2006
f)
Care should be taken to avoid contaminating the skin with blood. If this
should occur, disinfect the skin with 70% ethanol or a disinfectant containing
a quaternary ammonium compound/chlorhexidine such as Savlon. Then wash
the affected area thoroughly with soap and water. If sodium chlorate(I) is
splashed onto the skin, wash it off immediately under running water.
g)
It is a sensible precaution to ensure that people with open wounds on the
hands or other exposed parts of their bodies do not come into contact with
blood during any of the procedures described above.
Incubators & other temperature-stabilised equipment
Over 24 hours, the temperature in a school laboratory can vary a great deal, particularly in winter when heating is on only when the premises are occupied. After this
period, the rooms cool quickly, especially those with a large area of windows, and the
temperature can typically drop to 10 °C or even less in very windy or frosty weather,
further at weekends when there is no daytime boost from the heating system. The
opposite problem can occur in summer, with overheating in laboratories facing the
sun, again particularly at weekends when no-one is there to open windows. Apart
from refrigerators and freezers, systems for maintaining small enclosures below room
temperatures are extremely expensive and are not discussed here. However, various
systems for maintaining higher temperatures are used in school laboratories and are
discussed below.
Work requiring a
temperaturecontrolled
environment
A temperature-controlled environment is required in the following biological
investigations:
a)
Maintenance of cultures and other experimental material at closelycontrolled temperatures so that variations with temperature and other factors can
be studied.
b)
Maintenance of stocks of organisms, sometimes at the optimum temperatures
to encourage breeding or at closely-controlled temperatures so that their life
cycle times can be predicted. [Note, however, that it is no longer recommended that most microorganisms should be incubated at temperatures
above ambient although thermophilic bacteria, such as Bacillus stearothermophilus, require an incubation temperature above 60 °C.]
c)
Maintenance of poultry eggs at the required temperature for hatching.
The requirements of particular organisms or operations are summarised in
Table 14.6.
14.14.1 Controlling the environment
Temperature
In everyday usage, the word ‘thermostat’ means any device which attempts to control
temperature and covers energy regulators such as ‘Simmerstats’ as well as true
thermostats. It is important to appreciate the differences between these devices when
considering temperature-stabilised equipment.
True thermostats A true thermostat switches on the electricity supply to the heater when the temperature of the sensor is less than t1 and switches it off when the temperature
reaches t2. The more sophisticated the design of the thermostat, the smaller t2 - t1
becomes and the smaller the temperature fluctuations in the chamber as a whole.
© CLEAPSS 1991
1463
Thermostats have a sensor either inside the chamber or attached to the surface of
the chamber itself. Mechanical devices using the vapour pressure of volatile
liquids, the expansion of liquids or the expansion of metals to open and close electrical contacts are common. In these first two types operated by pressure, the
sensor can be a sealed tube, about 5 mm in diameter, fixed vertically at the back
of the chamber or horizontally under a shelf support or it can be a sealed disc with
corrugated surfaces. Rod thermostats, which work on the differential expansion of
metals, have an Invar rod surrounded by a brass tube; a length of 300 mm and a
diameter of 10 mm are typical dimensions. These rods are often fixed horizontally
near the top of the chamber.
Electronic sensors are smaller than mechanical sensors and are often incorporated
in a small button that can be seen inside the chamber. If no sensor can be seen, it
may be attached to the outside surface of the inner chamber.
Energy regulators Sometimes called ‘Simmerstats’, these are controllers in which the electricity
supply is switched on and off at regular intervals by a small independent heater
affecting a bimetallic strip just behind the control knob which governs the timing
of the intervals. An energy regulator keeps the temperature of the chamber
several degrees (depending on the setting) above ambient temperature, ie, temperature of the surroundings. The temperature of the chamber will vary with
ambient temperature and be steady only when ambient temperature is steady.
Human
intervention
Some disasters caused by low night temperatures can sometimes be avoided if the
organisms or experiments are placed near a low wattage heater at the end of the
afternoon and removed the following morning or if an insulating cover such as a
corrugated cardboard box is used overnight.
