Welcome to the course Working With Amphibians in Research Settings.

The class Amphibia is a highly diverse class of 'cold-blooded' (ectothermic) vertebrates comprising over 4000 species, that have been used widely in teaching and research. The frog was one of the first vertebrate organisms most people have used to learn anatomy in secondary school. Frogs and other amphibians have also been important models for embryology, physiology, studies of tissue regeneration, and as sentinels for environmental toxicology.

Salamanders are discussed in mythology as creatures that are born of fire, are impervious to fire, and are able to extinguish fire by virtue of their coldness. They were widely believed to poison the fruit of trees upon which they climbed. The image shows a crowned salamander in a bed of fire, the heraldic insignia for King Francis I of France.

The word amphibian derives from the two greek words amphi meaning 'both' and bios meaning 'life'. Amphibians are unique in that in their larval forms they are totally aquatic while, as adults, most are primarily terrestrial (though some species are aquatic throughout their life cycle).

This is the amphibian module in a course series on the use of animals in research. Each course in this series refers to a different animal class, order or species.

• Lessons that are similar in all the species courses relate mainly to the regulatory coverage, housing requirements, and zoonotic hazards of each animal species.

• The remaining lessons present information that is more specific to each animal species, such as biological features, anesthetic doses, and biomethodologies.

The goal of this course is to cover important information about using amphibians in biomedical research settings. If you are responsible for handling amphibians or if you must write an animal use protocol, this course will be useful by providing you with:

• Information on key regulatory issues.
• Guidance on searches for alternatives in the care and use of animals.
• Highlights of unique biological features of these animals.
• Overviews of acceptable basic methodologies.
• Requirements for supportive care procedures.

Hypertext links in this course provide you with supporting information, such as regulatory sources, drug doses, practical tips, etc.

This course will not provide you with detailed information on how to conduct the methods and procedures described. For this, you should use other courses offering in-depth information and hands-on instruction from your institution's animal facility staff.

Click here to view the credits for this course.

The class Amphibia is a diverse class of vertebrates that comprises three orders and more than 4000 species. The orders are: Gymnophiona, Anura and Caudata.

Gymnophiona

These are limbless, burrowing amphibians with poorly-developed eyes and annular grooves on the body. They resemble earthworms and occur in in tropical areas of Africa, the Americas and Asia. Also called caecilians, some species are oviparous and some are viviparous. Most species have aquatic juvenile forms but some larvae are terrestrial. Members of this order are rarely used in the laboratory.

Anura

Anurans are the frogs and toads and at more than 3400 species they account for the bulk of amphibian species. These tailless amphibians usually have long hindlimbs that are adapted for jumping. The eggs are usually laid in water, although some are carried by the adult and the larvae are aquatic. Adults may be terrestrial or aquatic. Commonly used laboratory genera include:

• Bufo (the true toads) which have thick, warty skin and well-developed parotid glands.
• Rana (bullfrogs and leopard frogs) are semi-aquatic with smooth skin and are best known for their use in teaching anatomy and studies of physiology.
• Dendrobates (poison dart or arrow frogs) which are brightly colored, arboreal frogs that produce cutaneous toxins which are used by aboriginal peoples and studied by scientists.
• Xenopus (African clawed frogs) which are primitive, aquatic frogs used as a sourse of oocytes for developmental and genetic studies.

Caudata

The order Caudata (formerly classified as Urodela and still sometimes referred to as urodeles) includes three general groups: sirens, 'primitive' salamanders and 'advanced' salamanders. Sirens are completely aquatic, lack hindlimbs and have external gills. Salamanders are long-tailed amphibians that usually have two pairs of limbs of approximately equal size some are totally aquatic and some are terrestria as adults. Some species (such as the axolotl) exhibit 'neoteny' or the persistence of larval characteristics into adulthood.

Fertilization is internal except in sirens and Cryptobranchiae(a group of primitive salamanders). Eggs are usually deposited singly or in clumps or strings in water and hatch into aquatic larvae, though some species salamanders have direct development of terrestrial eggs.

Important Laboratory Genera:
• Ambystoma spp. - These are heavy bodied aquatic or burrowing animals and include the tiger salamanders and axolotls.
• Necturus spp. - Mudpuppies. These are obligate aquatic animals and are frequently used in developmental biology studies.

Not surprisingly, amphibians possess myriad anatomical and physiological characteristics that differ greatly from mammals. However, there is a biological diversity within the class that is remarkable as well. Some of the more important or interesting characteristics follow:

Reproduction and Development

Reproductive stragegies are remarkably diverse among amphibians. Fertilization of eggs is commonly external, but may be internal in many salamander species. In these, the male deposits a "spermatophore" which the female collects into her cloaca. Fertilized eggs may develop free in the environment, or they may be nurtured by either the male or female depending on species. In some anuran species the eggs are carried and develop on the female's back. In others, they are nurtured in the buccal sacs of the male or even in the stomach of the female.

