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Reptile Euthanasia
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Old 12-04-2013, 06:43 PM   #1
Reptile Euthanasia

CH-3003 Berne-Switzerland, February 2013
Federal Department of Home Affairs FDHA
Federal Veterinary Office FVO
International Affairs
Analysis on humane killing methods for reptiles in the skin trade
1 Introduction 2
2 Criteria for the evaluation of a humane killing method 3
3 Killing methods for reptiles: current knowledge 3
3.1.1 Decapitation 3
3.1.2 Cervical Dislocation 3
3.1.3 Shooting: Free Bullet, Gunshot 4
3.1.4 Captive-Bolt Pistol 4
3.1.5 Stunning: Blow, Concussion 4
3.1.6 Exsanguination 4
3.1.7 Rapid Freezing, Supercooling, Cooling, Hypothermia 5
3.1.8 Heating, Hyperthermia 5
3.1.9 Suffocation 5
3.1.10 Drowning 5
3.1.11 Pithing 5
3.2 Chemical Methods 5
3.2.1 Inhalation 5
3.2.2 Injection 6
4.1 4.2 5 6
Conclusions 6 Animal Welfare Considerations 6 Recommendations 6
Expert Panel 8 References 9
Mechanical Methods 3
Editor: Swiss Federal Veterinary Office (FVO), 2013
Expert Panel (2013): Analysis of humane killing methods for reptiles in the skin trade, ed. Swiss Federal Veterinary Office (FVO)
Analysis on humane killing methods for reptiles in the skin trade
1 Introduction
Trade in reptile skins, their parts and derivatives provides an important economic activity for a diverse range of stakeholders along the value chain from the hunters/breeders to skin traders, tanners, leather refineries to tailors and commercial luxury brand houses and finally consumers. Details of this trade, in particular relating to trade in Python snakes, can be found in the recently published report by the In- ternational Trade Center16. Exact trade figures for all reptile species are difficult to obtain, in particular for species for which no international trade controls exist. However, thanks to the trade data gathered by the Convention on International Trade in Endangered Species of wild Fauna and Flora (CITES), the volume of trade in those species listed in the Appendices to CITES can be assessed over the past 40 years. According to figures on the dashboard of CITES’ official website, this volume has seen very high figures of between 1.5 and 2.25 millions skins per year in the past, but seems to be decreasing since 2006 to a level of around 800’000 skins traded per year. However, taking into account those species not listed by CITES and for which no trade data exist, the number of skins traded per year is likely to be considerably higher.
Increasingly in recent years, concerns have been raised about the potential welfare implications for snakes and lizards caught and killed for the skin trade. To improve the welfare of the animals involved in the trade and to reduce their suffering caused by inhumane slaughtering methods, it was felt that guidelines on how to humanely euthanize reptiles are needed. This need was also recognized by vari- ous stakeholders along the value chain starting from the countries of export, to the countries of import, international organizations and NGO’s but, in particular, by the public and the companies which sell the finished products to the final consumer at the end of the value chain.
In 2011, Switzerland as an important hub in the trade of finished products obtained from reptile skins, directed it’s Federal Veterinary Office to set up an Expert Panel composed of internationally recog- nized experts in the field of animal welfare, euthanasia and trade in reptile skins with the aim to exam- ine the current literature and draft recommendations for the humane killing of reptiles. The list of the members of this Expert Panel can be found in the paragraph 5 of this document. In particular, the Ex- pert Panel was tasked with assessing the humaneness, appropriateness and acceptability of the vari- ous existing euthanasia methods available for reptiles irrespective of the setting under which the killing takes place.
The Panel set about the task in three phases. The first phase was to review published scientific and gray literature on current methods used to euthanize, slaughter and kill reptiles. The second phase was to rate the list of published information according to the usefulness and relevance for the purpose of the panel’s work. In the third and final phase the selected publications were used to gain consensus amongst the panel and to create a document, which represents the panel’s recommendations for hu- mane slaughtering of reptiles. These recommendations are therefore based on a thorough review of the current level of available knowledge. The analysis of the expert panel also revealed that there are gaps in our knowledge about various aspects that may influence the humaneness of the methods of euthanasia recommended in this document. Should new information become available the recom- mendations of this panel may have to be reviewed and updated accordingly.