Humidity
Humidity levels are quite critical for the well-being of some organisms but, unfortunately, automatic humidity control is very expensive. Where necessary, the humidity in
temperature-controlled enclosures should be checked with a simple hygrometer and
adjusted by increasing ventilation or adding water vapour. This can be achieved by
placing an open dish containing water in the chamber or, if the organisms are likely to
drown, by cotton wool, kept moist by an arrangement like a birds’ drinking fountain.
14.14.2 Incubators
These are the usual method for obtaining a temperature-controlled environment.
Models able to maintain up to 80 °C will be needed for the incubation of thermophilic
bacteria.
These have good insulation and an inner glass door so that the contents can be
Incubators for
higher education observed without their cooling. They have a very sensitive thermostat and possibly a fan to circulate the warm air, both of which help to keep temperature flucetc
tuations in the chamber to 0.1 degree C or less. An incubator of this type, with a
capacity of 75 l, costs between £700 - £1200 at the time of writing.
Incubators for
school
biological work
Available from the main school suppliers, these are without fan circulation but
with thermostats allowing temperature fluctuations of up to 0.5 degree C; a 75 l
capacity model currently costs approximately £500. Many schools may want a
capacity greater than 75 l and so need to purchase a more expensive incubator. It
is important to assess uses carefully before purchase.
Incubatorsterilisers
With these, the greater temperature range from room temperature to 200 oC
allows dry sterilisation as well as incubation. Usually a thermometer can be fitted
through a hole in the top to help set the temperature; the control knob is not precisely calibrated. The thermometer can break if it slips out of its clip or if its limit
is exceeded by switching temperature ranges. The subsequent contamination
1464
© CLEAPSS 1991
by mercury of the aluminium lining is almost impossible to remove. On a Philip
Harris model tested, day-night fluctuations were negligible but there can be
variation of around 2 degree C from place-to-place inside.
Egg incubators
There are two main types of egg incubators: the still-air variety which relies on
convection for ventilation and the forced-air type. In most situations, the number
of eggs to be incubated will not be very large and the still-air design, which is
much cheaper, will normally be quite adequate. Automatic models, with some
form of motor drive that turns the eggs, are also available. Some models allow
much easier observation of the eggs than others; the importance of this in a particular situation should be considered before purchasing. Avoid incubators which
are too small or too large; there is often a minimum order size for fertile eggs and
too many hatched chicks increases the problems of finding homes for them.
Also avoid most egg incubators which are inexpensive; these often fail to meet
current safety standards expected of mains-powered equipment for use in schools.
14.14.3 Other equipment for maintaining stable temperatures
Environmental
chambers
These are aluminium boxes with transparent fronts and perforated zinc shelves,
designed for housing locusts, stick insects and butterfly larvae in the bottom
section, with space for further material above the shelf. Heat is provided by a light
bulb controlled by a thermostat. Temperatures from ambient to around 40 oC can
be achieved with variation within the enclosure of 4 to 6 degree C and day-night
fluctuations around 5 degree C. Some improvements can be made by insulating
the outside of the chamber.
Glasswaredrying cabinets
These are similar to the food-warming cabinets seen on self-service counters. They
are enamelled-metal cabinets with sliding glass doors front and back and wire
shelves. The heater under the base is controlled by an energy regulator and so the
day-night fluctuations can be large. It is difficult to achieve the low temperatures
needed for biological work and variation inside can be as much as 30 degree C.
They are best used for drying glassware!
Locust cages
These are of similar construction to environmental chambers; they are heated
with light bulbs but have no temperature control of any kind. Temperature variations inside and day-night fluctuations will be greater.
Plant propagators These consist of a plastic tray or base which often incorporates a heating element
in which sand or soil can be warmed; plant pots or trays planted with seeds or
cuttings are placed on this. A transparent top keeps the air round the plants
warm. Heaters in propagators are sometimes controlled either by a true thermostat or an energy regulator; day-night fluctuations will depend considerably on
which is fitted. Temperatures reached will depend on the make and model but are
unlikely to be much more than 12 degree C above room temperature; variation
inside will be as much as 5 degree C.
