Topics in Medicine and Surgery

Anesthetic Considerations for Amphibians
Mark A. Mitchell, DVM, MS, PhD

Abstract
The popularity of amphibians in research, zoological exhibits, and as pets is on therise. With this increased popularity comes a need for veterinarians to develop
methods for managing these animals for various diagnostic and surgical procedures. For many of these procedures, the provision of anesthesia is a must.
Fortunately, there are a number of different anesthetics available to the veterinary
clinician for anesthetizing amphibians, including tricaine methanesulfonate, clove
oil, propofol, isoflurane, and ketamine. In addition to the variety of anesthetics at
our disposal, there is also a wider range of methods for delivering the anesthetics
than are generally available for higher vertebrates, including immersion, topical,
and intracoelomic routes of delivery. The purpose of this article is to review thedifferent methods that can be used to successfully manage an amphibian patient
through an anesthetic event. Copyright 2009 Elsevier Inc. All rights reserved.
Key words: amphibian; anesthesia; anesthetic; anuran; monitoring; urodelanA

mphibians are commonly used in research
and are popular exhibit animals in zoologicalcollections. Although they are not as popular
as reptiles in the pet trade, they are being presented
to veterinarians in clinical practice with increased
frequency. Because veterinarians are being asked towork with these animals, it is important that wedevelop standard protocols for managing them incaptivity.
Historically, anesthesia has been a specialty inherpetological medicine that has received little attention. Procedures were often completed with ananimal being manually restrained or cooled to minimize movement. Because these methods would not
be acceptable for birds or mammals, they should also
not be considered acceptable for reptiles or amphibians. Fortunately, in the past 2 decades we have seen
an increase in the number of scientific studies evaluating anesthetics in amphibians. Although many ofthese articles have been directed toward a specificspecies, Xenopus laevis, because of its value in research, the information obtained does have application across amphibian species.1-4 However, it should
not be assumed that all 4500⫹ species of amphibians

40

will respond to an anesthetic the same way. In cases
in which there is no reference for a particular species, special precautions should be taken to minimize the likelihood of complications.
Regardless of the species, success with anesthesia
can be achieved by following a consistent, wellplanned protocol. For the author, the most pressing
points to consider when establishing a protocol are
the preanesthetic examination, the selection of ananesthetic that will allow the veterinarian to performthe desired procedure safely, an anesthetic-monitoring protocol that will help to determine the level ofanesthesia, and a postanesthetic plan that considers
the recovery and management of the animal afFrom the University of Illinois, College of Veterinary Medicine,
Department of Veterinary Clinical Medicine, Urbana, IL USA.
Address correspondence to: Mark A. Mitchell, DVM, MS, PhD,
University of Illinois, College of Veterinary Medicine, Department
of Veterinary Clinical Medicine, 1008 W. Hazelwood Dr, Urbana,IL 61802. E-mail: mmitch@illinois.edu.
© 2009 Elsevier Inc. All rights reserved.
1557-5063/09/1801-$30.00
doi:10.1053/j.jepm.2008.11.006

Journal of Exotic Pet Medicine, Vol 18, No 1 ( January), 2009: pp 40 – 49

Anesthetic Considerations for Amphibians

ter the completion of the procedure. The purposeof this article is to review the methods that can beused to successfully manage anesthesia in captive
amphibians.