The eggs develop through a process known as "metamorphosis" into larval forms (most of which are aquatic) which themselves progressively transform into either aquatic or terrestrial adult forms. Salamanders tend to exhibit a characteristic called "neoteny" which refers to the retention of larval traits such as tails and gills by the adult.

Respiratory System

Respiratory strategies vary widely among amphibians. Larval forms breathe via gills while adult forms may have gills or primitive lungs. Among gilled adult forms, the gills may be external (as in axolotls) or internal. Some salamanders have neither lungs nor gills and respire transcutaneously. Even when other respiratory structures are present, the skin is the principal respiratory organ for most amphibians. Therefore, it is important that they be able to keep the skin moist.

Blood and Cardiovascular Systems

Amphibians have nucleated red blood cells and nucleated thrombocytes.

Larval amphibians have a two-chambered heart (atrium and ventricle) while most adults have a three-chambered heart (right and left atria and a single ventricle).

Blood plasma has a lower osmolarity in amphibians (approximately 200mOsm/kg as compared to 250-300 mOsm/kg in mammals.) This needs to be considered when providing replacement fluids for dehydration.

Amphibians possess lymphatic sinuses throughout the body and especially in the subcutaneous tissues. In anurans, the sacral lymph sacs are sometimes used for "intravascular" injections. Lymph drains into the venous system, assisted at lymphatic/venous junctions by small contractile structures.

Gastrointestinal System

Adult amphibians are carnivorous. Larval forms may be carnivorous or herbivorous depending on species. Most adult anurans have an extensible tongue used for capturing prey whereas Xenopus spp. and salamanders prehend food directly with the mouth.

Frogs often vomit as a defensive strategy and some may actually evert the stomach.

Melanin is a normal pigment in the liver of many amphibians.

Nervous System

Amphibians have a primitive brain compared with mammals and have ten paired cranial nerves instead of twelve. The bulk of the cerebral cortex consists of olfactory lobes.

Larval forms possess a "lateral line" system of cutaneous sensory cells that are sensitive to low frequency stimulation and differences in water pressure. The lateral line is preserved in many species in which the adults remain aquatic.

Anurans often have an exposed tympanic membrane for sensing high frequency sounds. Lower frequency sounds are perceived through the bone structure of the skull. The tympanic membrane may exhibit sexual dimorphism, with the male structure approximately twice as large as the female's.

Integumentary System

Amphibian skin is moist and serves multiple functions besides the barrier function of mammalian skin. Amphibian skin is highly permeable and is a principal respiratory organ for most species. In addition to gas exchange, it is permeable to many other molecules and it is through the skin that anesthetics are frequently administered. Because of its permeability, care must be taken to avoid cutaneous exposure to potentially toxic chemicals.

There are many glandular structures in amphibian skin that produce the mucus layer that protects amphibians from water loss, abrasions and infections. Many anurans and caudates also produce toxic skin secretions that protect the animal from predators. Toxins vary among species but may include neurotoxins, cardiotoxins, hallucinogens, emetics and paralytics.

Amphibians periodically shed their skins as do reptiles.

Excretory System

Amphibians, like birds and reptiles, possess a common collecting chamber called the cloaca. Products of the urinary tract, the intestinal tract and the reproductive tract empty into the cloaca before finally exiting through the anus.

Amphibians have a mesonephric, rather than the metanephric kidney seen in mammals, which is unable to concentrate urine beyond the normal plasma osmolarity. Urine is collected in a urinary bladder before excretion. The bladder of many amphibians empties when the animal is stressed. This may serve to deter predators.

As ectothermic (cold-blooded) animals, amphibians are not subject to regulations promulgated by the United States Department of Agriculture (USDA) under the Animal Welfare Act. However, they are covered by Public Health Service (PHS) policy and they are subject to the general policies described in the Guide for the Care and Use of Laboratory Animals (Guide) published by the National Research Council (NRC).

The Guide considers amphibians to be "non-traditional" laboratory species and as such does not offer specific recommendations for housing and handling. It does, however, state that "expert advice" should be sought in that regard when use of particular non-traditional species in laboratory or field studies is anticipated.

Several sources offering more detailed recommendations for housing, handling and use of amphibians are available. These include:

• Guidelines for Use of Live Amphibians and Reptiles in Field and Laboratory Research from the American Society of Icthyologists and Herpetologists.
• Amphibians:Guidelines for Breeding, Care and Management of Laboratory Animals, 1996 from the National Academy Press, National Research Council
• CCAC Guide, volume 2, 1984 from the Canadian Council on Animal Care.

The amphibians most commonly used in laboratory research are available from various captive breeding programs but many species of amphibians are considered to be threatened or endangered in their natural habitats, and their use is regulated in order to protect these populations. Investigators should be aware of the regulatory status of species they intend to use in their studies

The United States Department of the Interior (USDI) delegates responsibility for oversight of threatened or endangered amphibians in international and interstate commerce to its Fish and Wildlife Service (FWS). The FWS exercises its authority through the CITES agreement (Convention on International Trade in Endangered Species of Flora and Fauna) for international commerce and the Endangered Species Act (ESA) for interstate commerce in NHP.

