Below are the first 10 and last 10 pages of uncorrected machine-read text (when available) of this chapter, followed by the top 30 algorithmically extracted key phrases from the chapter as a whole.
Intended to provide our own search engines and external engines with highly rich, chapter-representative searchable text on the opening pages of each chapter.
Because it is UNCORRECTED material, please consider the following text as a useful but insufficient proxy for the authoritative book pages.
Do not use for reproduction, copying, pasting, or reading; exclusively for search engines.
OCR for page 46
V FshysIcal FacilNIes
A. RELATION TO NATURAL HABITAT
Two choices are offered when the decision is made to use a wild living ani-
mal in the laboratory and to develop it as an animal adapted to the labora-
tory. The first choice is to provide a laboratory habitat that is, in as many
particulars as possible, a close mimic of the natural habitat. The second is
to develop a laboratory habitat in which the animals live and which is at
the same time compatible with the laboratory environment, minimizes
labor and material costs, and may be managed on the basis of principles
already familiar to animal husbandry personnel.
Many amphibian hobbyists have chosen the first alternative. With ani-
mals from a known and limited geographic territory, it is possible to con-
struct reasonable facsimiles of a particular natural habitat in the labora-
tory. Usually, this is expensive, either in materials (container fabrication,
water, soil, plants, inserted objects), or in labor, or both. This is difficult
to do, however, when designing quarters suitable for any representatives
of a species whose range places them in many quite different habitats.
Also, such naturalistic quarters usually will not accommodate animals in
densities that significantly exceed those found in nature.
For these reasons the second alternative was chosen, and facilities are
described that are only now under critical test (Boterenbrood, 1966;
Frazer, 1966~. Many of the details will change as experience broadens.
It is asked that experiences that may aid in satisfying the requirements
of this second choice be communicated to the institute of Laboratory
Animal Resources.
The principles for the second alternative are as follows:
. No matter how varied the environments within which a given species
is found, each of those environments contains common features that make
46
OCR for page 47
47
it possible for the species in question to exploit it. Only these common
and essential features need or should be incorporated in the design of
laboratory quarters for the species.
· Laboratory quarters and management protocols for the species in
question should not require unusual expenses and should not require train-
ing that is totally different from the typical training experience of person"
net in animal facilities.
B. THE AMPHIBIAN QUARTERS
1. General Description
Amphibians should not share quarters with mammals or birds, but may be
in rooms with aquaria containing fish or other aquatic forms. The high hu-
nudity of amphibian quarters and the optimal temperatures for these ecto-
therms are not usually compatible with the requirements for endotherms.
The work area requirements for an amphibian unit remain the same re-
gardless of the size of the unit. The following describes a unit suitable for
the maintenance of several thousand or more animals of several species.
Smaller units should contain equivalent work and animal areas even
though these areas are not in separate rooms. The major difference be-
tween smaller and larger units is that smaller units may not have as many
options for maintaining animals at a variety of temperatures and under
various lighting regimens. Facilities for dealers will differ in size and in
the proportion of areas for temporary holding as distinct from long~term
animal culture.
As noted below, the amphibian unit should be provided with inflow
and outflow entrances to a suite of rooms or functional areas that include:
· The animal rooms
Breeding area
Isolation quarters
General laboratory area
Examination and autopsy area
Insectarium
General service area
Storage area
Information control area
Office area
a. Isolation Quarters
Isolation quarters for newly arrived animals are advisable. Because of the
difficulties inherent in regulating temperatures for different species, spe
OCR for page 48
48
cific areas within a room should be set aside for new arrivals. The new
arrivals should be kept relatively isolated from other animals in the room.
Since amphibians do not introduce as much contamination into the envi-
ronment as do mammals and birds, their physical isolation is satisfied by
placing them on the lowest cage rack level; in this position, the effluent
water from their containers will flow directly into the drain.
Isolation and acclimatization for an incubation period is desirable,
particularly if juveniles or adults are brought into the laboratory. Isola-
tion not only provides disease protection to existing laboratory stocks
but also gives the new arrivals a chance to adapt to their environment
with a minimum of disturbance. These animals often will not feed for
several days and will be easily startled by routine care activities. Mini-
mizing activity around the isolation quarters for several days will reduce
mortality.
If symptoms of disease appear, treatment and container care as dis-
cussed in Chapter IX should be followed. Diseased amphibians in isola-
tion should not be incorporated into existing stocks until the disease
has been controlled for at least 2 weeks.
b. Heating, Ventilation, and Size Specifications for Rooms
Because of the several temperature requirements for different amphibians,
both during hibernation and periods of activity, the unit should be pro-
vided with animal rooms at different temperatures. This includes rooms
for adult and larval amphibians and for insect culture; the size of each will
depend on the size and objectives of the animal colony. To facilitate ser-
vicing, however, these should be at least "walk-in" size.
For hibernation of northern species, temperatures of 0-2 °C (32-35 °F)
and 2 - °C (35-39 °F) must be available. This requirement can be met in
two ways: A sufficiently large room held at approximately 18 °C (64 °F)
could contain fiberglass circulating refrigeration units to attain the tem-
perature desired (see Section C.5~. Alternatively, two hibernation rooms-
one maintained at 0-2 °C (32-35 °F) and the other at 2 - °C (35-39 °F)-
could be used. The latter would have the advantage of allowing the use of
hibernation containers for smaller groups of animals (see Section C.6.a).