Preanesthetic Examination
A preanesthetic examination should be performedon each patient to determine whether an animal is asuitable candidate for an anesthetic procedure. The
examination should include both a physical evaluation of the patient and supportive diagnostic tests,
such as bloodwork, to help to characterize the physiologic status of the patient. Before performing an
examination, however, it is important to consider
the different techniques that can be used to safely
restrain the animal for the examination.
Amphibians, like fish, require some special consideration when planning for a procedure that requires restraint. The integument of both of thesegroups of animals is an important component totheir innate immune system, and damage to the skinduring an examination could compromise the patient. To minimize the likelihood of causing problems, an amphibian should be handled with care.
Latex or vinyl examination gloves are strongly recommended when handling an amphibian. The
smooth texture of the gloves will minimize the potential for injury to the amphibian’s integument.
The gloves should be premoistened with distilled ordechlorinated water. Wearing gloves also has a benefit to the individual restraining the amphibian.
Many amphibians can secrete toxins that can beabsorbed through the skin of the handler.5 The
integument of an amphibian can also be covered
with a variety of opportunistic pathogens (e.g.,
Pseudomonas species, Aeromonas species). The examination gloves can serve as a protective barrier against
both potential toxins and pathogens.
The physical examination of an amphibianshould include both a hands-off and hands-on reviewof the patient. The hands-off examination will enable the clinician to evaluate the breathing pattern,
general demeanor, and locomotion of the patient.
Animals that are dyspneic or depressed should be
handled carefully. Amphibians should have a naturalescape response when approached. Anesthetic procedures should be delayed in animals displaying abnormal behaviors.
For the hands-on examination, the animal may be
manually restrained. For small or fractious amphibians, chemical restraint may be necessary to complete a hands-on examination without injuring thepatient or handler. Amphibians do not generally

41
require a high degree of restraint for an examination. Urodelans can often be examined while being
held in an open hand. If restraint is required, thenthe animal can be gently grasped behind the forelimbs with an index finger and thumb, with theexaminer’s remaining fingers encircling the rearbody (legs and tail). Care should be taken whenhandling the tail, because tail autonomy can occur.
Because of their natural escape response, anuransneed to be secured quickly. For most anurans, they
can be secured by grasping them in front of theirrear legs. For larger specimens, such as the horned
frogs (Ceratophrys species), care should be taken tominimize the likelihood of being bitten. The
amount of restraint required for a caecilian depends
on the species and general demeanor of the animal.
Some caecilians can be examined similar to snakes,
whereas others need to be chemically immobilized
because they cannot be secured for an examination.
The hands-on examination of an amphibianshould be done in a thorough but rapid manner.
The oral cavity should be opened atraumatically.
Processed radiograph film or a rubber spatula can beused to gently pry open the mouth of an amphibian.
The mucous membranes should be moist and pale
pink in color. Increased tackiness may be an indicator of dehydration. The integument of the amphibian should be moist and free of defects. Animals thatappear dry should be kept moistened with distilled
or dechlorinated water. Body condition can be used
to determine the general health status of the amphibian. The muscling over the limbs of anurans andurodelans and the tail of urodelans can be evaluatedto assess body condition. In many amphibians, transillumination can be done to evaluate the coelomic
cavity/viscera. Positioning a pen light or transilluminator on the lateral body wall will enable the examiner to see various organ systems. For most species,
respirations can be measured by counting gularmovements or rib excursions. A Doppler ultrasoundcan be used to measure the heart rate. The probe
can be placed on the ventral aspect of the pectoral
girdle to obtain the heart rate.
Preanesthetic bloodwork can be used to assess thephysiologic status of an amphibian patient. Unfortunately, many of the animals we work with in captivity
present challenges for collecting samples or obtaining sufficient volumes of samples for screening.
Blood samples can be collected from the ventral tailvein of larger urodelans, the ventral abdominal veinof anurans and urodelans, the femoral vein of anurans and urodelans, and the sublingual vessels ofanurans. Cardiocentesis can be done, but can resultin cardiac arrest in some species. If only a small

42volume of blood can be collected, then a packed-cellvolume or blood smear can be done at a minimum.
This type of data can be used to determine thegeneral health of the patient and provide insight
into potential complications. For example, if a patient is anemic, then recovery from the anesthetic
event might be expected to be prolonged and theclinician can plan for it. Plasma chemistries can provide insight into the hydration status of the amphibian and general organ function. In the past, there
were often limitations with processing small sample
volumes; however, there is now equipment that can
provide complete biochemistry analysis on samples
as small as 100 ␮L of whole blood (VetScan 2; Abaxis, Union City, CA USA). If electrolytes are foundto be imbalanced, they can be corrected with fluids
before initiating a procedure. Likewise, if enzymes
are found to be elevated or there are alterations in
the minerals (e.g., inverse calcium to phosphorus
ratio suggestive of renal compromise), then a plan touse an anesthetic that will not affect that system(s)can be made.

