Unique Anatomy of Frogs and Toads^3,4

MUSCULOSKELETAL

Four limbs are present, but there may be variable hindlimb lengths. The length depends on species locomotory modes; longer hindlegs occur in animals that jump (Figure 5-4). The forelimbs possess a fused radius and ulna (radioulna), whereas the hindlimbs have a fused tibia and fibula (tibiofibula). The forefeet possess four phalanges on each foot, whereas the hindfeet have five phalanges. The hyoid apparatus in some species is adapted to eject the tongue for prey capture. Vertebrae are separated into three fused regions: presacral, sacral, and postsacral (note that a sacrum is not present). The pelvic girdle is fused with the last presacral vertebra, and a fused urostyle (or coccyx) is found caudal to the girdle. Some animals possess pigments that result in a blue coloration of the bones (Phrynohyas resinifictrix). Amphibians do not have diaphragms.

Figure 5-4 Skeleton of Rana catesbeiana.

GASTROINTESTINAL SYSTEM

Teeth are absent in some frogs; others lack tongues (e.g., Pipa pipa, which uses negative pressure to create a vacuum to catch prey). Occasionally, frogs evert their stomachs in order to wipe noxious substances from their stomach mucosa. It is not unusual to see undigested insect skeletons, bones, or plant fibers in the feces of anurans. All amphibians (note: within this text, “all” amphibians will include caudates and caecilians) have a simple, short alimentary system that empties into a cloaca (Figure 5-5). A large bilobed liver (which can fill up to 50% of the coelom) encases the pericardium. There are typically two lobes, though a third will occur in some species. The color and shade varies from pale gray to brown to black. Melanomacrophages are present and can result in the dark coloration of the liver. In the larval amphibian, the liver is the primary hematopoietic organ, switching to bone marrow after metamorphosis. Anurans have a large gall bladder.

Figure 5-5 Gastrointestinal system (liver removed) of the frog (Rana pipiens).

Unique Anatomy of Newts and Salamanders^3,4

GASTROINTESTINAL SYSTEM

The liver is elongate and single lobed; it may have scalloped edges. The intestinal anatomy of newts and salamanders has no relevant clinical differences from other amphibians (Figure 5-6). The oral cavity hinges such that the maxilla is mobile while the mandible is static.

Figure 5-6 Basic anatomy of the caudate (Ambystoma mexicanum).

Unique Anatomy of Caecilians^2–5

REPRODUCTIVE SYSTEM

Male caecilians possess a phallodeum (intromittent organ), enabling them to practice internal fertilization. Caecilian ovaries are paired and are closely connected with the kidneys. Viviparous animals have an oviductal lining where embryos feed off the lining (Figure 5-7). Oviparity occurs in most caecilians, with some species-specific parental care occurring until hatching.

Figure 5-7 Caecilian (Typhlonectes natans) uterus with fetuses.

Basic Premetamorphic Anatomy^2–5

CAECILIANS

Aquatic caecilians are viviparous. Larval animals are born fairly precocious, although they do have gills (Figure 5-8) present for the first day after birth. Oviparous caecilian larvae also have external gills that are quickly resorbed, but they also possess larval fins, which disappear shortly before hatching. The larval caecilian has a pronephric kidney that changes to a mesonephric kidney during metamorphosis. This change alters the kidney’s function from a filtration system to a more complex excretory and water resorptive system.

Figure 5-8 Caecilian (Typhlonectes natans) newborn with external gills.

General Clinical Physiology of Amphibians³

Environmental Considerations^4–8

Amphibians can be aquatic, semiterrestrial, fossorial, terrestrial, or arboreal; thus, it is important for veterinarians to emphasize the importance of species-specific requirements.

AQUATIC

Animals that live in ponds versus streams have very different tolerance levels for water quality. Stream-dwelling animals generally have a greater need for more dissolved oxygen and higher quality water. In general, water quality should be maintained as for fish. Table 5-1 provides basic parameters to observe with most amphibians.