The Panel stresses that these recommendations are only a first step towards assuring the humane treatment of reptiles in the production of leather for this trade. We hope that these recommendations will be put to work, closely monitored and research conducted to test their humaneness and effective- ness.
Switzerland will start by submitting these recommendations to the World Organization for Animal Health (OIE) for adoption as standards for humane slaughtering of reptiles and sharing them with the relevant working groups dealing with animal welfare issues in the fashion industry. The Panel also recommends the wider incorporation of humane killing methods into Best Management Practices (BMP’s), and assessment of compliance through wildlife management authorities as well as capacity
Analysis on humane killing methods for reptiles in the skin trade
building activities in the countries of origin of the species involved in the trade.
2 Criteria for the evaluation of a humane killing method
The methods used should1:
- Attempt to cause no pain
- Lead to rapid unconsciousness and death
- Be performed with minimum restraint
- Avoid excitement
- Be appropriate for the age, species, and health of the animal
- Attempt to cause no fear and psychological stress to the animal
- Be reliable, reproducible, irreversible
- Be simple to administer
- Be safe for the operator
- Be aesthetically acceptable for the operator or observer
Considerations about the practicability of a method2:
- Compatibility with requirement and purpose
- Drug availability and human abuse potential
- Ability to maintain equipment in proper working order
- Economical3
3 Killing methods for reptiles: current knowledge
3.1 Mechanical Methods
3.1.1 Decapitation
This procedure involves the severing of the neck of the animal, exactly between the skull and the first cervical vertebra, using a sharp instrument (guillotine1,2,3,4,5, axe or blade1) ideally with a single very swift cut4 that leads to severance of the spinal cord.
Some reptiles may remain conscious for up to an hour after decapitation4,6,7, which makes this proce- dure acceptable only if the brain of the severed head is immediately destroyed by pithing2,4,6,8 or by blunt trauma.
3.1.2 Cervical Dislocation
This method involves separation of the skull and the brain from the spinal cord by applying pressure3 in a simultaneous ventral-cranial motion at the base of the skull with an appropriate tool.
The operator must be confident of performing this technique quickly and effectively. It requires master- ing of technical skills to ensure that loss of consciousness is rapidly induced3. Although suitable for small rodents, rabbits and birds, the method is not appropriate for larger reptiles (> 200 grams) owing to the resistance of the reptilian brain to hypoxia4. Also, for taxon specific anatomical reasons, and es- pecially in large specimens, it is extremely difficult to dislocate vertebrae.
3.1.3 Shooting: Free Bullet, Gunshot
Shooting should be performed by personnel trained in the use of firearms.
A high level of skill is required in order to hit the brain through the two brain cases found in many rep- tiles9. In addition, with small species, and/or where the target is moving, shooting may not be effective. However, apart from this, shooting in the head to ensure immediate destruction of the brain is an ef- fective and humane way of killing large reptiles1. It is occasionally recommended that even when this method is used the spine is severed and the brain destroyed by pithing 10,11.
Analysis on humane killing methods for reptiles in the skin trade
3.1.4 Captive-Bolt Pistol
Captive bolt pistols are powered by gunpowder or compressed air and must provide sufficient energy to penetrate the skull2 (penetrating captive bolt) or cause fatal stunning (non-penetrating captive bolt) of the species on which they are being used.