These instruments categorize different species according to the degree to which wild populations are at risk of extinction and they establish rules for trading in such species or prohibit trade altogether. These regulations may be reviewed in Title 50 of the Code of Federal Regulations, part 17 for ESA regulations and part 23 for CITES regulations.

Finally, virtually all 50 states have their own laws regarding taking of amphibians from the wild. Investigators should become familiar with state wildlife laws if they intend to collect amphibians from the wild or conduct field studies involving amphibians.

The Public Health Service Policy requires institutions to have an occupational health and safety program for individuals working with laboratory animals. This requirement is also reiterated in the Guide.

It is the responsibility of principal investigators to assure that their laboratory staff are informed of and participate in their institution's occupational health and safety program.

Elements of an occupational health and safety program, including institutional responsibilities, are described in the guideline, Occupational Health and Safety in the Care and Use of Research Animals (shown at right), published by the National Research Council.

Although the risks are generally small, amphibians may pose certain hazards to individuals who work with them. Some of the more significant hazards are listed below:

Allergies

Allergy to amphibians is uncommon but has been reported. Individuals with known allergies or with asthma might be at increased risk of developing allergy to amphibians and should use protective clothing when working with these animals and they should make the health professional in charge of the institution's occupational health program aware of their history of allergy.

Physical Injury

Amphibians do not pose the same degree of risk for bites or scratches as certain mammals such as dogs, cats or monkeys, but larger species of anurans and caudates are capable of causing painful bites and care should be exercised when handling them.

Perhaps a more serious potential for physical injury relates to the generally wet environment in which they need to be housed. In such circumstances, there is a shock hazard when electrical outlets, switches and appliances are not properly constructed. It is particularly important that electrical fixtures be moisture resistant and equipped with ground fault interruption devices.

Toxins

Amphibians are somewhat unique in that there are so many species that are capable of secreting toxic compounds from dermal glands. The parotid glands of toads in genus Bufo and some salamanders and other dermal glands of anurans such as poison arrow frogs are moderately to highly toxic. The image shows a specimen of poison arrow frog.

Staff working with amphibians should be aware of the potential for toxicity of the species with which they are working and should wear powderless gloves and or eye protection when appropriate.

Infectious Disease

There is a short list of infectious agents of which amphibian users should be aware:

• Salmonella spp. may be carried by amphibians, particularly if wild-caught. These are capable of causing enteric or systemic disease in humans ranging from asymptomatic to potentially fatal.
• Several species of Mycobacterium including M. marinum, M. xenopi and M. fortuitum are present in wet environments and may be capable of causing skin and lymphatic infections in humans.
• Chlamydia psittaci, the agent causing human psittacosis, has been isolated from some amphibians. Although no human cases have ever been definitively associated with amphibian exposure, personnel working with amphibians should be aware of the risk and protect themselves by frequent handwashing and use of personal protective equipment where appropriate.

Federal Animal Welfare Act (AWA) regulations require investigators to provide a written assurance that alternatives are not available for proposed procedures in animals that are likely to cause pain and/or distress. Although amphibians are not covered by the AWA, Public Health Service (PHS) policy expects investigators to justify their choice of animal model. As a consequence, most Institutional Animal Care and Use Committees (IACUCs) will expect investigators to consider alternatives to using live amphibians in their studies. If alternatives are available but are not satisfactory for the proposed research, the investigator is required to explain why the proposed procedures must be used instead of the less distressful alternatives.

Your protocol form, therefore will ask you for an assurance that you have considered alternatives to the use of animals if painful or distressing procedures are proposed in order to satisfy the mandate by the PHS Policy to avoid or minimize discomfort, pain, and distress consistent with sound scientific practices.

Alternative procedures are those which may replace animals with nonanimal methods, reduce the number of animals used, or refine the methodology to minimize animal pain or distress.

For more information on what is meant by alternatives to the use of animals, please refer to the course Working with the IACUC, which is part of this series.

The assurance must take the form of a written narrative that describes which sources were used to determine that alternatives were not available.

Although other methods may be acceptable (for instance, expert knowledge in a very specialized field) the USDA considers a computer database search to provide the best information, and typically, you may be asked to provide the results of a database search.

If a database search is used, the following elements of the search must be described:

1. The databases that were searched.
2. The date the search was performed.
3. The years of citations covered by database searches.
4. The key words and/or search strategy used when searching a database.

The IACUC is responsible for determining whether or not a thorough search for reasonable alternatives was conducted and the USDA looks at the above elements when it reviews the IACUC's performance. It is strongly recommended that this information be sought during development of a protocol.