TABLE 5-1 Water Chemistry Parameters for Amphibians

Dissolved oxygen should be greater than 80% (>5 mg/L) to maintain the biological filter (e.g., aerobic cycle). Water filtration (bacterial and mechanical) should be maintained as with fish. Under-gravel filters are acceptable if the tubing does not allow animals to get underneath the plates. Some amphibians do not tolerate vibration well; therefore, care should be taken to place filters away from the enclosure if possible. A 10% to 20% water change should be done every week or two with aged or dechlorinated water. For caecilians, water depth should be shallow enough to allow animals to obtain air without much effort, especially newborns. For most amphibians, there should be a basking/dry-out area where the animals can leave the water.

Creating a Vivarium

Glass tanks are commonly used to house amphibians. They are well accepted because they allow excellent visualization of the amphibians, are relatively inexpensive, and hold moisture. The primary disadvantage associated with them is that they are not well ventilated. One way to improve ventilation is to have a secure, well-ventilated lid. The lid also helps prevent escape. Plastic tanks are also appropriate for making vivaria and may be more flexible for modification, though scratches can occur that obscure direct visualization of the animal. Lids that are hinged allow servicing to a portion of the enclosure while maintaining cover for the rest of the area. Solid plastic or glass lids should be considered when maintaining a high humidity is important. When these types of lids are used, it is important to still place some ventilation holes in them to prevent stagnant airflow.

Volatile organic compounds, such as those found in glues, need to be cured appropriately (under strict manufacturer conditions) before being used around amphibians; otherwise the fumes can leach into the environment and be irritating to the skin and respiratory epithelium.

Deionized water should not be used in an amphibian’s vivarium, as it can disrupt osmolarity. Chlorinated tap water also is not recommended. Distilled water or reverse osmosis water is ideal.

ACCESSORIES

It is important to take time and consider what accessories are needed in a vivarium, as these items can be used to mimic an animal’s natural habitat and reduce stress. Overplanting and overfurnishing a vivarium may make prey capture difficult for the animal. Plastic plants can be used to landscape a vivarium and are easily disinfected. Live plants can also be used and have the benefit of increasing environmental humidity. It is important to only use live plants that are known to be nontoxic, as prey being offered to amphibians may eat the live plants and transfer any toxin to the amphibian.⁶ If live plants are chosen, the pet owner should check to see if harmful pesticides or fertilizers have been used. Live plants may be contaminated with parasitic ova (e.g., from Florida, where tree frogs live in the plants). Recently acquired live plants should be cleaned and their soil should be replaced to reduce the likelihood of contaminating the vivarium with unwanted pathogens. Aquatic plants can harbor snails, which can introduce parasites to a vivarium as they serve as intermediate hosts for a number of parasites.

“Furniture” considerations for a vivarium may include large rocks, branches, shelters, and waterfalls. Traumatic injuries may occur with such items if they are not secured, and all furnishings should have smooth edges. These items should also be conducive to sterilization or disinfection. Bear in mind that porous objects may retain disinfectants. Proper rinsing is critical. Shelter type will depend on the natural camouflage of the animal and should be provided. Such accessories are easily created (Figure 5-9).

Figure 5-9 Frog housing made out of PVC and moss.

Hospital Tank

Veterinarians working with amphibians should maintain hospital tanks to treat cases presented to their facility. Smaller plastic containers with snap-on lids and small doors on the lid make access to the animal easy and safe. Bubble wrap placed on the bottom of the hospital tank provides a soft substrate, which stays moist and is atraumatic. Plastic wrap placed around the entire tank can be used to help maintain humidity, but the habitat should be serviced enough to provide adequate ventilation. This is not recommended for permanent or semipermanent housing, where the air may stagnate and predispose amphibians to conditions like pneumonia. Sphagnum moss, if used, should be thrown out weekly or when soiled, or more frequently if the patient has a known infectious disease. The hospital tank should be easily disinfected.