The animal must be properly restrained to ensure that only a single shot is required. Both penetrating and non-penetrating captive bolt guns must be placed directly on the skull over the brain cavity to en- sure their effectiveness. All personnel must be trained in these techniques to ensure the correct posi- tioning of the weapon to ensure a direct hit into the brain1. An appropriate charge of the gun (air or gunpowder) must be selected to match the size of the animals. It has been shown to be very efficient for the slaughter of Pantanal caimans and American alligators, and can be used for all sizes of croco- dilian12,13 given that appropriate charge is selected. It is considered an acceptable and humane meth- od for large reptiles but should only be carried out by trained personnel who know where to position the pistol5 and thereby ensure a direct hit into the brain. In snakes the captive bolt would have to be shortened to avoid wrist injuries and damage to the equipment. However, there are ways of using the standard equipment in snakes by placing soft materials (foam, etc.) beneath the animal to soften the trajectory of the bolt after penetrating the head. Alternatively the non-penetrating bolt would not need modification although the head is quite large for some species.
3.1.5 Stunning: Blow, Concussion
This involves striking the head of the animal directly over the cranium with some hard implement or object and with sufficient force to cause immediate loss of consciousness and/or death9,14.
If many animals are to be killed within a short time by the same operator it is difficult to ensure con- sistency in performance and therefore only a few animals should be killed by the same person using this method at any time1. Larger reptiles (crocodilians) may be rendered unconscious by this method but are less likely to be killed. The brain must be destroyed before the return of consciousness3,4,9,14, either by a further blow or by some other method such as pithing. It is considered an acceptable method for all reptiles but should only be carried out by experienced operators6 who know exactly where to strike.
3.1.6 Exsanguination
By cutting the major blood vessels in the neck i.e. the carotid arteries and jugular veins3.
This method of euthanasia is not acceptable for reptiles and other ectothermic vertebrates because of their slow metabolic rate and hypoxic tolerance1,3,6.
3.1.7 Rapid Freezing, Supercooling, Cooling, Hypothermia
Killing of animals by placing them in very cold temperatures such as deep freezers.
Immobilization of reptiles by cooling is considered inappropriate and inhumane even if combined with other physical or chemical methods of euthanasia2. Quick freezing of deeply anesthetized animals is acceptable2. In the laboratory situation dropping an animal into liquid nitrogen at minus 196°C – a very extreme form of freezing, far removed from a domestic freezer may be acceptable for animals of less than 40g bodyweight (i.e. less than 1cm in diameter) as liquid nitrogen would freeze an entire body of that size instantaneously4. This methods as a standalone method is not acceptable for euthanasia of reptiles1,9,15
3.1.8 Heating, Hyperthermia
Raising the temperature above the critical temperature of the species4. This method is not acceptable for euthanasia of reptiles1,9,12.
3.1.9 Suffocation16
Analysis on humane killing methods for reptiles in the skin trade
Depriving animals of oxygen. This method is not acceptable for euthanasia of reptiles.
3.1.10 Drowning
This method is not acceptable for euthanasia of reptiles4.
3.1.11 Pithing
Carried out by inserting a sharp metal rod or probe through the foramen magnum into the base of the brain to ensure quick brain destruction3.
Method acceptable for unconscious reptiles1,2,9 (e.g stunned, anaesthetized). It may also be accepta- ble when performed immediately after decapitation or cervical dislocation. Pithing can be carried out in reptiles without crushing the skull4.
3.2 Chemical Methods
3.2.1 Inhalation (Halothane, Enflurane, Isoflurane, Methoxyflurane, Ether, CO2, CO)
With inhalant anaesthetics, the animal can be placed in a closed receptacle containing cotton or gauze soaked with an appropriate amount of the anesthetic, or the anesthetic can be introduced from a va- porizer. The latter method may be associated with a longer induction time. Vapors are inhaled until respiration ceases and death ensues2.
Many reptiles are capable of holding their breath and converting to anaerobic metabolism, and can
survive long periods of anoxia (up to 27 hours for some species)2,4,6. Because of this ability to tolerate
anoxia, induction of anaesthesia and time to loss of consciousness may be greatly prolonged when