Selecting an Anesthetic for an Amphibian
When selecting an anesthetic for an amphibian procedure, it is important to consider the type andduration of the procedure being performed. Procedures of short duration, such as diagnostic sampling,
can be managed using any of the possible anesthetic
methods, including immersion, topical, or parenteral administration; however, selection of the anesthetic in these cases should be determined based ona desire to have a rapid recovery. Procedures thatdemand a longer anesthetic period, such as surgery,require an anesthetic that will maintain a longer
surgical plane of anesthesia. The type of procedure
can also have an affect on the anesthetic selection.
For example, the application of a topical anesthetic
may be preferred over an immersion anesthetic for a
coeliotomy.

Mitchell

PropofolT (PropoFlo; Abbott Animal Health, Abbott
Park, IL USA), have also been used.
The water used to make anesthetic water for anamphibian should be of high quality. The author
always prefers to use water from the animal’s enclosure for the base of the anesthetic solution. To determine the value of the water, it is important toevaluate the ammonia and nitrite levels, hardness,
alkalinity, and chlorine/chloramine levels. If the water is deemed to be of good quality (e.g., no ammonia, no nitrite, neutral pH, good buffering capacity
and hardness, and no chlorine or chloramines), it
should be used as the base for the anesthetic. If it isof poor quality, then dechlorinated tap water can beused. The temperature of the anesthetic solution
should be similar to the preferred range of the amphibian. Anesthetic solution that is too cold may
result in a delayed anesthetic induction and prolonged recovery. If the anesthetic solution is too
warm, the amphibian may process the drug too rapidly and have an inconsistent anesthetic event. Aerating the water with an airstone and aquarium pumpwill ensure that the water is well oxygenated.
The induction chamber used for an immersion
anesthetic should be water tight and have a secure
lid to prevent escape. Plastic storage boxes with lids
work well as induction chambers (Fig 1). Sealable
plastic bags, glass tanks, and Styrofoam coolers havealso been recommended.5 Animals may become agitated during the induction period, and injuries associated with an escape response (e.g., jumping into
the side of the induction chamber) are possible.
Reducing the light and noise in the room can help
to limit complications. Lining the induction chamber with a protective material (e.g., bubble wrap) has

Immersion Anesthetics
Amphibians present veterinarians with a uniquemethod for delivering an anesthetic that is not available in the higher vertebrates: immersion. In manycases, the highly vascular integument of an amphibian is capable of absorbing different anesthetics at a
rate that can provide surgical anesthesia. The mostcommon compounds used to anesthetize amphibians via immersion are tricaine methanesulfonate(MS-222; Argent Laboratories, Redmond, WA USA)and clove oil (Spectrum Chemical Corp., Gardena,
CA USA), although other compounds, such as

Figure 1. A Ceratophrys species being anesthetized in an immersion of clove oil. A plastic shoe box was used as the induction
chamber.

43

Anesthetic Considerations for Amphibians

also been recommended.5 When inducing an amphibian in an immersion anesthetic, it is importantto keep the animal’s head and nares above the water
line. Failing to do so may lead to accidental drowning. To limit the likelihood of problems, the author
recommends keeping the anesthetic solution below
the level of the humero-scapular joint.6,7 It is important to monitor the patient closely during the induction period to reduce the likelihood of complications. Once anesthetized, the amphibian should
be removed from the anesthetic solution for theprocedure. If additional anesthesia is required, it canbe “dripped” onto the animal.
Apnea is a common finding in amphibians thatare anesthetized with immersion anesthetics. In
higher vertebrates, anesthetic-induced apnea canhave fatal consequences. In amphibians, this is also a
concern, but it is presumed that most amphibians
rely on cutaneous respiration during these types of
events.6,7 Therefore, it is important that the immersion solution have appropriate oxygen levels to limit
complications such as hypoxia. Aerating the anesthetic solution with a pump used for aquariums canbe done to ensure that oxygen saturation levels areappropriate (⬎5 ppt).