For hibernating animals from the intermediate geographic ranges, for
conditioning northern animals to hibernation, and for maintaining cer-
tain of the urodeles, a room maintained at 7-12 °C (45-55 °F) is ad-
visable.
For certain larvae and the adults of other urodeles, a room of 18-20 °C
(65-68 °F) is advisable. For axolotls and for R. pipiens in the process of
acclimatization to hibernation, a room at 2~22 °C (68-72 °F) is needed.
OCR for page 49
49
For active R. pipiens and other species, rooms at 22-25 °C (72-78 °F) are
optimal.
For tropical species and for use as an isolated insectarium, two rooms
should be available at 26-30 °C (78-86 °F). The insectarium must be pro-
vided with a strong vent fan, high-capacity air intake, and a thermostati-
cally regulated heater to prevent undue temperature fluctuations. Such
venting is needed to minimize odors and to reduce the possible occur-
rence of insect allergies among personnel (see Chapter X, Section B.2~.
c. Description of Ancillary Rooms
The breeding and general laboratory rooms [maintained between 20 and
22 °C (68-72 °F)] need equipment typical of a biological laboratory as
well as extra shelf space for holding pans of embryos from fertilization
to hatching. In addition to the usual equipment for autopsy and exami-
nation, this room needs a small refrigerator for the storage of carcasses
until time of autopsy and disposal.
The equipment for the insectarium as described above, depends on the
insect species under culture. An ample storage and general service room
should be provided for the storage of cages, food, sundry supplies, and
washing equipment as well as simple shop tools. It will need a refrigerator
and a deep freeze. In a large unit the washing operation should have a room
separate from the storage and general service functions. An automatic tun-
nel washer is advisable, but care must be exercised to test the toxicity of
detergents used in such washers. Larvae may be particularly sensitive to
detergent "buildups."
Equipment in the office and information control center will depend
on the nature of the records and the information to be handled (see
Chapter VIII). Adequate space for file cabinets and shelves for record
books is mandatory. The room should be suitable for computer terminal
installation. In addition, this room or an adjacent room should contain
photographic equipment and space to store preserved animal specimens.
Each of the "wet" rooms should be equipped with sinks that have heavy"
duty industrial capacity disposals and with facilities that meet standards
for animal waste disposal.
d. General Specifications
Other specifications for the amphibian quarters include hot and cold water.
steam lines for cleaning, plentiful waterproof electrical outlets at 1 10 V,
gas, and high-pressure air and vacuum lines. The floors must be designed
to permit thorough cleaning, minimize slipping, and allow for good drain
OCR for page 50
50
age. Lighting adequate to the function of each room must be provided as
discussed below. The distribution of these facilities to the several rooms
will depend on the geometry of the suite and the uses of the several rooms.
Each room associated with the amphibian quarters should be made as
insectproof as possible because insects may inadvertently escape into the
room when insect or amphibian enclosures are opened. Insect proofing
should include special baffles and sealing materials around doors. The
arrangement used for weatherproofing doors is suitable. Doors should be
equipped with self~closing devices to ensure prompt and effective closure.
The amphibian quarters should be provided with entrance ways with two
sets of doors that form an entrance chamber-another aid in the control
of escaped insects. Floor and ceiling moldings should be carefully inspected
for possible routes of insect escape, as should the points of entrance of
pipes, wiring, etc. Measures must be taken to control insects that have es-
caped, but insecticides must not be used as they are a danger to the food
insects and the amphibians. Cleanliness is the best control, but it may be
aided by the appropriate use of flypaper.
Floors, walls, and ceilings should be of water-resistant material to per-
mit hosing or steam cleaning at regular intervals. One of the major con-
taminating arthropods in the insectarium and the amphibian quarters is
the spider, which finds the environment highly compatible because of the
available food. The only method to control spiders is cleanliness. Other
contaminating insects include parasitic Hymenoptera (wasps) and Diptera
(flies). These, too, can only be controlled by cleanliness and care to pre-
vent their introduction from the outside. In one amphibian installation,
bats, thriving on the insect population, have constituted a nuisance, even
during the winter months.
2. Environmental Control
a. Water
The water supply is most critical to a successful amphibian colony. For
aquatic forms, e.g., anuran larvae and the aquatic urodeles, the require-
ments for water quality are as critical as those for fish; for the terrestrial
and semiterrestrial juvenile and adult forms, water remains an important
component of the environment. Thus, though critical tests have not been
published, we recommend that water standards for both larvae and adults
be held within the limits prescribed for fish. Although we review here some
of those standards that seem particularly applicable to amphibians, we
recommend reference to fish standards (Committee on Standards, in press)
and texts on the engineering and biological aspects of facilities for the
OCR for page 51
51
long-range and large-scale maintenance of fish (Spotte, 1970; Clark and
Clark, 1971; Bardach et al., 1972~. McKee and Wolf (1963), American
Public Health Association (1965), and Culp and Culp (1971) are valuable
references on the evaluation of water quality.
Water quality is highly variable between geographic locations and is af-
fected by its source, quantity, method of transport, type of food placed
in it, and amount of waste released into it (Bennett, 1962~. In particular,
differences may be expected between water from subterranean and surface
sources, the latter more commonly possessing deleterious characteristics.
The quantity of water consumed depends on the size of the facility, the
number and types of animal and the flushing efficiency of the animal con-
tainer (see Sections C.2, 4, and 6~. Since container designs have not been
standardized, the quantity of water needed for proper care of amphibians
must be determined for each facility. Water should be available in quan-
tities greater than use expectation. Where adequate supplies of running
water of high quality are not available, facilities for recirculating water
are essential (Spotte, 1970; Clark and Clark, 1971; Cullum and Justus,
1973~; the capacity of such systems for supplying large facilities, however,
is limited by the economics of filtering, treating, and pumping used water.