inhalants are used2,17. Death in these species may not occur even after prolonged inhalant exposure2.
Therefore euthanasia by inhalation of toxic gases is not an acceptable method for Euthanasia in rep- tiles1,3,4,18.
3.2.2 Injection (Barbiturate, Tricaine methane sulfonate, T-61, Others)
Various methods of applications are available (e.g. intravenous, intraperitoneal, intrapulmonic, intra- muscular, subcutaneous, intracardiac, oral, rectal).
Sodium pentobarbitone is an effective and humane method of euthanasia in reptiles3,4,9,12. The intra- venous route can be used by well-trained personnel9 and result in quicker death1. Where intravenous injection is difficult the intraperitoneal route may be used but it is slower acting3,4,9. Intracardiac injec- tion may only be used on a fully anaesthetized animal as this is very painful and is therefore not con- sidered acceptable3,9. Intramuscular or subcutaneous should not be used as they are not effective and may cause pain4.
Tricaine methane sulfonate (TMS, MS-222) may be administered by various routes to euthanatize. These are expensive means of euthanasia2 and because there is little information on the humaneness of this method, it is not considered acceptable for reptiles9.
T-61 must only be injected intravenously and slowly as it is otherwise painful1. The animal must be se- dated prior to administration of T-611 and is acceptable in all groups of animals15.
Some authors recommend the intramuscular injection of ketamine as a premedication minutes prior to intravenous injection of sodium pentobarbitone, but ketamine should never be used as a sole agent for euthanasia2.
Analysis on humane killing methods for reptiles in the skin trade
4 Conclusions
4.1. Animal Welfare Considerations
We exclude the following methods (alone or in combination with other methods) as being inappropri- ate and inhumane:
- Exsanguination
- Freezing
- Heating
- Suffocation - Drowning
- Inhalation
4.2. Recommendations
Key issue in the appreciation of humane methods of Euthanasia for reptiles is that the brain has to be destroyed by either chemical or mechanical methods.
Based on the current knowledge, we consider the following methods as acceptable at this time:
Captive-bolt pistol
Alone or with a subsequent method to ensure death (pithing) if the brain is not immediately destroyed.
Blow to the head with a hard imple- ment
In combination with a subsequent method to ensure death (pithing) if the ani- mal is only stunned.
With a subsequent method to ensure death (pithing or blunt trauma).
With appropriate bullet for size of the animal and in line with relevant legisla- tion, training and safety protocols (effective, quick and humane). Particularly in conjunction with spinal severance and pithing. Minimizing the distance be- tween the animal and the shooter will reduce margin for error for “missing” the brain.
After prior stunning (captive-bolt or blow) or decapitation and as method to ensure death.
Cervical Dislocation, if performed in the correct size animal (<200g)
With proper technique and followed by another procedure to ensure death.
Depending on the context and the experience/training of the person (e.g. vet- erinarian, researchers).
Analysis on humane killing methods for reptiles in the skin trade
The selection of killing methods for reptiles depends on the expertise, proper administration, trained personnel and setting for humane euthanasia. A heavy blow to the head and the use of the captive bolt pistol are simple and fast to perform. Cervical dislocation requires a proper technique and can not be easily carried out, even in small reptiles. Injection could only be considered practicable in facili- ties where the necessary equipment, man-power and capacities are available (laboratories, zoos etc.).
Analysis on humane killing methods for reptiles in the skin trade
5 Expert Panel
Composition of the expert Panel on humane euthanasia of reptiles:
- Ashley Don (Alligator and Crocodilian Trade Consultant; International Alligator/Crocodile Trade Study (IACTS), IUCN Crocodile Specialist Group (CSG), Industry Committee, Louisiana Dept. of Wildlife & Fisheries and Alligator Advisory Council).
- Auliya Mark (PhD, Dept. Naturschutzforschung - Dept. of Conservation Biology).
- Briner Alexandra (, Scientific Officer for Food Hygiene, Veterinary Office, Switzer-