TopicalAlthough immersion anesthetics are certainly applied “topically,” there are a select number of anesthetics that can also be applied directly onto theintegument of an amphibian. A potential advantage
of using an anesthetic topically versus as an immersion is related to the volume and dose of anesthesia
required. Because topical anesthetics are applied
directly onto the integument, there is no dilutional
effect, and a smaller volume and dose can be used. A
potential disadvantage associated with this method,
however, is a greater potential for toxicity whendrugs are rapidly absorbed through the integument.
The most common anesthetics used topically areisoflurane and benzocaine.

Parenteral
Parenteral anesthetics have been evaluated and advocated in amphibians, but are less commonly usedthan immersion or topical delivery routes. The parenteral routes for administering anesthetics in mammals, including intramuscular, intraosseus, and intravenous methods, can also be used in amphibians.
In addition to these common routes of drug delivery,
injections into the dorsal lymph sacs and coelomic
cavity (Fig 2) can also be used in amphibians.5,7,8
Amphibians do have a renal portal system, so mostclinicians avoid intramuscular injections into the

Figure 2. Certain anesthetics, such as propofol, can be injected
intracoelomically in amphibians.

hindlimbs to limit the likelihood of complications.
Intravenous routes of anesthetic delivery are generally reserved for larger amphibians. The author has
had some success delivering anesthetics into the sublingual vessels, ventral abdominal vein, and femoral
veins of different anurans. Intraosseus routes of delivery are generally limited to the tibia or femur inanurans or large urodelans. Whether the renal portal system has an effect on anesthetics being delivered via this route is unknown. Intracoelomic injections appear to be well absorbed by amphibians,
although there is often a delayed clinical responsewith this method compared with that of other delivery routes.

Inhalant
Inhalant anesthetics are routinely used to anesthetize amphibians via immersion or topical routes;
however, they can also be delivered with the standard route via oxygen flow through the respiratory
tract. One of the limitations with the standard
method of delivery is related to the small size ofmany of our captive amphibian patients. The author
will commonly intubate and maintain larger amphibians with the standard route of delivery. In manycases, intravenous catheters, feeding tubes, or nasogastric/endotracheal tubes (1.0-1.5 mm tubes; Specialty Vet Products, Daphne, AL USA) can be used to
intubate amphibians. Because many amphibians
have a proportionally small cervical region, it is important to premeasure endotracheal tubes to prevent intubating a single lung. Amphibians are prone
to dessication when induced with an inhalant in an
induction chamber; therefore, it is important to line
the chamber with moistened paper towels to minimize the likelihood of complications.

44

Anesthetic Options for Amphibians
Tricaine Methanesulfonate
MS-222 is a water-soluble compound that is a derivative of benzocaine. This anesthetic is primarily used
as an immersion anesthetic in amphibians, although
it has been used parenterally too.9,10 Although mostof the original references to MS-222 anesthesia in
amphibians were related to anurans, more recent
work suggests that this anesthetic is also useful inurodelans.11,12 MS-222 is strongly acidic (pH: 3.0) in
solution, so it is important that the anesthetic wateris buffered to a neutral pH (7.0-7.5) before adding
the amphibian. Not buffering the anesthetic water
can have 2 negative impacts: 1) the acidity is toxic to
the amphibian and can lead to cutaneous lesions,and 2) the anesthetic becomes ionized and cannot
be absorbed. Buffering can be accomplished by adding sodium bicarbonate at a 1:1 ratio (dry weight) inmost cases; however, it is important to measure the
pH before starting an anesthetic event.
The recommended range of doses for MS-222
immersion anesthesia in amphibians is wide, withdoses of 0.2 g/L to 5.0 g/L depending on the age(e.g., larval form vs adult), size, and life history (e.g.,larger terrestrial bufonids may require a higher dosethan an aquatic species) of the animal.5,9 Downes9
has suggested that the doses used for amphibians
could be reduced if the water is buffered appropriately. Likewise, the author has found that lower dosing regimens can be used if the anesthetic solution isbuffered before use. In addition, induction itself
appears to be smoother when the anesthetic solution
is buffered, because some animals appear to be irritated by a more acidic solution. Most amphibians
can be induced within 10 to 30 minutes with thesedosing regimens.
It has also been suggested that MS-222 can bedelivered by the parenteral route, with recommendations for injecting the anesthetic into the dorsal
lymph sacs and intracoelomically.9,10 Although theseroutes can be used, it is important to consider theacidic nature of the drug. If the drug is to be used via
these routes, it should be buffered. Downes9 has
described a method for preparing MS-222 for parenteral delivery, although the additional compounds
required may not be readily available at most practices. Unless there is a specific need for delivering
MS-222 via a parenteral route, such as an experimentor limited access to high-quality water, the author
recommends delivering this drug via an immersion
method.