Vivarium
temperature
controllers
These are sold commercially to control the temperature inside a vivarium by
turning on and off a light bulb. One model, supplied by Bio-Pet, reduces the disturbance to animals in the vivarium caused by sudden changes in light intensity;
the light bulb is dimmed rather than switched off when the correct temperature is
reached. A suitable vivarium can be achieved by placing a lid fitted with a light
bulb on top of a glass aquarium; see section 14.14.5 for details of suppliers.
Waterbaths
Temperatures between ambient and 90 oC can be achieved and day-night fluctuations are negligible Some waterbaths are made with built-in stirring devices to
reduce temperature variations but these are expensive and can make the water so
turbulent that flasks etc will fall over unless held very firmly. The temperature
variations in unstirred waterbaths are small enough for biological work.
© CLEAPSS 1991
1465
14.14.4 D-i-y systems
An improvised incubator will consist of a container, a heater and, possibly, a thermostat. The main problem is matching heaters and containers so that suitable temperatures are achieved. Use of a thermostat can enable a more powerful heater to be used
in a container without damaging the contents. Suitable combinations are suggested
below.
Monitoring
temperature
Ideally, a thermometer should be placed so that it can be read without opening the
container but this usually involves boring a hole which can be inconvenient. It is
satisfactory to place a thermometer so that it can be read in situ immediately the
container is opened. Contrary to popular opinion, a thermometer reading does not
begin to change as soon as the temperature of its surroundings alters. A maximum-and-minimum thermometer can give more helpful information especially if
used in conjunction with a second one placed in the room near the container.
Whatever type of thermometer is used, it should be fixed securely.
Electronic sensors, for example a thermocouple or a temperature probe, are useful
because the temperature can be noted without opening the container. The leads
for the temperature-measuring device can leave the container alongside the mains
lead for the heater. It is particularly helpful if, while initial adjustments are made
to a d-i-y system, the temperature-measuring device can be connected to a
datalogger.
m Electrical safety
Wiring must be carried out only by someone competent to do so and any LEA
rules must be followed. Double-insulated cable of the correct current rating must
be used with the line (brown) wires of the heater and the thermostat controlling it
connected in series using a junction box. Do not use insulating tape! It is usually
convenient to put the junction box inside the container as then only one cable
needs to emerge and this simplifies heat insulation. If either the heater or
thermostat has an earth wire, this must be connected to the earth wire of the lead
from the plug.
The rating of the fuse in the plug should be 3 A unless the power of the heater is
greater than 500 W. The whole system should be inspected to check that cables
are securely clamped and that no bare conductors are accessible. Any wires passing over sharp edges should be protected by grommets.
Any lamp holders used should be brass batten-holders fitted with an earthing
point which must be connected to the mains earth.
Heaters
The wattage required depends on the desired temperature, on the capacity of the container and on its insulation. As a rough guide, for small containers 1 W per litre is
required and for larger ones 0.75 W per litre. Some animals such as locusts benefit if
day and night conditions are simulated. This can be achieved by a combination of a
light bulb with a heater, such as a tubular heater or a seed tray heater.
Soil/air warming
cable
This is a very useful and inexpensive heater for a biology department to have. It is
available in several lengths, the shorter ones of 6 m (75 W) and 12 m (150 W)
being the most useful for schools. (NB A long warming cable should not be cut.)
These cables can be laid in a zig-zag over the bottom of a container and also held
in rows round the sides using cable clips. They are available from garden centres
and some laboratory suppliers.
1466
© CLEAPSS 1991
Tubular heaters
These are designed for use in greenhouses, garages, airing cupboards etc. The
diameter of the tube is 50 mm and they are available in several lengths; the
shortest is 300 mm but probably the most useful size for this application is
600 mm which would fit diagonally across an underbench cupboard. Tubes are
rated at either 60 W or 80 W per foot. They can be obtained from heating suppliers
eg Tube Heat Ltd.
Light bulbs
These are very useful for an emergency. Standard light bulbs are easily available
in a range from 15 to 150 W; 15 and 25 W pygmy bulbs are particularly useful in
small containers. Pygmy bulbs are made with either standard or small bayonet
fittings and appropriate holders must be used.