In view of these qualitative and quantitative differences between water
supplies, the quality and flow rates of water should be monitored and the
water treated to ensure qualitative acceptability. In planning the water
supply, care should be taken to ensure its quality with respect to:
Alkalinity and hardness as CaCO3
Ammonia and other nitrogen compounds
Carbon dioxide
Chlorine
Fluorides
Heavy metals
Microorganisms
Oxygen
pH
Polychlorinated biphenyls (PC B) and other toxicants from plastics
Toxicants
Municipal systems are a variable source of water. For example, in a city
with several wells and surface sources, it is not uncommon to shift between
these sources on a seasonal or even daily basis. Consequently, water chem-
istry should be monitored regularly. Acceptable procedures can be found
in the publication prepared by the American Public Health Association
(1965~. An inviolate schedule for such analyses should be established, and
OCR for page 52
52
a specific member of the staff should be assigned responsibility for this
service. Remember, however, that such monitoring can never cover mea-
surement of all possible changes; changes in water quality, especially in
the northern states, may be pronounced at the time of winter freezing
and spring thaw. The changes may involve organic contaminants not
normally detected by routine water quality assays. It is extremely impor-
tant to note that such sudden changes in water quality may be highly dele-
terious when mating procedures are being conducted. Thus, as described
in Chapter VII, we recommend that artificial media formulated from dis-
tilled water be used in containers for mating, artificial insemination, and
early development.
( 1) Alkalinity and Hardness as Calcium Carbonate Total alkalinity
(total ionic strength) and hardness should be maintained between 150 and
250 mg/liter, compared with a 60-120 mg/liter standard sometimes recom-
mended for public water supplies. The addition of food to rearing con-
tainers of larval amphibians should adjust the hardness and alkalinity values
in the water and provide the larvae with the necessary minerals. If alka-
linity or hardness must be adjusted because of deficiencies that may occur
in some public water supplies or in places such as some of the mountain
states or because of excesses that clog valve systems, water treatment
specialists should be consulted. Rather sophisticated equipment is avail-
able, but even then it will be necessary to maintain tight control over
the system.
Well water may be high in iron and a variety of salts and deficient in
oxygen.
(2) Ammonia and Other Nitrogen Compounds Concentrations of
ammonia above 0.2 mg/liter as nitrogen are detrimental to fish and may
also affect amphibians. Ammonium carbonate and ammonium hydroxide
form in waters high in carbonates. At 4 mg/liter these compounds are toxic
to fish and can cause stressful pH changes. Thus, in recirculating water sys-
tems, ammonia must be kept at a minimum, especially in hard water.
Nitrates and nitrites are produced by bacterial decomposition of organic
materials. Values for these compounds should not exceed 0.3 mg/liter as
nitrogen; in the presence of phosphorus and wide spectrum lights algae
growth, which may clog water pipes, will be promoted. Although prob-
lems with these compounds occur most frequently in recirculating sys-
tems, they will also occur in those areas where public water supplies are
high in nitrates. Water should be checked for nitrates before use as they
may be harmful to both larval and adult amphibians.
In flow-through systems the frequency of flushing or the rate of flow
OCR for page 53
53
must be adjusted to prevent the accumulation of waste products and the
bloom of putrifying bacteria whose actions have a variety of deleterious
consequences. If such adjustment is difficult, it may be necessary to use
"conditioned water" systems in which nitrifying bacteria control am-
monia levels and in which populations of putrifying bacteria are depressed.
An excellent account of the dynamics of well-established conditioned
water systems is available in Atz (1971~. Poorly defined "control agents"
that seem to regulate R. pipiens populations (Richards, 1958, 1962; Rose
and Rose, 1965; Gromko et al., 1973) may accumulate in noncirculating
water systems and limit the density of the tadpoles that may be cultured.
If nitrogen compounds must be removed, water treatment specialists
should be consulted. However, it will be difficult to control nitrogen com-
pounds without affecting other water quality characteristics. The water
supply system may need reconstruction to obtain desired qualities.
(3) Carbon Dioxide Carbon dioxide should not exceed 5 mg/liter.
It is doubtful that fish, and possibly amphibian larvae, can survive long
periods exposed to 12 mg/liter of carbon dioxide.
In a well-aerated, flow-through system carbon dioxide is unlikely to
reach detrimental levels. Increases in carbon dioxide, however, may de-
press pH values below desirable levels [see Section B.2.a(9) for a further
discussion of pH] .
(4) Chlorine Both chlorinated and nonchlorinated water must be
available. Close attention to chlorine levels is needed as chlorine in public
water supplies often will exceed lethal tolerance limits for aquatic am-
phibians.
Aquatic Water supplied to aquatic, gill~breathing larvae and
adults or to hibernating or skin-breathing adult amphibians must be free
of chlorine. Although some larval amphibians can tolerate chlorine levels
as high as 3.8 mg/liter over a period of time, growth and other physio-
logical processes may be affected. Thus, concentrations should be well
below this level. Activated charcoal filters, aeration or sodium thiosulfate
easily remove chlorine. For small operations, holding the water in tanks
with large surface areas, bubbling air through the water, or agitating the
water will remove the chlorine in a few hours. For intermediate-size op-
erations, sodium thiosulfate (6-8 mg/liter of water) can be added to large
reservoirs or metered into continuous flow systems. However, control of
the sodium thiosulfate level is essential as concentrations of S mg/liter are
known to be toxic to some fish.