MitchellBenzocaine
Benzocaine can be used as an anesthetic for both
diagnostic and surgical procedures in amphibians.
Induction and maintenance of anesthesia with thisimmersion anesthetic are similar to those methods
described previously. Dosing regimens for adult amphibians are generally between 0.01% and 0.03%,
with lower doses recommended for larval and paedomorphic amphibians (0.005%-0.01%).12-15 Benzocaine, like the other immersion anesthetics, can induce apnea in amphibians. A recent study in leopard
frogs found that immersion in 0.02% benzocaine
resulted in apnea and a need to provide respiratory
assistance.15 It is important that the clinician is prepared to manage these complications to ensure success with their cases. Although MS-222 is generally
preferred over benzocaine for most amphibians, a
recent study comparing benzocaine with MS-222 inmetamorphic and paedomorphic tiger salamanders(Ambystoma tigrinum) suggested that benzocaine wasmore effective than MS-222. However, it is important
to note that the authors used a lower dose of MS-222(0.02%) than is generally recommended for amphibians and that they did not buffer the MS-222.12
Because of the availability and consistency of MS-222
and clove oil, the author does not routinely use
benzocaine for amphibians. However, individuals interested in anesthetizing large numbers of animals
may want to consider benzocaine over these othercompounds because it is less expensive.12

Clove OilClove oil is a naturally occurring compound that has
been used as an anesthetic in humans, mammals,
fish, and amphibians.3,6,7,16-19 The active ingredientof clove oil is a phenolic compound, eugenol. Cloveoil has been found to produce surgical anesthesia inboth frogs and salamanders.3,6,7
Dosing regimens for clove oil may vary dependingon the species and the life history of the animal. For
example, dosing requirements for leopard frogs(Rana pipiens) and African clawed frogs, both aquatic
species, were lower (318 -350 mg/L) than thoserequired for tiger salamanders (450 mg/L), a terrestrial species.3,6,7 The immersion methods used toanesthetize amphibians with clove oil are similar to
those described previously.6,7Attempts to deliver
clove oil topically or parenterally through the dorsal
lymph sacs in African clawed frogs were unrewarding.3 Induction times for anesthesia with immersion
in both frogs and salamanders were less than 15
minutes. Surgical anesthesia was achieved in all(12/12) of the leopard frogs and the majority(67%; 8/12) of the salamanders exposed to clove

45

Anesthetic Considerations for Amphibians

Figure 3. Gastric prolapse was a common side effect noted inleopard frogs anesthetized with clove oil.

oil. The salamanders were not exposed to the anesthetic solution for as long (10 minutes) as the frogs(15 minutes), and this may have accounted for thedecreased success with inducing surgical anesthesia.
All of the animals in both studies had uneventful
recoveries from the clove oil anesthesia, with recovery rates in the frogs and salamanders averaging 89
minutes and 75 minutes, respectively.
Clove oil was found to cause respiratory depression in the leopard frogs but not in the tiger
salamanders. Bradycardia was noted in both species,
although it was transient. Another side effect notedwith clove oil in leopard frogs was that 50% (6/12)of the frogs prolapsed their stomach after being
removed from the clove oil solution (Fig 3). The
gastric prolapse resolved spontaneously in all 6 animals and was attributed to the bad taste of the compound. The topical application of clove oil has been
associated with cutaneous necrosis in African clawedfrogs, so care should be used when considering theapplication of clove oil directly onto the skin of anamphibian.2