Feeding Mechanisms

Larval anurans are filter feeders.¹¹ Water is pumped through the mouth, and then planktonic algae are extracted and entrapped by a mucus-covered filter in the pharynx and then directed into the esophagus. Adult, terrestrial amphibians use their tongues to capture prey. These animals have glands on their tongues, which produce a sticky substance that helps apprehend prey until it is securely placed in the mouth. Some anurans can shape their tongue into a tube to assist in feeding on ants, termites, and worms. Aquatic salamanders direct prey to their mouths by creating an inward flow of water into the oral cavity. Pipa pipa direct food and water into their mouth with hypobrachial pumping movements. Some aquatic amphibians use their long phalanges to direct the food into their oral cavity.

Feeding and Diet

Obesity can occur in amphibians. In captivity, there is a tendency to overfeed animals, either by offering too much food or offering food too frequently. As animals get older, the frequency with which food is offered can be decreased (e.g., ambush predators should be offered food every 2 weeks). Larval animals and smaller frogs should be fed daily. Nocturnal animals should be fed at night or late in the day; diurnal animals should be fed once a day, and sometimes twice daily, depending on species.

The list of possible invertebrates to offer amphibians is extensive.¹² In general, the most common food items offered to terrestrial amphibians include earthworms, red worms, mealworm larvae, waxworm larvae, crickets, fruit flies, other larvae, occasionally small reptiles (e.g., anoles), and amphibian eggs and larvae. For aquatic species, these same foods are offered, but bloodworms, blackworms, and crustaceans, such as crayfish and brine shrimp, may be offered as well. Some animals have very specific diets, like the termite-eating caecilian (Boulengerula spp.), and can be difficult to maintain in captivity.⁵ Avoid brightly colored insects,⁶ as they may be naturally toxic. Finally, although wild-caught invertebrates would provide enrichment and balance for an amphibian, caution must be exercised not to feed out organisms that have been subject to pesticides or other such toxins.

Feeding pelleted diets intended for fish or reptiles may not be appropriate for amphibians, as these diets may be intended for herbivores or omnivores and will not meet the fat, protein, and vitamin requirements of amphibians.¹² Rodents can be offered on occasion, but vitamin A levels are often too high in these prey items, and secondary nutritional hyperparathyroidism can occur without supplementation of vitamin D₃. On the other hand, oversupplementation with vitamin D₃ may cause mineralization of organs. Unfortunately our knowledge regarding vitamin D₃ metabolism in amphibians is limited.

Supplementing amphibian diets with vitamins and minerals is a questionable but necessary practice. With a truly well-balanced diet, supplementation is unnecessary and may even cause problems (e.g., hypervitaminosis A and D). Unfortunately, with the limited number of commercially available food items, providing a well-balanced diet is difficult. Based on this limitation, it is best to offer the largest variety (e.g., multiple prey species) of food types to provide the best nutrition. Most supplements target calcium and vitamin D₃ in an attempt to balance the Ca:P ratio (1.5 : 1 or 2 : 1). Contrary to that target, most invertebrates, with the exception of earthworms, have an inverse Ca:P ratio.¹² Invertebrates must therefore be “gut-loaded” with vitamins and minerals by feeding them a diet that is high in Ca and low in P (available commercially) in the 24 hours before offering that prey item. Some of these diets can result in death of the prey item if fed longer than 24 hours. Additionally, these diets are not balanced for invertebrates and should not be used long term. Another product used for increasing the nutritional value of invertebrates is to “dust” them with a supplement. Placing the supplement and invertebrates in a brown bag and gently shaking the bag will result in the invertebrate being covered with the supplement. Dusting can be used with small invertebrates where gut loading is not practical or lethal, and this should be done immediately before feeding.¹² Dusting larger invertebrates with these supplements does not successfully achieve the calcium levels that gut loading does. When feeding frozen fish, it is necessary to supplement the diet with thiamine as the fish produce natural thiaminases that destroy the thiamine in the tissues.