Organizations that can assist you in performing an alternatives search are:

• ALTWEB, Center for Alternatives to Animal Testing, Johns Hopkins University
• Animal Welfare Information Center, National Agricultural Library
• Center for Animal Alternatives, University of California – Davis

The following is a case study of alternatives searches that may guide you in the development of a search strategy that is pertinent to your own research.

Click on a link below for a sample search on key terms for this example.

The links will take you to the named database where you can conduct a search in real time.

Example Search:
Models for tissue regeneration

PubMed

BIOSIS

It is a part of a recurring mantra, given the diversity in Class Amphibia, that ideal housing conditions vary widely depending upon the species being housed. Some general principles do apply though.

• First, amphibians are all ectothermic. The requirement for appropriate environmental temperatures is universal. As a rule, amphibians are more sensitive to heat stress than to cold stress.
• Next, it is important to understand the biology of the particular species being housed and to consult with experienced users on details of housing requirements for that species.
• Finally, it is important that caregivers are familiar with signs of illness or distress - particularly those that might be environmentally related.

Primary Enclosures

Cages or tanks housing laboratory amphibians may be may be simple or complex arrangements. The following general considerations should be observed:

• Enclosures should prevent the animals from jumping out (in the case of frogs) or climbing out (some salamanders). Often this is accomplished by means of a weighted cover. Metal lids such as screens or grids may be sources of abrasive dermal injury so plastic or glass covers may be preferable.
• Enclosures should be made of materials that are easily sanitizable. The presence of a drain in the bottom may be convenient. Care should be taken to avoid materials that are dyed or contain chemicals that could leach into the water and pose a risk of intoxication to the animals.
• Some amphibian species may be group-housed while others are intensely territorial. Obviously, the latter species should be housed separately. When group housing is used, care should be taken to avoid overcrowding. With Xenopus species, volume recommendations commonly range between 3-4 gallons per frog.

Water Quality

Water is essential for preservation of life and health for all living things, but it is particularly critical for amphibians. It is a primary requirement for aquatic larvae and adults but it is critical for terrestrial species as well. Highly permeable amphibian skin puts the animals at increased risk of dehydration in the absence of adequate moisture, and exposes them to water potentially toxic components that are benign in mammals. Chlorine in municipal water supplies is likely the most common problem in this regard, but minerals, residues from cleaning chemicals, and ammonia or nitrites from decomposing feed and excreta may also be problems.

As noted previously, amphibians do better in relatively cool temperatures, generally 10-20 degrees C, though the ideal temperatures will vary by species. Water generally does not need to be aerated, with the possible exception of water housing larval forms or gill-breathing adults. The pH of water (and other environmental elements for terrestrial species) is best kept in the neutral or slightly alkaline range (7-8.5). A very acidic environment (less than 4.0) is likely to be toxic. Water in static systems should be changed after feeding.

Light, Temperature and Ventilation

Lighting

Most amphibians are nocturnally active and ideal light cycles will depend on species and intended use. For breeding or behavioral studies, light cycles may need to be matched to natural cycles. For most biomedical uses, a 12-12 light/dark cycle is satisfactory. Light sources that provide a full wavelength spectrum are usually recommended, expecially where natural light is not available.

Temperature

Temperatures are most important at the level of the primary enclosure and tanks or cages usually should be kept cooler rather than warmer. Room temperatures may dictate primary enclosure temperatures, especially when bulk-type tanks are used. For terrestrial species, it is desirable for a temperature gradient to be maintained so that the animals are able to modulate their body temperatures by changing their location within the cage. A single, incandescent bulb at one end of the primary enclosure and a sheltered area at the other are simple ways to achieve this. Water should be available for terrestrial animals to immerse themselves. Room temperatures within a range of 20-25 degrees C will usually be adequate.

Ventilation

Air flow rates recommended for mammals (15-20 changes per hour) are likely to be dehydrating in terrestrial amphibians. Ambient humidity for these animals should be maintained at 70-80% to help animals maintain skin moisture. Ventilation rates should be decreased from mammalian standards and/or air should be humidified for amphibians.

Sanitation

Sanitation of amphibian enclosures can pose special challenges because of territorial behavior and pheromone marking, and because of the sensitivity of amphibians to chemical agents.

Frequent sanitation of terrestrial species can disrupt territorial markings and lead to behavioral distress. On the other hand, ignoring sanitation can lead to accumulation of toxic metabolites and opportunistic bacteria. A balance must be found between the competing goals of a clean environment and minimizing disruption of the animal's normal behaviors. Singly housed amphibians in terraria may be managed effectively by cleaning cages approximately every two weeks. Amphibians in an aquatic environment require cleaning more frequently, at intervals ranging from daily to weekly. Xenopus species are commonly cleaned after feeding, approximately three times weekly.

Sanitization should be accomplished without using the chemicals normally employed with mammals (e.g. detergents and quaternary ammonium or phenolic compounds). If the cages will tolerate it, standard cagewashers without chemicals are effective. If a disinfectant must be used, dilute bleach (e.g. 1:32 ratio with water) may be used if it is thoroughly rinsed after use.