Isoflurane
Isoflurane is a popular anesthetic that can be delivered to amphibians as an immersion, topical, or
inhalant. Attempts at administering isoflurane parenterally (e.g., intracoelomically, intramuscularly,
and subcutaneously) have been made in African
clawed frogs, but the results were not as consistent orsafe as those reported for immersion or topical exposures.1 Stetter and coworkers20 used isoflurane as
an immersion anesthetic in both African clawedfrogs and Bufo species. Direct immersion in a 0.28%
solution of isoflurane was found to provide surgicalanesthesia. The duration and depth of anesthesia

with a direct bath can be prolonged, so caution
should be taken when using isoflurane with thismethod. The best results can be achieved by removing the animal from the solution once it shows signsof sedation (e.g., loss of righting reflex, slowed response to superficial and deep pain). Aerating theisoflurane (5%) into the immersion water was also
found to be effective, but induction and recovery
rates were faster and longer, respectively, than thosedescribed for topical application or a direct bath of
the inhalant. Aerating the anesthetic into the immersion solution is not recommended because of thehealth risk to the humans performing the procedure. Topical application can be achieved by dripping the isoflurane directly onto the animal, saturating a pad with isoflurane and placing the pad on the
animal, or mixing the isoflurane with a water-soluble
carrier and applying it on the animal.1,20 Different
doses are generally required for amphibians basedon their life history. For example, African clawedfrogs require a lower dose (e.g., direct application:
0.007 mL/g body weight; mixed with water-solublegel: 0.025 mL/g body weight) than toads (e.g., direct
application: 0.015 mL/g body weight; mixed withwater-soluble gel: 0.035 mL/g body weight).20 For
larger specimens, isoflurane can be used as it was
intended, as an inhalant. Induction rates for amphibians are generally best achieved at 5%, whereas
maintenance levels generally range between 1% and
3%, depending on the type and duration of theprocedure.

Propofol
Propofol is a nonbarbiturate anesthetic that is routinely used to anesthetize reptiles, birds, and mammals. The primary benefits associated with propofol
are that it is noncumulative, not a perivascular irritant, and is not associated with prolonged recoveries.
The primary disadvantages associated with propofol
include cardiopulmonary depression and route ofdelivery (e.g., in higher vertebrates, it must be given
intravenously or intraosseously). Although propofol
can be used intravenously or intraosseously in amphibians, it has also been evaluated via intracoelomic
injection and immersion.
Intravenous injections of propofol can be given in
the femoral vein, sublingual vessels, heart, ventral
abdominal vein, or ventral tail vein of amphibians.
Caution should be used when injecting compounds
into the heart of an amphibian, because this canprove fatal. Intraosseus injections can be given in thetibia or femur. Intracoelomic delivery of propofol
has been attempted in White’s tree frogs (Pelodryas
caerulea) and tiger salamanders.7,8 In the White’s tree

46
frogs, the propofol was administered at 3 different
doses: 9.5 mg/kg, 30 mg/kg, and 53 mg/kg. These
doses produced moderate anesthesia, surgical anesthesia, and death, respectively. In tiger salamanders,
propofol was dosed at 25 or 35 mg/kg. Surgicalanesthesia was achieved in the majority (83%; 5/6)of the animals given the higher dose (35 mg/kg),
whereas less than half (40%; 2/5) of the animals
given 25 mg/kg achieved surgical anesthesia. Cardiopulmonary depression was observed in bothgroups, with extended periods of apnea. Immersionof African clawed frogs in a propofol solution was
found to have few benefits.4 Propofol doses of 88 and175 mg/L were found to cause sedation and death,respectively. However, at the lower dose (88 mg/L)
the animals were still responsive to noxious stimuli,
suggesting that this route of delivery would haveminimal benefit.

Dissociative Agents
Ketamine hydrochloride (HCl; KetasetŒT, Fort Dodge
Animal Health, Fort Dodge, IA USA) and tiletamineHCl (TelazolŒT, Fort Dodge Animal Health) have bothbeen evaluated in amphibians. Although ketamine
HCl has been found to provide effective and consistentanesthesia in amphibians, tiletamine HCl has beenfound to have limited value.5,21 The primary advantages
associated with using ketamine HCl are that it isinexpensive, can be administered intramuscularly,
and provides good somatic analgesia. Disadvantages
associated with this compound include cardiopulmonary depression and prolonged recoveries. Dosing
recommendations for ketamine hydrochloride
range from 20 to 210 mg/kg.5 It is important to use
caution with this compound at very high doses, because it cannot be reversed. The author generally
recommends ketamine as a preanesthetic, administering doses to animals at 20 to 40 mg/kg intramuscularly. Because dissociatives are not considered toprovide visceral analgesia or muscle relaxation inhigher vertebrates, the author does not recommend
using these compounds as a sole anesthetic for asurgical procedure.