Quarantine¹³

All new animals must be quarantined before being added to an extant collection of animals. Transporting animals and moving them into a new environment are exceptionally stressful events. Even though it is impossible to completely remove these stressors, they can be minimized once the animal is in quarantine. This can be accomplished by providing a quiet room, appropriate furniture in the new enclosure (e.g., a hiding spot, such as a film canister or plastic plants), and not overcrowding animals. Accessories must be easily disinfected. Partially covering the enclosure can also help reduce visual stressors. Multicolored papers (comic sections of newspaper) taped to the outside of the enclosure are excellent covers because a single color may not be easily perceived; avoid black, as it may act as a mirror and stimulate striking behavior.

Quarantine should last a minimum of 30 to 60 days for captive raised animals but should be extended to 90 days for wild-caught specimens. All in/all out principles should apply. Animals in quarantine should be serially examined for pathogens and parasites. A standard schedule may include three fecal examinations conducted a week apart. Prophylactic treatment is warranted when an animal is in poor body condition, has a normal appetite, and has a fecal exam that shows excessive numbers of flagellates, nematodes, trematodes, or cestodes. Whereas some of these organisms are normal or incidental flora, large numbers in a compromised patient may warrant a course of antiparasitics (e.g., metronidazole). Utensils used for an enclosure should not be shared with another group of animals.

Fecal samples can be collected by placing the animal into a disinfected and rinsed opaque, covered plastic container. The container should be well ventilated, and animals should not be kept in the containers longer than 12 hours. Several fecal samples should be collected to evaluate the status of the animal. It is important to remember that some parasites shed transiently, so multiple (serial) samples may be required to confirm a diagnosis. I perform the following exams when screening amphibian feces: fecal floatation with commercial hyperosmotic agent, direct saline smear, acid-fast stain, and Gram stain.

Routine Exams

A baseline examination on any amphibian presented to a veterinarian is recommended. In large colonies, or cases with exceptionally small animals, this may be impractical. The minimum database collected during an amphibian examination should include a physical exam, radiographs, weight, and, if possible, blood examination. Individual or group fecal examinations should be performed at least yearly. Visual examinations of every animal should be performed daily.

Disinfection and Sanitation

Utensils, accessories, enclosures previously occupied by amphibians, and enclosures with known disease entities, should be disinfected to prevent the spread of disease. Removal of all debris is necessary before application of a disinfectant, as some products are deactivated in the presence of proteins and other organics. Warm water and surfactants can be used to help break down organics during the cleaning process. Numerous classes of disinfectant are available, each with variable effectiveness against certain organisms (Table 5-2). Some products absorb the chemicals and can leach out while the amphibian is present (e.g., iodine products). Ammonia and chlorine bleach are good general disinfectants. With all chemical products, thorough and repeat rinsing is mandatory. For most disinfectants, a minimum contact time of 10 minutes is recommended. For more fastidious organisms, longer contact times are recommended. Porous objects should be discarded rather than reused, not only because they harbor pathogens but also because it is difficult to thoroughly rinse and remove the chemicals from these materials. Humidifiers should be disinfected weekly, as should any aerosolizing tools. Povidone iodine, chlorhexidine, isopropyl alcohol, and quaternary ammonium compounds are known to cause skin lesions and to be toxic in amphibians at high concentrations, and thus should be used with caution.¹³

TABLE 5-2 Disinfectants Commonly Used to Clean Amphibian Environments

In a healthy biologic environment (vivarium or aquarium) with normal microflora, the opportunity for pathogens to be a problem is minimal. To assist in maintaining a healthy biologic system, unnecessary organic loads can be reduced by spot-cleaning feces and other organic material in the enclosure and by changing out sections of soil, sand, and leaf litter regularly. Water changes of 10% to 20% should be performed weekly in aquatic systems to reduce both organic and inorganic loads on the system.