Nutrition

Adult amphibians are carnivorous and most require moving (live) food, although some of the more commonly used laboratory species such as Xenopus and Ambystoma do well on chopped heart muscle, liver, or commercial fish diets (such as Purina's Trout Chow) or frog diets (such as Nasco's Frog Brittle).

Live foods may include crickets, mealworms, tubifex worms, fly larvae, earthworms, and "pinkies" (immature rodents). Crickets and other arthropods are usually deficient in calcium and vitamins. This may be supplemented by dusting the insects with a calcium/vitamin mix, or by raising the feed insects in-house and feeding them with diets enhanced with calcium and vitamins.

Larval amphibians are mainly herbivorous or omnivorous, becoming carnivorous as they undergo metamorphosis. They feed on copepods and other microscopic invertebrates, or flake-type fish food, alfalfa pellets or boiled spinach.

Wild populations of amphibians are on the decline and except for a few species, captive-bred populations are not readily available. In order to avoid pressure on wild populations, serious consideration should be given to whether wild-caught species are the best or only available model for a given study. If animals must be collected from the wild or if vendors who capture their animals from the wild are used, it is important to comply with all applicable federal laws (the Endangered Species Act), international agreements (the Convention on International Trade in Endangered Species of Flora and Fauna - the CITES Agreement) and state fish and game laws.

A few sources of captive-bred specimens are available for some species such as Xenopus and Ambystoma. Your institutional veterinarian can assist you in locating sources of good quality animals.

Acclimation, quarantine and conditioning are procedures that serve related but distinct functions.

A period of acclimation allows a newly aquired animal to become accustomed to its new laboratory environment.

A quarantine period is an interval in which newly acquired animals are housed and cared for separately from existing colony animals. Its purpose it to attempt to identify and correct any infectious diseases that new animals might introduce into stable, existing colony animals.

A conditioning period is one in which an animal is permitted to recover from the stress of transport and for animals of poor health and nutritional status to become stronger and more robust before they are used in research studies.

Although they accomplish different goals, periods of acclimation, quarantine and conditioning may be completed simultaneously. For example, a new animal may be acclimated to its laboratory environment while it undergoes quarantine to protect existing colony animals. An acclimation or quarantine period may also serve as an interval during which an animal's physical condition or nutritional state is improved - i.e. a conditioning period.

There is no well-documented basis for establishing consistent time periods for acclimation, quarantine or conditioning of new amphibians. The appropriate intervals will need to be established based on source, species and assessed risk to existing colony animals.

For example, captive-bred animals obtained from high quality commercial sources may require only short acclimation periods of a few days to adapt to new enclosures and diets, whereas wild-caught specimens may take much longer to adapt to captive conditions.

Similarly animals obtained from the same commercial or laboratory sources may pose little risk to an existing colony and quarantine periods may be short or absent. However, all wild-caught specimens should be quarantined and tested for parasites, cultured for infectious agents, and observed for clinical illness before mixing with existing colonies.

Finally animals obtained from good quality sources are likely to arrive well-nourished and healthy, requiring little or no additional conditioning. Animals from the wild or from sources of questionable quality may be in poor health or badly nourished and require extensive conditioning before they are used in research studies.

Collection

As noted previously, captive-bred animals should be used if at all possible. However, if wild animals must be collected, live-capture techniques should be used that prevent or minimize damage to the animal.

Many amphibians are slow-moving and can be collected by hand. Care must be taken to avoid removal of the protective mucus layer covering the skin of amphibians. If nets that are used, they should be made of soft cloth materials. Animals should be handled gently and transported in containers that protect them from trauma and dessication. Small amphibians can be temporarily restrained in plastic bags containing a small amount of water, filled with air and sealed. The soft sides cushion the effects of bumping or jumps.

Traps may be useful for collecting some species. If traps are used, they should be designed to protect the animals from injury or access by predators. They should be checked at least daily or more often when environmental conditions might threaten the well-being of trapped animals.

Handling

Experts vary on recommendations for handling amphibians with bare hands. Some biologists argue that delicate species are placed at greater risk when gloves are used because of a loss of tactile sensitivity by the operator. If amphibians are handled using bare hands it is extremely important for investigators to ensure that they have not applied insect repellents, perfumes, lotions, or other potentially toxic substances that might be absorbed through highly permeable amphibian skin.

Others argue that wearing disposable gloves when handling amphibians will protect the animals’ skin from abrasion, chemicals and the spread of infection. However, gloves containing talc should not be worn as they could irritate the amphibian’s skin. Gloves should either be ‘talc-free’ or rinsed in warm water prior to use. When handling highly toxic amphibians or where bites from animals are possible, gloves should be worn to protect the handler and contact with bare skin or mucus membranes should be avoided.