For large facilities, e.g., 20 gallons of water/in or greater, cylinders of
OCR for page 54
54
activated charcoal can be installed directly in the water supply system.
Most commercial water treatment services can supply, install, replace, and
service the charcoal cylinders for a nominal fee.
Copper chloride at 9 mg/liter has been reported as toxic to fish. Thus,
copper water lines should be avoided; if this is not possible, water flowing
through them should be closely monitored (see below).
Terrestrial The presence of chlorine in water provided for
lung-breathing amphibians will retard bacterial growth and is thus bene-
ficial. Nonhibernating adult R. pipiens can tolerate levels between 4 and
6 mg/liter and levels as high as 12 mg/liter for Tort periods (Kaplan, 1962~.
At the Louisiana State University facility, R. catesbe~ana also tolerate
4 mg/liter. Higher levels have not been tested.
Because chlorine may be lost between the chlorination plant and the
point of water usage, it may be necessary to meter chlorine into the water
supplied to adults to control the levels of microflora. Chlorine gas is ex-
tremely dangerous and should be handled only by trained personnel. A
high-quality flow meter is essential for accurate metering of chlorine into
the water supply, although such meters require regular maintenance to
ensure accurate delivery.
Amphibians such as R. catesbeiana-which tolerate chlorine but are fed
underwater such living foods as fish or earthworms that cannot tolerate
chlorinated water-must be handled differently with regard to chlorinated
water. Their water supply must be shifted from the chlorinated to the de-
chlorinated line at the time of feeding.
(5) Fluorides Concentrations of fluorides should not exceed 1.5
mg/liter (Kaplan et al., 1964~. In northern climates, concentrations in
water supplies may slightly exceed this value.
(6) Heavy Metals Heavy metals such as zinc, copper, mercury, and
lead may enter food or water systems from many sources and must be
evaluated before amphibians are reared.
Zinc may be leached from galvanized pipes, copper from copper or brass
pipes, etc. Copper is toxic to gill-breathing organisms (see above), and zinc
is known to be toxic (Pickering and Vigor, 1965) and to accumulate to
lethal levels when fish are exposed to ZnCl2 (McKee and Wolf, 1963~. A1-
though the toxicity of these metals to amphibians has not been evaluated,
the aquatic forms may be at risk; pipes in the water system should be made
from black iron or high-density polyethylene or polypropylene or nylon
[see also Section B.2.a(10~]
OCR for page 55
55
Well water may be high in iron. Upon aeration the iron normally pre-
cipitates and is thus nontoxic. However, large quantities of precipitated
iron may clog sensitive water valves or enhance growth of iron bacteria
that may, in turn, clog valves or deplete oxygen in the water.
Municipal water departments may add copper sulfate to water-supplies
to control algal growth, particularly in the fall and spring. Copper sulfate
is an inhibitor of tadpole growth. It may be removed by adding versene
(EDTA) at 50 mg/liter of water (Richards, 1958~.
(7) Microorganisms Some aspects of the role of bacterial flora in
water supplies are noted above. Fecal coliform densities should not exceed
2,000/100 ml and total coliform not exceed 20,000/100 ml.
(8) Oxygen Gill-breathing amphibians must have an adequate sup-
ply of oxygen to survive normally. Since the oxygen requirements for
aquatic stages of amphibian species have been poorly documented, the
oxygen requirements for fish should be followed. For warm water fish,
oxygen levels should not fall below 5 mg/liter and for cold water fish,
8 mg/liter. This suggests that larval stages of amphibians from northern
climates may have higher oxygen requirements than the same species
from more temperate regions.
Though gills are replaced by lungs during metamorphosis, oxygen should
still be maintained at the recommended levels to prevent other complica-
tions in water quality. Should the water become anaerobic, bacterial pop-
ulations will increase, and the chance of disease outbreak increases. Also,
ammonia levels may increase to toxic levels.
Well water, depending on its source, may be either deficient in oxygen
or contain an excess of oxygen that leaves in a gaseous form as the water
warms. In either case, the water should be stabilized by aeration before use.
(9) pH A pH value outside the range of 6.5-8.5 may be detrimental
to amphibians that remain in water for extended periods, and there is evi-
dence that tadpoles of some Rana species develop best if the pH is around
6.5. However, many amphibians have evolved in natural environments with
lower or higher pH levels and may have other optimum levels. For wild-
caught animals, especially eggs or larvae, pH levels similar to those of the
environment in which the animals were collected should be maintained.
Changes in ammonia and carbon dioxide concentrations can cause
changes in pH isee Sections B.2.a.~2) and (3~] . In cases of pH values being
depressed by excess carbon dioxide, correction can be attained by the ad"
dition of calcium sulfate or sodium hydroxide. Conversely, the water in
OCR for page 56
lo:
am
some public supply systems may reach pH values as high as 9.5-10.5 be-
cause of the source or method of treatment. These pH values stress am-
phibian larvae; this state can be reduced by the monitored addition of
acetic acid, although caution should be exercised as '`iron bacteria" may
clog pipes at lower pH values.