Manual Restraint¹⁵

Animals can be placed into containers for close visual examination without touching the animal. Suitable containers include small clear jars or boxes and plastic bags. Occasionally animals will feign death, but this must be distinguished from exhaustion. Under such a circumstance, place the animal in its traveling enclosure and wait. Avoid handling larval amphibians if possible.

Large frogs should be handled by firmly grabbing the animal around the inguinal region just cranial to the hindlimbs (Figure 5-10, A). Smaller frogs can be cupped into the palm of the hand for limited access or, while in this grip, gently restrained by the hindlimbs and caudal coelom using the opposing hand. When handling an animal in this way, it is important to take care not to squeeze or traumatize the animal (Figure 5-10, B). Expect anurans to urinate when handled, and be prepared to collect the sample if desired. Additional behavioral manifestations identified after handling may include breath holding or air sac insufflation, color change, and vocalizing.

Figure 5-10 A, Handling technique for a large frog. B, Handling technique for a small frog.

Salamanders and newts should be handled at the pectoral girdle initially, then the hindlimbs (Figure 5-11). Tail autotomy can occur in these animals, and care should be taken to not hold the animal by its tail.

Figure 5-11 Two-hand technique for handling a newt.

Caecilians are very difficult to manage with manual restraint. Clear tubing or a plastic bag can limit the mobility of these animals and facilitate examination (Figure 5-12). For a more thorough examination, the animal should be sedated.

Figure 5-12 Handling technique for a caecilian using a sealed plastic storage bag.

Chemical Restraint

Before immobilizing an amphibian, fasting the animal is generally recommended to avoid aspiration pneumonia; however, this has not been described as a common complication. In an urgent situation, immediate anesthesia is recommended over delaying to fast the animal.

Local anesthesia can work very well on amphibians. Lidocaine can be injected or placed topically in the perimeter of a lesion. Emla cream (Astra Pharm, Westborough, MA) may also be used and can induce local anesthesia in minutes. Both may result in general anesthesia or death if caution is not exercised and too much product is used.

The most common general anesthetic agent used for amphibians is tricaine methane sulfonate (MS-222) (Finquel, Argent Chemical Laboratories, Redmond, WA). The drug comes in a powdered form and is usually reconstituted in distilled water. MS-222 acts by reverting transient sodium ion permeability and decreasing excitability and blocking conduction of nerve impulses.¹⁶ This compound is very acidic and should be buffered with an equal volume of baking soda to stabilize the pH. Unbuffered MS-222 can be highly irritating to the amphibian, causing excitability and erythema. Unbuffered MS-222 can also result in a more rapid induction because uptake across the skin is more rapid in the ionized form. Although buffering the agent will result in slower uptake, it is far safer for the animal (e.g., hyperglycemia and tachycardia have been reported with unbuffered MS-222).¹⁷ Induction at 100 to 200 mg/L will achieve steady state levels in most amphibians, but induction may be prolonged for more than 25 minutes. Higher doses (1-5 g/L) may induce anesthesia faster, but these levels are considered potentially lethal. Higher dosing levels can also prolong recovery. At induction, animals may try to jump out of the solution, as it can be irritating. Anesthetic deaths are rare with MS-222, but they are possible. Under MS-222 anesthesia, pulmonary ventilation is diminished or absent. Gas exchange across the skin is usually adequate to prevent hypoxia at low (<74° F) environmental temperatures. Once anesthetized, the animal’s nares and oral cavity must be kept out of the water to avoid drowning. Keeping the animal at room temperature (74°–78° F) usually results in acidosis and hypercarbia. Oxygen can be aerated into the immersion solution, and this usually helps the animal compensate. MS-222 has been used intramuscularly (IM) and intracoelomically (ICe); however, I have not used these routes. When using these alternative routes, expect prolonged recoveries.