The quickest way to use a light bulb is to position it outside the container, eg with
an Anglepoise lamp outside a glass aquarium, or to hang it from the top of a
container (single insulated twin flex should not be used). This can, however,
produce very uneven heating unless the bulb is near the bottom of the container.
It is better to use a batten-type bayonet fitting screwed to a piece of wood so that
the light bulb is standing upright at the bottom of the container.
The life of a light bulb, roughly 12 weeks continuous use if reasonably ventilated,
can be extended if two bulbs are wired in series so that they run below normal
temperature; then the power of each will be roughly 2/3 nominal value; ie 65 W
instead of 100 W.
When light bulbs are used as heat sources, even when connected in series, they
emit much radiation which contributes to the uneven heating associated with this
source. If a shield made of aluminium foil, for example, is placed between the
bulbs and the items being heated, the heating is likely to be more even.
When using light bulbs special care must be taken to avoid spills of liquids.
Single seed-tray
heaters
Propagator heaters, described in section 14.14.3, can be used with containers
other than the seed trays for which they were designed. Their main advantage is
that they are rigid and Petri dishes can be stacked directly on them or they can be
placed under containers such as an aquarium or a biscuit tin. Similar heaters are
also sold as ‘heat pads’ for warming vivaria, for example by Bio-Pet.
Containers
A wooden under- The capacity can be approximately 200 litres. Any wooden shelf should be rebench cupboard moved and replaced by a metal mesh which is open but sufficiently rigid; as a last
resort chicken wire fixed to a wooden frame can be used, although some supports
across the middle may be needed. It is best to drill a hole near the bottom of a side
for the mains lead, rather than squashing it in the edge of the door.
One arrangement we tried was to screw the thermostat to the inside back of the
cupboard, making the device more secure from casual tweekers and with the
‘straight on’ view simplifying setting the required temperature. We used 12 m of
soil/air warming cable, spread in a zig-zag on the bottom and with three rows
fixed to the sides with cable clips. The junction box was fixed inside the cupboard
and the one mains lead passed through a hole drilled near the bottom of the
cupboard. Tests suggested that it should be possible to maintain a temperature of
at least 25 oC on all but the coldest nights. Temperature variations within the
cupboard and day-night fluctuations were found to be negligible.
A wooden drawer This gives a capacity of approximately 50 l. A drawer is really too shallow to allow
low temperature variation to be achieved and only a flat heater, eg a soil warming
from an undercable or seed-tray heater, can be used, not light bulbs. A lid can be made of chip
bench unit
board, several layers of corrugated cardboard, expanded polystyrene sheet (20 mm
thick or more if possible) or any combination of these. If a drawer is used within
its unit, care must be taken not to squash the mains lead.
© CLEAPSS 1991
A picnic box or
cold box
1467
A typical capacity is 13 l or more. One problem with these containers is that,
unless a hole is drilled through the wall, the mains cable prevents the lid from
closing properly. However, the gap can be filled with adhesive foam strip (draught
excluder), cotton wool etc. Such a container is quite effective because it is quite
deep for its capacity and well insulated. The purchase price depends on the size
but picnic boxes can often be obtained at jumble sales for a pound or two.
The bulbs are mounted horizontally as shown so that the shelf can be low to give
as much working capacity as possible. This is made of expanded metal but perforated zinc etc would do instead. It can be fixed to the wooden base of the bulb
holders or separately supported above the bulbs by being bent into an inverted ‘U’.
Tests have shown that a steady temperature about 13 degree C above ambient can
be achieved which would be rather high when the school heating is working. It
could be switched on only at night, perhaps by a time switch if one were available.
Any condensation which occurs can be reduced by increasing ventilation but then
there is a risk of a significant drop in temperature.
Side view
Cotton
wool
Mains
lead
Junction
box
Aluminium
foil heat
shield
End view
shelf
16 watt pygmy
bulbs
Wooden base
A plastic or
glass aquarium
Some aquaria are supplied with hoods but tops made of wood, expanded polystyrene etc are effective for this purpose. The heater lead can be passed through
the aperture in the lid provided for feeding and for air lines. D-i-y tops can be cut
to provide a suitable entry point. The main advantage of this container is that the
contents can be seen without opening the top; this could be a useful feature with
some organisms. See section 14.14.5 for details of converting an aquarium into a
heated vivarium.