Medium and large frogs and toads may be grasped cranial to the hind limbs with the hind limbs fully extended. This helps prevent them from kicking. Larger animals may require a second grip around the forelegs. Larval forms and caudate species with gills should never be restrained around the neck because the gills will be damaged. Similarly, tails should not be used for restraint because they may easily detach. Some amphibians (such as small plethodontids which overheat easily in the hand) do not tolerate physical restraint well.

Procedures

Blood Collection

Blood may be taken from anurans from the femoral vein, the ventral abdominal vein or the lingual vein. In salamanders, the ventral tail vein is generally used for blood collection.

Blood may also be collected by cardiac puncture however, there is a high mortality associated with this technique and it should be avoided if other approaches are feasible. Cardiac puncture should be done using appropriate anesthesia.

Injections

Injection routes are generally similar to those of other laboratory species and include intramuscular, subcutaneous and intravascular approaches. Amphibians have no diaphragm separating the pleural and peritoneal cavities and instead have a single coelomic cavity. Intracoelomic injections are given at the midline in the caudal aspect of the abdomen.

Anurans have accessible subcutaneous lymph sacs over the dorsal sacral region. Since the lymph drains directly into the venous system, injections into the dorsal lymph sacs are comparable to intravenous injections. The lymph sacs can be located by observing their pulsations under the skin in the sacral region.The image shows a frog being restrained for an injection into a dorsal lymph sac

Oral Administration

Liquid agents may be administered by gavage using ball-tipped needles as for rodents or soft catheters.

Identification

Techniques used for identification of individual animals vary widely and depend to some extent on whether the animals are used in the laboratory or in field studies. The following techniques may be considered:

• Pigmentation patterns may be documented by means of photographs or drawings. This approach may avoid using more invasive procedures.
• Tattooing has been used with success in amphibians but it is time consuming, involves potentially toxic dyes, and tattoos may fade due to diffusion of the dye.
• The injection of colored inert plastics into the toe webs.
• Visible Implant Elastomer (VIE), a technique created for fish, may be used to mark amphibians. Problems associated with this technique include: migration of the mark when injected into the thigh, the lack of fluorescence of the mark due to the dark pigments of most amphibians, and the need to keep VIE cold until injected.
• Toe clipping should be avoided if at all possible. Toe clipping is painful and requires anesthesia, it may predispose the animal to infection and in many amphibian species toes may regenerate.
• In general, external tags are not recommended for amphibians because tags or wires could become snagged on environmental objects and improper insertion could cause tissue necrosis oor infection.
• Microchips (also called transponders) have been implanted subcutaneously or intracoelomically to mark amphibians. Migration of these transponders may make them more difficult to read. The image shows a close-up of a commercially available transponder.
• The attachment of small radio transmitters to amphibians may be used to identify and monitor the activity of individuals in field studies. Transmitter attachments that will impair reproduction, locomotion, behavioral interactions, thermoregulation, skin shedding or other normal activities, or that risk entanglement with vegetation or other obstructions should be avoided. Wherever possible, transmitters should be removed upon completion of a study or the transmitter attachments should be designed so that they are ultimately self-detaching.

There is little doubt that amphibians are capable of nociception - that is, response to potentially injurious stimulation. Indeed, an accepted model for assessing the analgesic activity of certain drugs involves dripping glacial acetic acid on the skin of frogs, in incrementally increasing concentrations, until the animal responds by rubbing at the affected area.

However, the nature of an amphibian's perception of pain is still controversial. Because they lack a limbic cortex and their cerebral cortex consists mostly of the olfactory lobes, it has been argued that amphibians cannot perceive pain in the same way as do mammals. Moreover, behavioral indicators of pain beyond simple avoidance responses are not well described for amphibians.

In light of the ambiguity of available information, humane practice dictates that in the absence of clear evidence to the contrary, it should be assumed that procedures that are painful to mammals will also cause pain in amphibians and measures should be taken to avoid or minimize that pain.

Terminology used for drugs that affect consciousness and pain perception is not consistent in the literature and is complicated by the fact that some drugs have effects that overlap categories. A further complication is that "consciousness" in amphibians is not well-defined and may be different than in mammals. The terms used here are terms that are commonly used in a research environment and are not meant to be rigidly definitive.

This section includes amphibian dose rates for the common drugs and drug regimens. If you need to use other drug agents, check with your institution's veterinary staff for assistance in determining a dose rate appropriate for use in the species you are using.

Postoperative Analgesics

These drugs relieve pain without necessarily obtunding consciousness and are available in three classes of drugs – the opioids, the alpha-2 agonists and the nonsteroidal anti-inflammatory drugs (NSAIDs).

Opioids are drugs that derive from the opium poppy or synthetic analogs and that are useful for relieving moderate to severe pain. The following agents and doses have been used in amphibians:

• Morphine 10 mg/kg by intracoelomic injection every 4 hours (q4h).
• Butorphanol 25 mg/kg by intracoelomic injection q12h.

Alpha-2 agonists are sedative drugs that also exhibit analgesic properties. The following agent and dose has been used in some amphibians:

• Xylazine 10 mg/kg by intracoelomic injection q12-24h.

