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Clinical Techniques in Amphibians

By Brent R. Whitaker, MS, DVM, National Aquarium

All possible routes of escape from the examination room, such as ventilation ducts and sink drains, should be blocked before handling amphibians. Recommended supplies include a mist bottle containing dechlorinated water, which can be used to keep amphibians moist when handled, dip nets, a small air pump with airline and air stone, a water quality test kit, a small room humidifier, tryptic soy broth blood culture vials, the anesthetic tricaine methane-sulfonate, and a microliter syringe. Fine-tipped culture swabs, glass slides, coverslips, scalpel blades of various sizes, an assortment of needles and syringes, sterile red-rubber tubes, and sterile saline should also be readily accessible.

The history should include a description of the animal’s diet and appetite; environmental conditions of the animal’s habitat, including humidity, temperature, water quality measures, and lighting regimen; social structure and reproductive status; the recent introduction or loss of animals; and the use of medications. Problems noted by owners should be described in detail. A review of food and water quality records is useful to identify important trends. A water sample from the animal’s enclosure should be analyzed for ammonia, nitrite, pH, hardness, alkalinity, and copper using a simple test kit readily available from most pet stores. Owners must take and record air and water temperatures at the time of water collection.

Before handling, the animal’s body condition, agility, posture, and behavior should be noted. Parasitic or microbial infections, malformation, or nutritional deficiencies may cause asymmetry. Loss of muscle mass commonly occurs as a result of improper nutrition, improper environmental temperatures, or chronic disease (eg, mycobacteriosis, chromomycosis, microsporidiosis). Neurologic impairment may be detected by first watching the animal move about its enclosure and then assessing its response to the introduction of stimuli. Neurologic impairment may also be suspected if an amphibian is unable to maintain equilibrium or exhibits an abnormal swimming pattern. When handled, most normal amphibians attempt to escape, withdrawing limbs that are grasped. Placed upside down, most species will attempt to right themselves. Touching the eyes typically elicits a blink reflex or withdrawal of the globe.

A cool, bright light and magnification are required when performing a physical examination. The mouth can be opened using the edge of an index card, a plastic card, or a rubber spatula. The color of the mucous membranes should be evaluated and any lesions noted (eg, retrobulbar injury, orogastric intussusception). Ulcerations, erythema, hemorrhage, and pigment loss in the skin may indicate poor husbandry, trauma, or infections (microbial or parasitic). Improper substrate or sanitation leading to bacterial and fungal infections can cause lesions on the feet. Touch preparations or skin scrapings of affected areas are easily made and can be stained with Wright-Giemsa and Gram stains for cytologic evaluation. Heart rate can often be determined by watching the skin overlying the xiphoid or using a hand-held 8-MHz transcutaneous Doppler system. Because pulmonic respiration (if present) depends on positive-pressure ventilation from buccal pumping, respiratory rate should be assessed by watching the rapid movements of the intermandibular space. The nares should be free of obstruction from mucus, which may indicate respiratory disease. Rhabdias spp, a nematode that has a direct life cycle, is a common cause of respiratory infections in captive amphibians. Eggs or larvae may sometimes be detected in oropharyngeal mucus. Ocular lesions are often detected and may include conjunctival, corneal, iridal, and lenticular changes. Corneal diseases, including nonspecific keratitis and lipid keratosis, are common. Corneal scrapings are easily collected for cytologic examination. Panophthalmitis and uveitis are associated with systemic or localized infection. A sample of aqueous or vitreous humor can be collected with a small-gauge needle for cytology and for bacterial and fungal culture. Coelomic palpation may detect retained egg masses, bladder stones, foreign bodies, or neoplasms. Hydrocoelom and subcutaneous edema (anasarca and ascites) are common and may be caused by lymph heart failure; cardiac failure; renal, GI, or hepatic disease; neoplasia; microbial infection; parasitism; toxicosis; improper environmental conditions; or other unknown factors. Collection of fluid for biochemical analysis, cytologic evaluation, and culture for bacteria and fungi is recommended. Blood collected from the ventral abdominal vein, lingual vein, femoral vein, coccygeal vein, or by cardiac puncture and placed into lithium heparin can be used for hematologic evaluation. A volume equal to 1% of the body weight of a healthy amphibian and 0.5% of the body weight of a sick amphibian may be taken. Normal values have not been established for most species of amphibians. Urine may be collected for analysis from those anurans that urinate when first restrained. Fecal samples uncontaminated by environmental organisms may be collected from species such as dart frogs by feeding the animal just before placing it on a clean, moist paper towel. Direct and float examinations are useful to identify protozoa and metazoa.

Treatments are administered orally, topically, or by injection. Oral administration requires firm restraint and opening the mouth with a piece of waterproof paper or thin piece of plastic (such as a credit card). Guitar picks, which come in a variety of thicknesses, work well. Small amphibians can be dosed accurately using a microliter syringe. Many drugs may be administered topically; the skin of most amphibians will absorb drugs directly. Some drugs, such as enrofloxacin, may be irritating, and alternative routes may be preferable. Treatments may also be delivered topically by placing the amphibian in a medicated bath. Bubble wrap or other nonabrasive material placed strategically over the amphibian may be needed to keep it in contact with the solution. Injections are typically given intracoelomically, into the lymph sacs, or intravenously.


Anesthesia may be required for further examination or for diagnostic and surgical procedures. Response to anethesthics depends on individual health and species of the animal. Tricaine methane-sulfonate, ketamine hydrochloride, propofol, halothane, isoflurane, and sevoflurane may be used. Routes of administration include immersion bath, topically, or parentally. Inhalant anesthetics are typically used topically or via an immersion bath, because they are readily absorbed through the skin. Larger amphibians can be intubated and maintained on anesthetic gas. Parental anesthetics are typically injected IV, IM, intracoelomically, or into the dorsal lymph sac. Injections into the rear limbs are avoided because of the presence of a renal-portal system. Heart rate can be monitored in most amphibians using an ultrasonic Doppler positioned over the heart. A baseline heart rate should be established before initiating anesthesia. Because amphibians can breathe through the skin, observing gular movement is less rewarding. Tricaine methane-sulfonate is a fine white crystal that is highly soluble in water. It can be prepared and stored as a 10 g/L stock solution, which is diluted just before use. Administration is by bath, because most amphibians will absorb tricaine through the skin. The dosage used for many large amphibians is 2–3 g/L for induction. For short procedures, the amphibian should be immediately removed and rinsed with fresh water. For longer procedures, the amphibian may be placed into a maintenance solution of 100–200 mg/L after it has been induced. In smaller amphibians, an induction dosage of 100–200 mg/L is safer. Aeration must be provided in the anesthetic solution to avoid hypoxia. Tricaine produces an acidic solution that must be buffered to a pH of ~7 using sodium bicarbonate, sodium hydroxide, or sodium hydrogen phosphate. Isoflurane gas can also be bubbled into an anesthetic bath placed in a sealed container; this will allow both percutaneous and inhalation absorption. Ketamine hydrochloride injected percutaneously or into the dorsal lymph sac at a dosage of 75–125 mg/kg body wt can be used; however, a surgical plane of anesthesia can be difficult to maintain, and recoveries are long. Intracoelomic injections of propofol at a dosage of 35–45 mg/kg body wt will also provide sedation to anesthesia.

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