A biscuit tin
This gives a capacity of approximately 5 l. This is quite a good container to use in
conjunction with an external heat source such as a seed-tray warmer because it
readily conducts heat to the contents. However, heat losses are great and
insulation is really essential. This container is small enough to be packed inside a
cardboard box with expanded polystyrene packing chips; this is quite effective
insulation. If the heater is inside, it is difficult to fit the lid of the tin over the
mains lead; again, it may be more effective to drill a hole in the side of the tin.
Additional
insulation
Expanded polystyrene, either solid or as chippings, and corrugated cardboard
have already been mentioned. Expanded polystyrene sheet can be cut with a
bread knife and pieces joined with wide brown parcel tape. Chippings can be used
loose or in a plastic bag. NB Polystyrene becomes sticky above 60 oC. Also
valuable for insulation is Aircap bubble polythene, a double-layer sheet of
polythene with pockets of air in between. often used in greenhouses.
1468
© CLEAPSS 1991
Thermostats
Rod thermostats Negligible temperature fluctuations are obtained with bi-metallic types sold for
use with soil/air warming cable. Being at least 600 mm long, they will, however,
not fit in some containers. These thermostats need to be mounted horizontally
near the top of the container. Currently available from Thermoforce Ltd.
Smaller rod thermostats are available, really designed to control the temperature
of boilers and hot water systems. They are likely to be less sensitive but worth
trying for some applications. Available from laboratory and heating suppliers.
Room
thermostats
Many makes are available and can be mounted vertically on a piece of wood and
hung or fixed near the top of the container. Good air circulation is necessary:
satisfactory results can be obtained in an underbench cupboard with negligible
fluctuation but, in smaller containers, 5 degree C fluctuation is typical.
14.14.5 Miscellaneous tips
Siting equipment, These should be sited in as even a temperature as possible, ie, away from windows, doors, radiators and hot pipes. It is sometimes helpful to erect temporary
organisms and
hardboard shields to protect experiments etc from draughts.
experiments
The siting of stocks of organisms needs special consideration in summer when,
over weekends, sunny rooms become very hot. It is worth consulting those in
charge of the security of the premises to see whether it is possible to leave some
top windows open.
Where a temperature close to room temperature is required, the equipment must
be sited with particular care. To achieve control, the equipment needs to be in an
environment with an ambient temperature at least 5 degree C below the desired
level. For example, if the thermostat is to be set at 20 oC, then the equipment
should be placed where the ambient temperature is 15 oC or lower for the
adjustments to be made. Possible locations include a bookstore, a caretaker’s
garage or even outside a ground floor room with an extension lead through a
window to the power supply. It is always best to check the setting overnight with
a maximum-and-minimum thermometer placed in the equipment or, better still, a
thermocouple linked to a datalogger.
Reducing
mischievous
adjustment of
control knobs
Some equipment has control knobs which can be locked with an Allen key. One
solution with equipment without this facility is to loosen the grub screw and
remove the knob. Another is to turn the equipment so that the control knob is at
the back towards the wall. A small mirror can be used to make legitimate adjustments if it is inconvenient to turn the equipment when fully loaded.
Conversion of old Suitable lids with a ventilation panel, sliding door and space for a lamp socket are
aquaria to vivaria available from Bio-Pet and Philip Castang. They are made for standard sizes of
aquaria but other sizes can usually be made to order.
D-i-y lids can be made from a wooden frame with chicken wire or smaller mesh
plastic Netlon available at garden centres.
© CLEAPSS 1991
1469
Table 14.6
Organism
Temperature requirements and ways of meeting them
Requirement /°C
Ideal equipment
Acceptable but
less effective
Comments
Hens’ eggs
38 ±1
Observation
incubator
Incubator
or incubator/
steriliser
No other commercial equipment tested gives
a sufficiently high, steady temperature.
D-i-y difficult.
Locusts
28 to 33
Locust cage,
because a diurnal
cycle is possible
Environmental
chamber
No other commercial equipment has lighting
and egg tube facilities and is secure enough
for hoppers. A d-i-y system may be possible.