NSAIDS are non-steroidal, anti-inflammatory drugs, and act (at least in part) by antagonizing inflammatory mediators such as cyclooxygenase. The following agent and dose has been used in some amphibians:

• Flunixin meglumine 25 mg/kg by intracoelomic injection q4h.

It should be noted here that clinical data on the use of analgesics in amphibians are scant. At best, the extended effects of these drugs on the animals and their physiology are poorly understood.

Sedatives

Sedatives may obtund consciousness but in normal doses do not do so sufficiently to ablate the perception of pain or other sensations. When combined with general anesthetics, they may be used to induce "balanced" anesthesia where muscle relaxation, unconsciousness, and analgesia are enhanced. Used by themselves, these drugs may decrease anxiety, fear or excessive activity.

Anxiety and fear are not conditions that have been consistently recognized in amphibians, so indications for using sedatives in amphibians(except as noted above for analgesia with xylazine) have not been evident.

Anesthesia refers to the complete blockade of pain and it may be applied by blocking local nerve receptors or by blocking the central perception of sensory impulses by the brain.

Local Anesthesia

For minor procedures such as tissue biopsies or microchip implantation, infiltration with local anesthetics has been found to be safe and effective. Lidocaine is probably the agent most commonly used. Care should be taken to limit the infiltration to the area involved, because large doses of local anesthetics can have systemic effects that may be toxic.

General Anesthesia

Surgical anesthesia is indicated by loss of righting reflex, loss of abdominal and gular respiratory movements, absence or corneal reflex, and lack of response to stimulation. Respiration will occur transcutaneously and so it is important that the skin be kept moist throughout the procedure and recovery.

Anesthetic agents may be injected or administered transcutaneously using a water bath. Agents that have been reported include:

• MS 222 (Tricaine methanesulfonate) is one of the more commonly used agents. It may be injected into the coelom or the dorsal lymph sacs, or it may be prepared as a bath of 0.1%-0.3%. MS 222 is acidic and should be buffered with bicarbonate to a pH of 7.0-7.5. The solution is rinsed away with fresh water when surgical anesthesia has been achieved. When used as a bath, induction usually takes 10-30 minutes, anesthesia lasts approximately 40 minutes and recovery may take as long as 24 hours. Anesthesia can be maintained by dripping additional solution on the skin. Solutions of MS 222 should be stored away from light and should not be used if discolored or if older than 30 days.
• Benzocaine in solutions of 0.2% and buffered with bicarbonate have been used as a bath and are reportedly preferred to MS 222 by some investigators.
• Injectable agents include pentobarbital 40-50 mg/ml intracoelomically or into the dorsal lymph sacs; ketamine 20-40 mg/kg and diazepam 0.2-0.4 mg.kg IM; and tiletamine/zolazepam 10-20 mg/kg IM. These agents have reportedly produced erratic results and high mortalities and may not be the best anesthetic choices.
• Halothane or isoflurane can be administered at induction doses of 3-5% and maintenance doses of 1-2%. The gas can be administered in an induction chamber or bubbled through bath water. The tracheal opening is located at the base of the tongue and tracheal intubation can be accomplished in larger amphibians using plastic catheters modified for use as endotrachial tubes and a non-rebreathing anesthesia circuit.

Hypothermia has not been proven to diminish pain perception in amphibians and is not acceptable for anesthesia.

The Guide and Public Health Service Policy require that major, survival surgery in amphibians be performed using aseptic technique. However, certain modifications of those procedures as applied to mammals are needed.

An important protective mechanism against infection and dehydration in amphibians is the mucus layer on the skin. Therefore, during surgical preparation, care must be taken to minimize disruption of this layer. Moreover, many skin preparation agents are readily absorbed though the skin and may be toxic. Therefore, soaps, detergents and products containing alcohol or iodine must not be used for amphibian skin preparation. Some investigators have recommended that amphibian skin be prepared for surgery in much the same way as mammalian eyes, using copious irrigation with sterile water or saline. Others have reported success with irrigation followed by very resticted application of chlorhexidine, limited to the immediate site of the incision and applied using a cotton-tipped applicator. The skin of the entire amphibian should be kept moist throughout the procedure.

Instruments should be sterile. If sterilized chemically, instruments should be rinsed thoroughly in sterile water before use. A sterile surgical field should be provided with appropriate draping, and sterile gloves should be used by operators.

Non-absorbable, monofilament suture material (such as nylon) should be used and sutures should be removed by 30 days after surgery.

Recovery from anesthesia may be hastened by rinsing the animal in fresh water, maintaining ambient temperature from 72 degrees to 85 degrees F, keeping the skin moist and humidity greater than 70%.

Amphibians should not be immersed in water until they have fully recovered from anesthesia because, paradoxically, that may result in drowning.