Mitchellfound that the animals achieved no detectable levelof sedation, but cardiopulmonary depression did occur.6 Additional studies evaluating different speciesof amphibians and different dosing regimens shouldbe pursued to determine if these compounds haveany value as anesthetics for amphibians.

Anesthetic Monitoring
Anesthetic monitoring of higher vertebrates has improved significantly over the past decade. We now
have the ability to use point-of-care anesthetic equipment to measure oxygen saturation, heart rate, respiratory rate, electrocardiograms, end-tidal carbon
dioxide, and indirect and direct blood pressures.
Unfortunately, for many of our amphibian patients,
this monitoring equipment is not sensitive enoughto detect their respiratory or circulatory outputs.
Although our ability to monitor amphibians may belimited compared with higher vertebrates, there arecertainly some methods available that can be used tomeasure the physiologic and anesthetic status of ouramphibian patients undergoing an anesthetic procedure.

Cardiac
One of the physiologic parameters that can be consistently measured in our amphibian patients is heartrate, and there are a number of different methods
available for measuring an amphibian’s heart rate.
The author’s preferred method is to use an ultrasonic Doppler. The heart rate can be measured byplacing the Doppler on the ventral pectoral girdle oraxillary region of urodelans or anurans (Fig 4).6,7 Anappropriate contact gel should be used to help facil-

Medetomidine
The alpha-2 agonists are popular anesthetics in veterinary medicine. The primary advantages associatedwith these compounds are that they provide sedation, muscle relaxation, visceral analgesia, and arereversible. They are commonly combined with dissociate agents to provide a more complete anesthesia.
The primary disadvantage associated with these compounds is cardiopulmonary depression. A study evaluating medetomidine (0.15 mg/kg) in leopard frogs

Figure 4. An ultrasonic Doppler can be placed on the ventral
pectoral girdle of an anuran to measure heart rate.

47

Anesthetic Considerations for Amphibians

itate the transmission of the heart beat. Because of
the sensitivity of amphibian skin, it is important toremove the gel after completing the procedure. A
physiologic saline solution (0.9%) can be used toirrigate the area. An electrocardiogram can also be
used to measure the heart rate of an amphibian. A
3-lead system can be used as in higher vertebrates.
Because alligator clips can damage the fragile skin of
an amphibian, the author prefers to place hypodermic needles (25 or 26 gauge) through the skin andto connect the alligator clips to the end of the needles. A pulse oximeter with an appropriate probe
can also be used to measure heart rate in larger
amphibians. Placing a C-clip probe over a peripheral
vessel (e.g., femoral) or inserting a linear probe in
the esophagus or cloaca with the probe directeddorsally toward the aorta can be done to measureheart rate. Caution should be used when interpreting oxygen saturation levels on a pulse oximeter inan amphibian. Pulse oximeters were developed forevaluating mammalian values, but can certainly be
used to determine oxygen saturation trends in an
amphibian.
It is important to obtain a baseline heart rate
before starting an anesthetic procedure. Amphibianheart rates are generally much lower (25-80 beats
per minute) than those found in higher vertebrates.
Knowledge of the baseline heart rate is especially
important when attempting to diagnose abnormalities such as bradycardia. The heart rate of an amphibian is dependent on its environmental temperature. Therefore, the animal should be kept at its
optimal temperature during the preanesthetic, anesthetic, and postanesthetic periods. This will ensurethat the animal can be induced, managed, and recovered from the anesthetic in a consistent matter.
Although measuring the heart rate of an amphibian can be invaluable during an anesthetic procedure, it is important to recognize that the heart of anamphibian can beat after it is clinically dead. Manyof us learned this unfortunate lesson during highschool biology. Because of this, it is important torecognize that heart rate is but one method we canuse to monitor anesthesia in an amphibian, and thatmeasuring heart rate should not be the only monitoring method used.

thought to be done to assist with olfaction. In anurans, it was found to be consistent with respirations.6 In some species, such as tiger salamanders,gular pumping did not occur for extended periodsof anesthesia with clove oil and propofol.7 Fortunately, amphibians have developed a highly vascular
integument that can assist with respirations. Although there is comfort in knowing that these animals can respire through their skin, it does tend tolead to difficulties when trying to determine whether
an animal is apneic or not.