(10) Polychlonnated Biphenyls {PCBJ and Other Toxicants
Mom Plastics The ease and favorable cost of plastic piping, containers,
and instruments recommend their use in many applications in amphibian
quarters. However, they are not without danger, especially when in con-
tact with the water supply. Phenolic and acrylic plastics may contribute
significant levels of polychlorinated biphenyls to the water. The toxicity
of these substances has been well documented for living systems and
should be avoided. Pliable plastics contain up to 40 percent by weight of
plasticizers, some of which are volatile. Phthalate esters of various kinds
may be leached into water from these plasticizers. These substances have
known toxic effects and should be avoided (Napier, 1968; Jaeger and
Rubin, 1973; Krauskopf, 1973~. Some plastics incorporate fungicides.
In the absence of tests evaluating the effects of these fungicides on am-
phibians, these plastics should be avoided.
Thus, we recommend that, where plastic piping is used to avoid copper
contamination, those made of high-density polyethylene, polypropylene,
or nylon be used. If plastic containers for embryos and larvae are used,
these should be of rigid plastics with reduced plasticizer content.
(1 1) Toxicants Toxicants are too numerous to describe here in de-
tail, but any source of water or food contains potentially toxic substances.
Insecticides are most likely to occur in commercial food preparations and
should be quantified and possibly removed (Stober and Payne, 1966~.
Amphibians normally are tolerant of the insecticide concentrations found
in commercial feeds. However, the insecticides may accumulate if appro-
priate precautions are not taken and may be detrimental in breeding colo-
nies where there is risk of pesticide accumulation in fat-rich ova. Cooke
(1971) has reported the toxic levels of pesticides for R. temporana and
Bufo larvae. It is not yet possible to define toxic levels of these substances,
particularly in the absence of information on their synergistic action.
Common table salt is widely used as a general treatment for diseased
fish and amphibians. Care should be exercised in use of such salt treat-
ments as amphibians, particularly juveniles and adults, rapidly absorb
these potentially lethal salts. Tolerance limits for the many species are
not known, and it is best to avoid the use of salts on a colony of amphib-
ians without first obtaining tolerance limits for a few individuals.
OCR for page 64
64
coldrooms as hibernation enclosures. These are more fully described in
Section C.6.a.
Regardless of the equipment selected, it is most important that am-
phibians that hibernate under ice be in water deep enough to cover them,
Mat temperatures be maintained between 2 and 3 °C (36-37 °F) and
not above 4 °C {39 °FJ, that the water be well aerated, that low-intensity
light levels be maintained for 8-10 h, and that agitation of the animals be
held to a minimum.
6. Enclosure Designs
In the absence of definitive standards for amphibian enclosures, this section
describes several aspects of the housing systems now in use. It is hoped that
it will meet the demand for specific designs to guide the planning of those
developing facilities for amphibians. Sufficient guidelines are given above
and in Chapter VI for the user of amphibians who handles only a few ani-
mals for short periods. Such a facility is not described here; the reader is
referred to Schmidt and Hudson (1969~. The descriptions presented are
not exhaustive (see also Boterenbrood, 1966; Frazer, 1966) and the reader
must evaluate them with respect to the animal- and facility-oriented desi-
derata listed in Section C.4. The serious planner should visit one or more
of these facilities before investing heavily in equipment for the care of
laboratory amphibians.
a. The Amphibian Facility of The University of Michigan
The housing and management system used in this facility is thoroughly
described in Nace (1968~; although improvements have changed some
specific operations, that document remains essentially current. It is a
flow-through system designed to house both anurans and urodeles that
number in the thousands. Major emphasis is on R. pipiens, but significant
colonies of X. Iaevis and B. orientalis are also under development. Small
test colonies of approximately 10 other anuran and urodele species are
also maintained. The colony is managed and data manipulated with the
assistance of computer-based techniques (Nace et al., 1973~.
Figure 14 illustrates the enclosure used to house larvae of all anuran
species from the initiation of feeding until the emergence of forelimbs.
Figure 15 shows a portion of the five-tiered rack that carries these enclo-
sures. One rack measuring 0.46 X 2.44 X 2.44 m (1.S X 8 X 8 ft) carries
130 enclosures. Each enclosure houses 50-75 larvae initially, which are
thinned to 15 per enclosure by the time of metamorphosis. Thus each
rack carries from 2,000 to 9,750 larvae, depending on their develop
OCR for page 65
65
mental stage, or 166-800 larvae per square foot of floor space occupied
by the rack. This configuration allows precise control of water flow,
easy cleaning by frequent flushing or by enclosure replacement, isolation
of small or large groups of larvae of known identification, ready access for
servicing, safety, efficiency of space utilization, modest installation costs,
long-term service, and low maintenance costs. The growth characteristics
of larvae in these enclosures closely resemble those of larvae in nature.
A rack of enclosures for juveniles and adults is shown in Figure 16.
The enclosures consist of transparent plastic mouse containers inserted
into similar opaque containers. Each measures 0.19 m (7.5 in.) deep,
0.24 m (9.5 in.) wide, and 0.45 m (18 in.) long, but in combination their
depth is 0.25 m (10 ink. Each combined enclosure may contain up to 30
or 40 juvenile R. pipiens. A rack measuring 1.13 X 2.44 X 2.44 m
(3.7 X 8 X 8 It) carries 76 of these enclosures for a capacity of approxi-
mately 30 juvenile frogs per square foot of floor space. Similar racks carry
containers that measure 0.20 m (8 in.) deep, 0.39 m (15.5 in.) wide, and
0.50 m (20 in.) long, but 0.35 m (14 in.) deep when combined. These are
used to house from 20 to 100 adults per combined enclosure. The smaller
number of occupants is used when maximum growth is desired; the larger
number when kidney tumors are being induced. A rack carries 36 such en-
closures for a floor density of 24-120 adult frogs per 0.09 sq m (1 sq It). An
illustration of a disassembled enclosure is shown as Figure 4 in Nace (1968~.