- Cooper Margaret (MARGARET E COOPER, LLB, FLS, Solicitor (not in private practice), Vis- iting Lecturer, Faculty of Veterinary Medicine, University of Nairobi, Kenya. Honorary Re- search Fellow, DICE, The University of Kent, UK).
- Cooper John (JOHN E COOPER, DTVM, FRCPath, FSB, CBiol, FRCVS Diplomate, Europe- an College of Veterinary Pathologists. European Veterinary Specialist, Zoological Medicine. Visiting Professor, Faculty of Veterinary Medicine, University of Nairobi, Kenya).
- Dublin Holly (PhD., Director and Special Adviser, sustainability at PPR to 31/12/2012. IUCN ESARO, Senior Adviser and Chair, IUCN/SSC African Elephant Specialist Group, P.O. Box 68200, Nairobi, KENYA, 00200, currently).
- Loup Fabien (Substitute Chief Animal Welfare, Veterinary Office, Switzerland).
- Manolis Charlie (Chief Scientist, Wildlife Management International).
- Martelli Paolo (Chief Veterinarian, Ocean Park Hong Kong).
- Micucci Patricio Alejandro (Biologist at the University of Buenos Aires. Member of the Croc- odile Specialist Group of the World Conservation Union (IUCN), Technical Director-Yellow Anaconda Conservation and Management Plan, Formosa-Argentina).
- Morgan Guy (BSR, Manager, Advisory Serices).
- Nevarez Javier (DVM, PhD, Dipl ACZM, Dip ECZM (Herpetology), Assistant Professor of Zoological Medicine, Wildlife Hospital of Louisiana Director, LSU School of Veterinary Medi- cine
Veterinary Clinical Science, Skip Bertman Dr., Baton Rouge, LA 70803).
- Karesh William (D.V.M, Executive Vice President for Health and Policy, President, OIE Work- ing Group on Wildlife Diseases, Co-chair, IUCN Species Survival Commission - Wildlife Health Specialist Group, Technical Director - Emerging Pandemic Threats - PREDICT program).
- Kelly Andrew (Head of Wildlife Department, Royal Society for the Prevention of Cruelty to Animals (RSPCA) to 31/12/2012, BSc(Hons) Zoology, PhD Ecology and Evolution, Visiting Researcher, Centre for Ecology and Conservation, University of Exeter, Cornwall Campus).
- Waller Tomas (Chair IUCN/SSC boa & Python Specialist Group).
- Wenger Sandra (; MSc. Dipl. ECVAA).
Analysis on humane killing methods for reptiles in the skin trade
6 References
1 Close B, Banister K, Baumans V, Bernoth EM, Bromage N, Bunyan J, Erhardt W, Flecknell P, Grego- ry N, Hackbarth H, Morton D, Warwick C. Recommendations for euthanasia of experimental animals: Part 1. Laboratory Animals. 30:293-316. 1996.
2 AVMA Guidelines on Euthanasia (Formerly Report of the AVMA Panel on Euthanasia). 2007.
3 Sharp T, Saunerds G, NSW Department of primary Industries. GEN001 methods of euthanasia. General. 2004.
4 Baines FM, Davies RR. The Euthanasia of Reptiles. 2010.
5 Euthanasia. The University of Western Ontario Animal Care and Veterinary Services. Standard Op- erating Procedure.
6 Mader DR. Reptile Medicine and Surgery. 2nd edition. 2006.
7 Warwick C. Letter to the editor: Euthanasia of Reptiles-Decapitation: An Inhumane Method of Slaughter for the Class “Reptilia”. Can Vet J. 27:34. 1986.
8 Cooper JE, Ewbank R, Platt C, Wawrick C. Euthanasia of tortoises. Vet Rec. 115(24):635. Dec 1984.
9 Close B, Banister K, Baumans V, Bernoth EM, Bromage N, Bunyan J, Erhardt W, Flecknell P, Grego- ry N, Hackbarth H, Morton D, Warwick C. Recommendations for euthanasia of experimental animals: Part 2. Laboratory Animals. 31:1-32. 1997.
10 Hutton JM, CSG Vice Chairman for Africa, PO Box HG 690, Highlands, Harare, Zimbabwe. Humane killing of crocodilians.
11 Natural Resource Management Ministerial Council. Code of practice in the humane treatment of wild and farmed Australian crocodiles. 2009.
12 Aleixo VM. Use of captive bolt pistol for human slaughter of crocodilians. Crocodile Specialist News- letter 27(4):24-25. 2008.
13 Campos Z. The “Zilca”, a new device for the humane killing of crocodilians. Crocodile Specialist Group Newsletter 19(1):20. 2000.
14 Code Of Practice for the Humane Killing of Animals under Schedule 1 to the Animals (Scientific Procedures) Act 1986.
15 Zwart P, de Vries HR, Cooper JE. The humane killing of fishes, amphibia, reptiles and birds. Tijdschr Diergeneeskd. 114(10):557-565.
16 Kasterine A, Arbeid R, Caillabet O, Natusch D. The Trade in South-East Asian Python Skins. Inter- national Trade Center (ITC), Geneva. 2012.
17 Girling SJ and Raiti O. BSAVA Manual of Reptiles. 2nd edition. 2004.
18 Kaplan M. Decapitation of Reptiles. Inhuman for euthanasia. Melissa Kaplan’s Herp Care Collection. 1997.
Analysis on humane killing methods for reptiles in the skin trade
Old 07-23-2014, 07:58 AM   #2
Evaluation of Hypothermia for Anesthesia in Reptiles and Amphibians