- egg pods
30 ± 4
Incubator or
incubator/
steriliser
Top of environmental chamber
At 32 °C: 11 days for hatching; at lower
temperatures, hatching is longer. To observe
when hatching, remove top 10 mm of sand
and cover with a Petri dish which helps to
preserve humidity.
Fruit flies
- observation of
genetic crosses
25 ± 0.5
Incubator or
incubator/
steriliser
Observation
incubator or
d-i-y system
with thermostat
25 °C gives 14 day life cycle, lower temperatures lengthen it. Curled -wing character is
somewhat temperature-dependent. 27 °C
recommended for last day of pupal life.
- for maintenance of stocks
19 to 21
As above
As above or any
propagator or
environmental
chamber set low,
or just in the open
lab
Below 10 °C, flies will die. Above 28 °C, flies
become sterile.
Flour beetles
- for genetics
30 ± 1
Incubator or
incubator/
steriliser
Observation
incubator or
d-i-y system
with thermostat
65% relative humidity recommended.
- for maintenance of stocks
Room
temperature
Bacteria
Ambient to 30
except for
thermophilic
species (65 °C)
Incubator or
incubator/
steriliser
Observation
incubator or
propagator
with thermostat
Yeasts and
moulds
25 to 30 ± 1
Incubator or
incubator/
steriliser
Observation
incubator or
propagator
with thermostat
Cabbage white
butterflies
18 to 25
during daytime
Large
insect cage
Keep in open lab unless temperature falls
below 5 °C. Beetles will not survive frost.
Many organisms will grow satisfactorily, if
more slowly, in this temperature range.
Incubation temperatures around 37 °C should
be avoided because they favour organisms
well adapted for survival in humans. Thermophilic bacteria requiring very high temperatures should not pose a significant risk.
Below 13 °C, adults will live but not lay and
eggs will not hatch. Over 30 °C eggs are
infertile.
1470
© CLEAPSS 1991
Table 14. 6 Temperature requirements and ways of meeting them (continued)
Organism
Requirement /°C
Ideal equipment
Acceptable but
less effective
Comments
Stick insects
22 to 32
Insect cage or
locust cage
Plastic sweet
jar
Room temperature is normally sufficient but
temperatures below 10 °C are undesirable.
Insulate cages or stand in propagators on low
setting for cold nights and weekends. Some
species need high humidity.
Crickets
25 to 35
Deep dustbin
- eggs
32
Any deep, closed
container with
some heating to
protect from
frost
Need very high humidity, hence lid. 250 ml
beaker containing damp cotton wool on which
eggs will be laid aids humidity. Immobile at
low temperatures. Die below 5 °C.
Reptiles
21 to 27
Vivarium
Aquarium with
suitable lid
Reptiles need a basking light, preferably near
one end of vivarium, so they can vary their
temperature. They need lower night temperature so use a time switch. In winter, extra
warmth needed, perhaps a second low
wattage bulb. Some species need uv
radiation.
Enzyme
experiments
Ambient to
50 ± 2
Waterbath
Incubator,
incubator/steriliser or observation incubator
For much work, 35 °C is sufficient when a
propagator with a thermostat might suffice.
For elementary work, a beaker of water with
hot or cold water added as required may be
adequate.
Digestion
experiments
25 to 37
Waterbath
Any equipment
with thermostat
Keeping agar
molten
50 ± 4
Waterbath or
incubator
Incubator/
steriliser
Action of
auxin on
coleoptiles
25 for 48 hours
Incubator or
incubator/
steriliser
Any equipment
with a thermostat
Pasteurisation
experiments
63 ± 1
for 30 minutes
Waterbath or
incubator
Incubator/
steriliser or
drying cabinet
Respiration in
woodlice etc
Ambient to 25
± 0.5
Waterbath
Incubator or
incubator/
steriliser
Seed germination experiments
4 to 30 ± 2
Propagator
with thermostat
Observation
incubator
Drying soilsamples
90 to 100
Incubator/
steriliser or oven
Drying cabinet
Light must be excluded.
Refrigerators for low temperatures.

Laboratory Handbook - Section 14 - MAINLY BIOLOGY, A - J

Transcription

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