Each combined enclosure contains water in the opaque portion to a
depth appropriate to the behavior of the amphibian species it contains. An
opening in the floor of the transparent component allows frogs to move
between the aquatic environment and the terrestrial environment of the
transparent component. The latter contains a high, dry shelf, neoprene
mesh on all floor surfaces, and several shards of unglazed pottery. One of
several insect-proof lid designs is shown. Each enclosure is placed on slid-
ing-arm runners to facilitate access and maintenance of the heavy, water-
filled containers. For ready access, water control valves are placed at the
front immediately above each enclosure, but a tube guides the inflowing
water into the back of the enclosure. Figure 17 illustrates a drain tube-
either a trombone-slide device or a wire reinforced hose-which drains
each enclosure toward a trough running the length of the rack at each tier
and receiving drainage from enclosures on each side of the rack. Access to
the animals is possible either through the opening in the lid or by separa-
tion of the transparent from the opaque component of the enclosure. Food
is introduced in appropriate containers placed on the floor of the upper
component.
Xenopus and other aquatic amphibians may be held in these enclosures
by removing the transparent component and placing the lid on the opaque
OCR for page 66
66
..~9 ~
~ h ~
I Jo
..
4:g~
.~..~.~7 ,0
c
~ l,
-
l
B
OCR for page 67
67
component. Removal of wastes by flushing may be supplemented by the
use of water vacuum tubes.
Figure 18 diagrams a hibernation enclosure. These enclosures are placed
In a "4 °C" coldroom in which temperatures oscillate between 0 and 4 °C
(32-39 °F). The 20-gal capacity of the enclosure minimizes the tempera-
ture fluctuation experienced by the frogs and usually is stable between
2 and 3 °C (36-37 °F). Frogs received from the wild during the winter
have usually been exposed to "room temperature" for at least several days.
They are washed free of packing material and placed in the hibernation
enclosure in water at room temperature. The capacity of the enclosure
ensures that the water temperature does not drop to hibernation tempera-
ture faster than the anunals can adjust to it. The pump and filter device en-
sure clear, aerated water circulated continuously by a gentle flow that does
not agitate the hibernating frogs. Every 7-10 days, 2-3 gal of water are
drained from Me bottom to remove collected debns. This water is re-
placed with fresh, prechilled water.
Frogs remain at the bottom of the enclosure, particularly when the light
is on. When the light is off, they sometimes swan to the surface. As many
as 100 wild-caught R. pipiens females that retained usable eggs have been
held in hibernation in this type of enclosure from October to July.
FIGURE 14 Diagram of the larvae enclosure. A. Flow-through configuration: (a) a
10-mm plastic stand-pipe fixes water level. It is set in a rubber stopper readily re-
tained when firmly inserted into the neck of the enclosure; (b) a 15-mm-diameter
stiff, open plastic tube with legs assures mixing of incoming water; (c) water level;
(d) a barrier screen of stainless steel (2 mesh sizes for different stages) retains tad-
poles but passes debris; (e) plastic cuffs on the screen assure contact with (b) and
with the sides of the enclosure to prevent escape of tadpoles into the neck of the
enclosure; If) debris collected in the neck of the enclosure. The bottle should be
one with a steep slope in this region to assure that debris is cleanly removed by
flushing; (9) a food pellet; (h) the pattern of water flow is indicated by the arrows.
B. Flush configuration: A stopper is placed in the open tube to change the drain
system from a flow-through, water-mixing to a flush device. The outer tube is
pumped up and down several times to initiate siphon action. Debris is siphoned from
the neck of the enclosure, an action aided by twisting the bottle to loosen the debris.
On completion of siphoning, water may be poured in to return the enclosure to
overflow level or it may return to this level more slowly by drip addition. C. Assem-
bled and disassembled: Each enclosure, fabricated of round 1-gal plastic bottles with
the bottoms removed, contains approximately 3 liters of water, and water is drip-
ped into the enclosure at a rate of approximately 3 vol per day. Flushing removes
and replaces approximately 0.5 vol and is conducted once or twice a day depend.
ing upon the developmental stage of the larvae. Thus, larvae are in an effective vol
of approximately 0.2-1.0 liters per larvae per day depending upon their stage.
OCR for page 68
68
'-v
~1
.~
~ -~--__ ~
::
FIGURE 15 A portion of a rack of tad-
pole enclosures.
i
_~ - ~ .~
_ :. ~
·. ~ __
-'it
"'~3
~ :::: ~ -- .. ~ :;
~~__, sit
~ ' '"
-'91Iqi 31a
FIGURE 16 A portion of a rack of en-
closures for juvenile and adult frogs.
b. The R. catesbeiana Facility at Louisiana State University
This housing and management system is a flow-through system designed to
house R. catesbeiana for a program to test management and husbandry con-
cepts. The colony is comprised of laboratory-reared and wild-caught con-
ditioned animals.
Rearing enclosures, fabricated from 1.22 m (4 ft) diameter round or
oval galvanized cattle watering tanks coated with epoxy paint, can be used
to house up to 100 juvenile frogs with a 76.2-mm (3 in.) snout-vent length.