Brent J. Martin
Brent J. Martin, D.V.M., is the Director, Department of Animal Resources, Vivarium, University of California, Santa Barbara, California.


When reviewing research proposals involving ectothermic animals, members of institutional animal care and use committees (IACUCs) are frequently required to evaluate husbandry methods and research techniques that are not well known to them. Gathering additional information on these techniques is often unrewarding due to the paucity of published accounts and the tremendous diversity among ectotherms.

One such topic of concern is the use of hypothermia for anesthesia. This anesthetic method has been used for over a century (Blair 1971) and is still often proposed by investigators who use amphibians and reptiles. IACUC members frequently express discomfort with the adequacy of this method; they may be unfamiliar with and lack specific knowledge about hypothermia and may suspect that it is insufficient to render an animal insensible to painful procedures such as surgery. Investigators may base their arguments for its use on past successful experiences where the hypothermic animal failed to respond to any stimuli. Investigators may also have experienced better survival rates with hypothermia than with traditional anesthetics. The scientific literature is replete with references to hypothermia used for anesthesia, and there are some concerns that anesthetics can adversely affect an experiment (Smith and others 1991). Also in the literature are counter-balancing statements that regard cold anesthesia as malpractice.1

The following review is an effort to present information that is known in the area of hypothermia in order to assist IACUC members in evaluating its use. Currently there is insufficient information to determine authoritatively when and if hypothermia is appropriate; however, there are some basic studies that are useful when reviewing the issue.

Research and Regulatory Guidelines

The regulations and guidelines for animal research are not helpful regarding the appropriateness of the use of hypothermic anesthesia for ectotherms. The guidelines for IACUC members that are currently available are not well referenced and are contradictory. Neither the Guide for the Care and Use of Laboratory Animals (Guide) (NRC 1996) nor the standards of the Animal Welfare Act address this issue. The report of the American Veterinary Medical Association (AVMA) Panel on Euthanasia (AVMA 1993) and the Institute of Laboratory Animal Resources (ILAR) report, Recognition and Alleviation of Pain and Distress in Laboratory Animals (NRC 1992, p 115), indicate that hypothermia is unacceptable for euthanasia. Both of these documents assert that while hypothermia decreases metabolism no evidence exists that it raises the pain threshold. The document cited in both cases states ‘there is no evidence that it [hypothermia] raises the pain threshold, i.e., makes the animal less susceptible to painful stimuli’ (Cooper and others 1989, p 11) but does not reference the statement. The listed references of Cooper and others (1989) consist entirely of review articles, books, and correspondence. Paradoxically, Recognition and Alleviation of Pain and Distress in Laboratory Animals further indicates that hypothermia is an effective analgesic for altricial neonates that have not acquired effective ther-moregulation and as an ‘adjunct to general anesthesia in cold-blooded animals’ (NRC 1992, p 83). The Canadian Council on Animal Care (CCAC) Guide to the Care and Use of Experimental Animals acknowledges that hypothermic amphibians appear to be in an unconscious state, but limits the use of hypothermia to nonpainful procedures because unconsciousness has not been assured (CCAC 1980, p 62-69). No references are offered. A newsletter article on pain relief in ectothermic animals indicates that ‘hypothermia is not an analgesic, since nerve conduction and thus the pain response is not abolished by temperature’ (Arena 1990). The statement is unreferenced; however, there is an article listed in the bibliography that relates to temperature effects on nerve conduction in a reptile. This article is an abstract that has been misinterpreted by Arena; both the abstract and the subsequently published article indicate a clear linear relationship between nerve conduction velocity and temperature (Rosenberg 1977, 1978). Decreasing temperature causes decreased nerve conduction velocities. Further, nerve conduction blockage was recorded at temperatures of 1-3.5°C in tortoises (Rosenberg 1978).

The Scientific Literature

A variety of scientific articles are available that shed some light on the interaction between temperature and nerve function.

The literature characterizes the anesthetic aspects of hypothermia in mammals very well. Cold is anesthetic in mammals; evidence for this is readily available (deJong and others 1966; Rossi and Britt 1984; Antognini 1993; Guerit 1994). Anecdotally, local anesthesia in humans by hypothermia is an exceptionally common procedure, for example, it is standard practice for ear piercing. Further, the human anesthetic literature has a wide variety of references to hypothermia. Specifically, peripheral nerve conduction velocity decreases linearly with decreasing temperature, down to 23.5°C in humans anesthetized with halothane (deJong and others 1966). Linear regression analysis predicted that complete conduction blockade would occur at approximately 9°C (deJong and others 1966). These findings agree with studies in a variety of mammalian species. The general mammalian condition is for hypothermia to induce a neuromuscular and cognitive condition comparable to surgical anesthesia at warmer body temperatures (Blair 1971). The evoked potentials of human and cat central nervous systems cannot be elicited below 20°C (Rossi and Britt 1984; Guerit 1994). A recent study has shown that anesthetized goats cooled to about 20°C do not react to painful peripheral stimuli when the anesthetic is removed (Antognini 1993).