Gas anesthesia has been used in a variety of forms.^16–18 Using the vaporized form results in direct exposure to the inhalant, as with a face mask (Figure 5-13), and can induce hypothermia and/or dehydrate the animal. Therefore, when using this form of anesthesia, it is important to monitor the animal’s hydration status and body temperature. Using direct exposure appears to be more successful in toad species than frog species. Dosing anurans percutaneously with topical isoflurane and halothane has also been successfully done, but it can induce a very deep plane of anesthesia and prolonged recoveries. Although some clinicians report that aerating an inhalant directly into water is useful, I have not had success with this method. Another potential complication with aerating the anesthetic into the water is that scavenging excess gas may not be possible, posing a human health risk. Finally, a gel mixture of a sterile KY jelly lubricant with isoflurane or sevoflurane and water mixed in a 3 : 3:1 ratio can be utilized. The mixture may be applied directly on the animal or via a gauze (Figure 5-14). This method results in rapid induction and recovery.

Figure 5-13 Anesthetic induction of a frog using a small animal facemask as a gas chamber.

Figure 5-14 Anesthetic induction of a frog using a gauze soaked in isoflurane and water-soluble gel.

Clove oil (eugenol) has been used to anesthetize leopard frogs (Rana pipiens) and tiger salamanders (Ambystoma tigrinum).^19,20 The dose required to induce surgical anesthesia in the salamanders (0.23 ml of 85% clove oil/500 ml dechlorinated water) is much higher than in the frogs (333 mg/L). Propofol was also evaluated in both species. Delivery in the frogs via the lingual vessels was difficult and produced only mild effects. When Propofol was administered intracoelomically in the tiger salamanders (35 mg/kg), surgical anesthesia was achieved in the majority (73%, 8/11) of test subjects.²⁰

Monitoring the Anesthetized Patient

The amphibian heart rate can be monitored using a pulse oximeter, Doppler, or ultrasound. Respirations can be difficult to monitor. The best methods to use are to evaluate the excursion of the rib cage or the gular area. Oxygen supplementation is important, as hypercarbia and hypoxia are common during anesthesia. However, too much oxygen may limit respirations, based on the theory that these animals are stimulated to breathe based on low oxygen levels and not high carbon dioxide levels. Based on this, it is generally preferred to recover these animals on room oxygen (21%). Readings of 95% oxygen saturation or greater are expected in a normoxic patient.¹⁵ If bradycardia occurs, reduce or end the anesthetic event. Atropine (IM or IV) can be used to increase the heart rate. If asystole occurs, begin cardiac compressions. Cardiac compressions can be used to re-stimulate the cardiac function. There are two methods that may be used: (1) Place the animal in dorsal recumbency. Then place an index finger on the cranioventral rib cage (e.g., level of the pectoral girdle) and compress downward. This will press the heart against the spine and act as a stimulus to restart the heart. (2) Grasp the lateral rib cage at the level of the pectoral girdle and squeeze inward. Epinephrine can be given intratracheally, intracardiacally, or intravenously to initiate cardiac function.

When monitoring the depth of anesthesia in amphibians, it is important to follow a consistent pattern and record all measurements. The following measurements are generally used: righting reflex, escape response, palpebral reflex, superficial pain response (e.g., pinch skin over dorsum of foot), and deep pain response (e.g., apply pressure to one of the rear digits). It is important to evaluate those reflexes before starting the anesthetic event, to provide a baseline for that animal. The generally accepted method for characterizing these reflexes is based on an ordinal scale: 0-present, 1-mild loss, 2-loss. Based on this scale, surgical anesthesia is achieved when all of the reflexes are recorded as a 2 (loss).

Recovery from water-bath–based anesthesia (e.g., MS-222, clove oil) can be accomplished by placing the animal into fresh, dechlorinated, anesthetic-free water or thoroughly rinsing the anesthetic off of the animal with the same type of water. When recovering the animal in a clean water bath, the animal should be monitored closely to prevent accidental drowning. The amphibian should be considered recovered when all of the reflexes have returned (0-present) and the heart rate and respiration rate are not different from the preanesthetic exam.