Animals should be observed until they are fully ambulatory. After recovery, they should be checked at least daily. Amphibians are highly resistant to hypoxia so simple unresponsivness should not necessarily be equated with death. Attempts to resuscitate an unresponsive amphibian may be effective. Death cannot be assumed until rigor mortis develops.

The term euthanasia is derived from Greek and means "good death." Animals should be euthanatized when killed for any purpose, including research. To euthanatize an amphibian, you must be trained in the concepts of euthanasia, the method to be used, and the proper handling of particular amphibian species involved.

The 2000 Report of the AVMA Panel on Euthanasia lists methods that are acceptable, conditionally acceptable and unacceptable for a large range of species, including ectotherms. This lesson will not cover them exhaustively, but a complete list and discussion of these methods may be reviewed by clicking on the link to the American Veterinary Medical Association.

Euthanasia of amphibians must take into account differences in metabolism, respiration and tolerance of cerebral hypoxia relative to mammals. Adult amphibians may be humanely killed using one of the following methods:

• An overdose of anesthetics such as injectable sodium pentobarbital, 60 to 100 mg/kg, intracoelomically or into the dorsal lymph sacs.
• Buffered MS-222 (0.1-1% solutions) or benzocaine hydrochloride (at concentrations >250 g/L) may be used in a bath or by intracoelomic or dorsal lymph sac injection.
• Decapitation with heavy shears or a guillotine is effective for some species that have appropriate anatomic features. It has been assumed that stopping blood supply to the brain by decapitation causes rapid loss of consciousness in warm-blooded species. Because the central nervous sys-tem of amphibians is tolerant to hypoxic and hypotensive conditions, decapitation must be followed by pithing the brain.
• Severing the spinal cord behind the head by pithing is used in some ectotherms. However, death may not be immediate unless both the brain and spinal cord are pithed. For these animals,pithing of the spinal cord should be followed by decapitation and pithing of the brain. This process of pithing the spinal cord and brain is sometimes called "double pithing". The pithing site in frogs is the foramen magnum, and it is identified by a slight midline skin depression caudal to the eyes when the neck flexed.
• Direct cranial concussion, though aesthetically objectionable, is humane when properly performed.

Many amphibians can hold their breath and survive periods of anoxia (up to 27 hours for some species). Therefore, euthanasia of amphibians using inhalation agents such as CO2, is not acceptable.

Cooling or freezing alone are not acceptable methods of euthanasia as formation of ice crystals in the tissues of an animal may cause pain and distress. Quick freezing of deeply anesthetized animals is acceptable.

Animals that have been euthanized by means of toxic substances or drugs (including euthanasia agents like pentobarbital or MS-222) must not be disposed of in areas where they may become part of the natural food web.

Federal Laws, Regulations, Policies:

1. Health Research Extension Act of 1985, Public Law 99-158, November 20, 1985, "Animals in Research".
2. Public Health Service Policy on Humane Care and Use of Laboratory Animals, Revised September, 1986, Reprinted March, 1996.
3. U.S. Government Principles For The Utilization And Care Of Vertebrate Animals Used In Testing, Research, And Training, Interagency Research Animal Committee.
4. Endangered Species Act Regulations, Code of Federal Regulations, Title 50, Part 17
5. CITES Regulations, Code of Federal Regulations, Title 50, Part 23

Guidelines:

1. Guide for the Care and Use of Laboratory Animals, National Research Council, 1998.
2. Guidelines For Use of Live Amphibians And Reptiles in Field and Laboratory Research, 2nd Edition, Revised by the Herpetological Animal Care and Use Committee (HACC) of theAmerican Society of Ichthyologists and Herpetologists, 2004.
3. Guide to Care and Use of Experimental Animals, Canadian Council on Animal Care, Volume 2, 1984.
4. Guidelines for Anesthesia, Postanesthetic Care and Surgery of Amphibians, Prepared by the ULAM Veterinary Staff July 20, 1998

Articles:

1. DeNardo, Dale "Amphibians as Laboratory Animals", ILAR Journal V37(4) 1995.
2. Machin, Karen "Amphibian Pain and Analgesia", Journal of Zoo and Wildlife Medicine, 30(1) 2-10, 1999.
3. Stevens, CW, MacIver, DN, Newman LC, "Testing and Comparison of Non-opiod Analgesics in Amphibians", Contemp Top Lab Anim Sci, July, 40(4):23-7, 2001.

Texts:

1. O'Rourke, Dorcas P and Schultz, Terry Wayne, "Biology and Diseases of Amphibians" in Laboratory Animal Medicine, 2nd Edition; Elsevier Scince 2002.
2. Lumb and Jones, Veterinary Anesthesia 3rd Edition, ed. Thurmon John C, Tranquilli William J, and Benson G. John, Williams and Wilkins, 1996
3. Kreger, Michael D., "Laboratory Housing for Reptiles and Amphibians" in Comfortable Quarters for Laboratory Animals, ed. Viktor Reinhardt, Animal Welfare Institute, Wash. D.C., 1997