Coating is necessary to prevent possible toxification by zinc. The enclo-
sures are inclined at an angle so that three fourths of the tank floor is covered
with water with a depth of from 10 to 40 mm at the deepest point. A
drain line is placed in the side about 40 mm from the bottom. A plastic
tubular net is inserted in the drain to prevent food (fish) from escaping.
If shallower water is desired, a drain can be cut in the bottom and an
overflow tube (vertical) installed at the proper height. Such a drain also
aids in flushing the tank periodically.
As the frogs grow, higher sides are required to prevent escape. Thin
aluminum or plastic sheeting is attached to the sides to increase the height
of the walls to 1 m (3 ft). The extension does not completely encircle
the tank as it would be impossible to lean over the tank and work with
the frogs. For this purpose a 0.6-m (2 ft) section is left open on one side.
Small enclosures used to house experimental groups of frogs are illu
OCR for page 69
69
h
A. Expanded
- _~
~ ,
B. Assembled
FIGURE 17 Drainage device used in juvenile and adult enclosures. (a) Nylon drain
20 mm in diameter; (b) set nuts and gaskets holding the drain in the floor; (c) of the
enclosure; (d) drain hose held in place by hose clamp (e); (f) variable length, stiff
garden hose which snuggly fits the offset in the nylon drain and whose length deter-
mines the depth of the water; (9) a plastic mesh sleeve which snuggly fits the inside
of the hose (f) and into the narrow portion of the drain. When the hose (f) is lifted
to rapidly drain the enclosure, this sleeve forms a barrier preventing juvenile frogs,
carried in the water flow, from passing through the drain or, if large enough, from
clogging its opening; (h) a plastic mesh sleeve formed into a cap by heat sealing. It
is larger in diameter than the hose (f) and long enough not to be dislodged. It pre-
vents juveniles from escaping through the hose. This cap need not be used in en-
closures for frogs too large to pass through or become lodged in the hose; (i) water
level. When only a film of water is desired, hose (f) is removed and cap (h) is used in
combination with a longer inside sleeve (9). The depth of the water is then no
greater than the thickness of the set nut (b).
OCR for page 70
70
A. . h
\~-~-~ ~
W1~1
A
FIGURE 18 Hibernation enclosure. {a) a 20~a1 plastic "garbage" container; {b) a
ring of cement blocks raises the enclosure and permits the weight of the water to
form the enclosure bottom into a shallow funnel shape; {c) nylon drain, hose, and
clamp; (d) a heavy plastic screen weighted down by a sealed ring of plastic tubing
filled with "shot" keeps the frogs from occluding the drain and forms a reservoir
for debris; (a) a 15-mm-diameter stiff plastic tube extends to a point just above the
screen; (f) a 10-mm plastic tube from a compressed air line bubbles air into the
larger tube (a). The combination of (e) and (f) forms an air-lift pump which aerates
the water while lifting it to the surface; (a) a plastic container filled with fiberglass
serves as a filter. The bottom of this filter is perforatecl. It is attached to the side of
the enclosure above the water-line to hold the filter and pump in place. Tube (e)
passes through the filter and the aerated water lifted through it is spilled over the
fiberglass and is filtered as it returns to the enclosure through the bottom of the
filter; (h) the plastic lid of the enclosure contains a suitable screen to admit light
whose duration is regulated by a timing device.
strafed in Figure 19. These enclosures house up to 25 frogs with a 76.2-mm
(3 in.) snout-vent length or 50 metamorphosing larvae or juveniles. Two
types are used. One measures 0.28 X 0.33 X 0.14 m (11 X 13 X 5.5 in.),
the other 0.45 X 0.73 X 0.20 m (18 X 29 X 8 in.~. By tilting these con-
tainers, the elevated section provides a terrestrial environment. These en-
closures require covers to prevent the escape of frogs but need not be
insectproof as only fish are used in the diet of these animals. It is impor-
tant, however, that the cover material not cause injury to the frogs. Plas-
tic netting is better than metal screen or hardware cloth; metal screening
may be used for strength if it is faced with neoprene mat.
If a shelter is provided in these enclosures, the frogs will move under it,
making escape less likely during feeding or cleaning operations. Restaurant
OCR for page 71
71
supply houses have a wide selection of plastic and fiberglass containers,
some of which have sliding tops, that can serve in place of the enclosures
shown in Figure 19.
The use of a ribbed, black, rubber floor padding in the dry portion of
the floor of enclosures eliminates cuts and skin abrasions and facilitates
frog movement without slipping. No flooring material has been found
that is easy to clean and maintain, but use of some type of padding is ad-
visable. Plastic or rubber netting is unsatisfactory as shed skin and dead
fish get caught, making cleaning difficult.
c. Southern Frog Company (J. M. Priddy, Dumas, Arkansas)
The husbandry facility for R. catesbeiana at the Southern Frog Company
uses a flow-through system that is a scale-up of the system used at the
R. catesbeiana facility at Louisiana State University and follows designs
originally developed by Stearns (1973~. It is designed to produce com-
mercially significant numbers of wild-caught conditioned and laboratory-
reared animals (Priddy and Culley, 1971~.
In place of cattle watering tanks, circular enclosures 6.1 m (20 It) in
diameter with 0.75-0.90-m (30-36 in.) walls of concrete block are used.