While the data in mammals clearly indicates the neuronal transmission blocking effects of hypothermia, the depressed and blocked function returns with rewarming (Blair 1971; Rossi and Britt 1984; Antognini 1993; Guerit 1994). Therefore, hypothermia would not be expected to supply analgesia once the cooled neurons warmed to functioning level. This would place hypothermia in the class of anesthetics that are not ‘analgesic,’ along with such drugs as pentobarbital and halothane. Hypothermia in mammals also has the potential to cause pain because cold temperature can be a noxious stimulus.

While the mammalian experience with hypothermia is useful for establishing anesthetic concepts, the specific details of the effects of hypothermia on reptiles and amphibians must be sought in literature involving ectothermic animals. Some aspects of hypothermia in ectotherms have been described. As is seen in mammals, peripheral nerve conduction velocities decrease in reptiles (Rosenberg 1978) and amphibians (Hutchinson and others 1970) with decreasing temperature. Hypothermia also reduces electroretinographic responses in amphibians (Schaefer and others 1978). The mechanism may be related to a similar reduction in ionic currents in Xenopus neuronal membranes (Frankenhaeuser and Moore 1963). The response of brain-stem auditory evoked potentials to hypothermia has been studied in alligators (Strain and others 1987). While the amplitude of the potentials decreased with decreasing temperature, potentials were recorded in the alligators at stabilized cloacal temperatures of 0.4°C. The authors further comment that ‘the presence of periodic head and limb movements’ was used to assure subject viability at ‘temperature extremes’ (Strain and others 1987). As is expected for these ectotherms, the temperature range in which neurons function is much cooler than is seen in mammals. Studies in hypothermic ectotherms show neuronal function below the temperature in which conduction blockade occurs in mammals (Hunsaker and Lansing 1962; Parsons and Huggins 1965; Walker and Berger 1973; Rosenberg 1977, 1978; Schaefer and others 1978; Strain and others 1987). While it has been shown that hypo-thermia decreases neuronal function and that ‘numbing’ can be anticipated, the critical point at which anesthesia occurs peripherally is when conduction blockade occurs. In a study of tortoises, peripheral neuron transmission generally blocked at 3.5°C but sometimes blocked as low as 1.2°C (Rosenberg 1977). Prior acclimatization of the tortoises to ambient temperatures that were warmer or cooler than room temperature had no apparent effect on the subsequent blocking temperature (Rosenberg 1977, 1978). A study in bullfrogs reported that in vitro, peripheral nerve conduction was blocked at temperatures of 0-2°C (Roberts and Blackburn 1975). Furthermore, in contrast to the findings in mammalian nerves, it has been reported that in bullfrog nerves, pain-carrying C fibers are blocked at higher temperatures than are the neuromuscular A fibers (Roberts and Blackburn 1975). These findings suggest there is a level of hypothermia that would block transmission of noxious stimuli. However, the temperature at which conduction blockade has been recorded is not substantially different from the ultimate body temperature that would be predicted by standard hypothermia-inducing technique (Smith and Varnold 1991). Ice water induction of hypothermia would leave a small margin for error between conduction blockade and frozen tissue. There is further reason for concern because in both endotherms and ectotherms, detectable muscular movements are blocked at warmer temperatures than the temperatures that block neuronal transmission (Hunsaker and Lansing 1962; deJong and others 1966; Rosenberg 1978). These findings suggest that techniques for monitoring the adequacy of anesthesia, such as testing for withdrawal reflexes, would not be useful for monitoring hypothermia as paralysis will occur before local anesthesia.