Reflexes
There are a number of reflexes that can be used toassess the anesthetic depth of an amphibian patient.
The methods most commonly used by the author
include the escape response, righting reflex, superficial and deep pain responses, and the palpebral
reflex. It is important to characterize these reflexes
in the patient before initiating an anesthetic procedure, because animals may show different responses
to each reflex. A rank scale can be used to assess thedifferent levels of a reflex. The author prefers torecord the reflexes as 1 ⫽ present, 2 ⫽ delayed, or
3 ⫽ absent. Surgical anesthesia is generally determined to be when the reflexes are all recorded as
being absent.6,7The escape reflex can be assessed by placing theanimal in an open palm and determining whether ittries to walk or jump away. Most amphibians have anatural desire to “escape” a perceived predator. Animals that are naturally stoic or long-term captives
may not display an active escape response. The righting reflex can be measured by placing the animal on
its dorsum and counting the time required to correct

Respiratory
In higher vertebrates, respirations can be visuallycharacterized by counting thoracic excursions. In
some amphibians this is possible, whereas in others it
is not. One of the methods that the author has used
to measure respirations is gular movements. In somespecies, such as chelonians, gular pumping is

Figure 5. The righting reflex can be assessed by placing the animal
in dorsal recumbency and measuring the time it takes for the animal
to recover. This anesthetized leopard frog had no righting reflex.

48

Mitchellthetic protocols for these animals. Although amphibian anesthesia has improved over the past decade,there is certainly much more that can be done. In
addition to finding new and consistent methods foranesthetizing amphibians, it is also important to develop better methods for monitoring amphibians tominimize the likelihood of complications.

References
1.

Figure 6. Superficial and deep pain can be assessed by pinching
the skin or digit of an amphibian.

2.

3.

its position (Fig 5). Animals that are sedate generally
have a delayed response, whereas those that are at asurgical plane of anesthesia will have no righting
reflex. The palpebral reflex can be assessed bytouching the upper or lower palpebrae with a cotton-tipped applicator. Animals that are alert will
blink on response to this stimulation, whereas animals at a surgical plane of anesthesia will not respond. Superficial pain can be assessed by pinching
the skin of an amphibian. The author prefers to use
the dorsal or ventral surface of the rear feet (Fig 6).
Deep pain can be assessed by grasping a digit. The
superficial pain reflex is lost before the deep painreflex; however, the deep pain reflex returns first.

4.
5.

6.

7.

8.

Postanesthetic Plan
9.

Amphibians should be recovered from an anesthetic
event with the same standard of care used for othervertebrates. The recovery area should be quiet andthe environmental temperature and humidity appropriate for that species. Lining the recovery enclosure
with paper towels or towels that are premoistenedwith dechlorinated water will help prevent dessication. Amphibians can be recovered on room air.
Return of the reflexes is important for determining
when an animal is fully recovered. Replacing an amphibian into an aquatic environment before being fully
recovered could result in an accidental drowning.

Conclusions
As amphibians are presented to veterinarians forvarious diagnostic and surgical procedures, it will be
important to develop complete and consistent anes-

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and benzocaine for metamorphic and paedomorphic tiger salamanders (Ambystoma tigrinum nebulosum). Am Midl Natural 155(2):417-422, 2006
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Anesthetic Considerations for Amphibians
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Sladky KK, Swanson CR, Stoskopf MK: Comparative
efficacy of tricaine methanesulfonate and clove oilfor use as anesthetics in red pacu (Piaractus brachypomus). Am J Vet Res 62(3):337-342, 2001
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