~ .~
~ - ~
it-- t ~
A
FIGURE 19 Small enclosures for test animals.
it--_
OCR for page 72
72
The flooring is concrete and slopes to one side with a drain line. Since
lime leaches from concrete and causes skin erosion, a concrete sealer is
painted on all inside surfaces and then covered with lead-free or other
heavy-metal-free epoxy paint or swimming pool paint. These enclosures
house up to 5,000 newly metamorphosed frogs or 1,500 with a snout-
vent length of 76.2-127 mm (3-5 in.) for a floor density of 2-16 frogs
per 0.09 sq m (1 sq ft). When animals in such numbers are placed in a single
enclosure, sets of two concrete blocks placed in V patterns at several loca-
tions in the container are useful to minimize animal pileups that may oc-
cur when the animals are alarmed.
R. catesbe~ana larvae are reared through metamorphosis in 4 months
from fertilization at an average temperature of 28 °C (82 F) using these
same enclosures. One or more frames measuring 0.31 X 0.31 X 0.92 m
(1 X 1 X 3 ft) are covered with nylon mesh and placed on 0.15-m (6 in.)
legs in these 6.1-m (20 ft) diameter enclosures that are flooded to 0.31 m
(12 in.) with continuously flowing water. The 200 larvae placed in each
frame are fed a finely powdered minnow meal that floats. To prevent this
feed from washing out of the nylon enclosures, it is placed inside floating
frames made from Styrofoam coolers with their bottoms removed. The tad-
pole swim under these frames and to the surface to ingest the feed. Food
is given twice a day at the same time each day to condition the tadpoles to
maximum feeding. R. pipiens larvae as well as other larvae may be reared
by this technique.
d. The Aquatic Animal Facility of Arizona State University
The stainless steel housing system for anurans and urodeles of the Aquatic
Animal Facility of Arizona State University utilizes recirculating water
filtered through dacron and passed under germicidal lamps. This design
requires a minimum of maintenance, provides control of temperature and
light, and has been successfully tested as a holding facility for axolotls,
frogs, toads, fish, crayfish, and turtles (Justus and Cullum, 1971; Cullum
and Justus, 1973~.
The volume of water in this system is 500 gal recirculated at the rate
of 850 gal/in and exchanged by the removal of 11 gal/in and the addition
of domestic water at 11 gal/in. Bacterial plate counts range from 600 in
water draining from adult frog holding tanks to 15 in water after ultra-
violet lamp treatment.
The design of the water circulation and processing system may be
found in Cullum and Justus (1973~. The following expands on their de-
scription of certain of the animal enclosures.
Figure 20 shows a breeding unit that measures 0.35 X 0.20 X 1.20 m
OCR for page 73
73
(14 X 8 X 48 in.). Newly fertilized eggs are placed in 0.23 m (9 in.) square
stainless steel frames with fine muslin bottoms. These are set in the breed-
ing tank until the tadpoles have reached swimming stages. They are then
released into the tank that can hold 6,0Q0 tadpoles up to 20 mm (0.78 in.)
in length for a density of 1,~50 young tadpoles per 0.09 sq m (1 sq It).
Members of different clutches are not separated and only one age group
is carried per cycle.
Tadpoles are then moved to a rearing tank until metamorphosis is com-
pleted. This tank (Figure 21) measures 1.5 X 3.1 m (5 X 10 It) and is
0.20 m (8 in.) deep. It can hold 8,000 tadpoles for a density of 160 larvae
per 0.09 sq m (1 sq ft).
Upon metamorphosis, the frogs are moved to adult holding tanks. These
measure 0.40 X 6.1 (1.3 X 20 ft) X 0.20 m (8 in.) and contain an aquatic
area and feeding trays (Figure 22) that may be moved to allow the use of
dividers to separate frogs of different categories. On the trays the frogs are
fed mealworms and other crawling insects or crickets with their hind legs
removed. The capacity of each adult holding tank is 80 frogs. Since these
tanks are stacked in two tiers, the capacity is six frogs per 0.09 sq m ( 1 sq
ft) of floor space.
The holding tank may be adapted to the housing of A. mexicanum.
This system is also useful for other aquatic amphibians such as Xenopus.
Perforated stainless steel baskets inserted in these tanks serve as enclosures
for the aquatic animals.
_. . ~
: ~
~1
'I-.....
rat
~.~;
~ ~ ~ . ~
· ~
_~
_~]
~:::.s~ id
~ 1~'~:~:~
Hi: ::~
~_....: .;
FIGURE 20 A breeding unit for eggs
and young larvae. Water is received in
the upper compartment, passes to the
larvae enclosure below, and is returned
to the recirculating system by the com-
ponents on the bottom level. Note the
positioning of lights.
OCR for page 74
74
1
:
Gil
.-...
:~ -A __
__-=-=~!
_ - ~ : ~ .
Fit _
FIGURE 21 Interior of a rearing unit. The top is removed from the drain com-
ponent. Water enters through the filter at the bottom of this component and exits
via the stand-pipe. Terrestrial areas for metamorphosing frogs are shown adjacent
to the drain component, the top of which also serves as a terrestrial area.
~ 1
"v .
~IF; i.
~-':~
--.
:-.~ ~
1., ~:~ l
_ f-~ ~ j a.
_::. Hi ; ~ ~ ~
~:~ f : -
~a_ ~ ~ ~ a .. -a
~_: ! ~ :,' ~.~"~'~ "A
I_ ~ ''' In'
`_ `
~ ~ ~X'~':~
~" ~ ~:47; ~^ ,,
FIGURE 22 Interior of an adult holding tank. A feeding tray with access ramps
extending both upstream and downstream is shown.
Representative terms from entire chapter:
water quality