The critical question whether hypothermia is able to render ectotherms unconscious prior to inducing peripheral neuromuscular blockade has neither been established nor adequately addressed in the literature. Some experimental evidence exists that suggests that hypothermia does not readily induce unconsciousness in these animals. Walker and Berger (1973) described 2 electroencephalographic states of tortoises: (1) behavioral activity and arousal, and (2) behavioral inactivity. EEG findings for both of these states were distinctly different. Cooling the tortoises, while causing behavioral inactivity, caused the EEG to record a state of arousal. EEG findings in the caiman have shown increased EEG activity in some brain regions at 2-4°C over those seen at room temperature (Parsons and Huggins 1965). Hunsaker and Lansing (1962) reported the EEGs of lizards at various body temperatures. Below 2°C, the EEGs were at isoelectric baseline and at 2-3°C, potentials were inconsistently seen. However lizards routinely had EEG responses to blowing air, flashing lights, and adjacent hand clapping at body temperatures of 2-3°C. One lizard was reported to have EEG responses to air puffs at a body temperature of 0.8°C. In this study, heart potentials were not consistently recorded until 5°C, and detectable body movements were generally not seen until body temperatures reached 9-10°C. It is further noteworthy that at room temperature, the use of curare lead to a marked, generalized reduction in EEG amplitude and an EEG response to air puffs remarkably similar to that seen in hypo-thermic conditions (Hunsaker and Lansing 1962). Although EEG responses to stimuli are not equivalent to consciousness, these findings in hypothermia are clearly different from those seen in mammals where isoelectric and unresponsive EEGs are found at much higher temperatures than are needed to block peripheral nerve function (Blair 1971). Since animals must be conscious to function in their environment and ectothermic animals routinely function at lower temperatures, the findings are consistent with an interpretation that hypothermia does not readily induce unconsciousness in reptiles. There does not seem to be EEG data for hypothermic amphibians, and the reptilian studies may not be indicative of amphibian responses to cold. However, since reptiles are known to use behavioral modification to maintain their body temperature above ambient temperature, there is no reason to suggest that reptiles would display better neural function during extreme hypothermia than amphibians who normally function at those lower ambient temperatures.

Hypothermia has been reported to cause brain necrosis in snakes and turtles (Northcutt and Butler 1974; Wang and others 1977). This may be the cause of clinical wasting that has been anecdotally reported to occur irregularly following hypothermia (Johnson 1992). Clinically apparent adverse effects have not been reported following hypothermia in amphibians. Female Xenopus spp. do very well following hypothermia even after repeated episodes including abdominal surgery (personal observation). A wide variety of alternative anesthetic regimes are available for amphibians and reptiles that do not use hypothermia (Kaplan 1969; Northcutt and Butler 1974; Wang and others 1977; CCAC 1980; Cooper and others 1989; Schaeffer 1994; Stoskopf 1994). Hypothermia may be a useful adjunct to the use of chemical anesthetic agents. The anesthetic requirement in both mammals (Eger and Johnson 1987; Antognini 1993) and fish (Cherkin and Catchpool 1964) has been shown to be dramatically less when hypothermia is induced. Studies have found that amphibians have a potent opioid in their skin (Braga and others 1984), which may have some influence on their recovery from surgical procedures. While it has been suggested to house ectothermic animals below their preferred temperature a day or two before using anesthetics (Arena 1990), this strategy could make amphibians less tolerant to pain and less susceptible to the analgesic effects of administered opioids (Stevens and Pezalla 1989).

MS-222 is often considered the anesthetic of choice in amphibians (CCAC 1980; Cooper and others 1989; Johnson 1992; Schaeffer 1994). An excellent review of its pharmacology and use in amphibians has recently been published (Downes 1995). However, a recent standard research methodology text advocates hypothermia, indicating that MS-222 induces the maturation of Xenopus oocytes (Smith and others 1991). While anesthetic interference with research can be a compelling argument against its use, it is unclear from the literature whether MS-222 actually adversely affects oo-cytes. Justification for Smith's statement is not supported following close examination of the citations. Smith initially cites himself in which he reproduces, in part, a table by Baulieu and others (1978). The Baulieu article reports that lidocaine, dibucaine, and tetracaine induce oocyte meiosis and that benzocaine and procaine do not. MS-222 is not reported. Further, the citations for tetracaine are articles dealing with calcium-membrane interactions. None of the references involve oocyte membranes nor do they model clinical application of tetracaine for frog anesthesia. Table 2 in Baulieu and others (1978) is an interpretation of drugs that could induce meiosis based on their interaction with calcium. The actual effects of the drugs on Xenopus oocyte maturation are not reported. While MS-222 may effect Xenopus oocytes, this has not been reported in the literature and its use should not be avoided based on these citations.

Amphibians and reptiles include thousands of very diverse species. Although the available articles related to the subject are inadequate for such a large and diverse group, they generally do not support hypothermia as a clinically efficacious method of anesthesia.


The author would like to acknowledge the community on COMPMED for their assistance and comments. Dr. Mark Suckow contributed valuable editorial assistance.


↵1 Helene N. Guttman reported in “The care and use of amphibians, reptiles and fish in research” (SCAW Newsletter 13[3]:11) that J Flanagan and V Lance both emphasized during a conference sponsored by the Scientists Center for Animal Welfare (held April 8-9, 1991, in New Orleans, Louisiana) that using hypothermia is malpractice.

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