Anesthesia can be critically unique based upon the type of animal involved. Understanding the basic anatomy and physiology of reptiles and amphibians is key for selection and assessment of anesthesia. Less than 30 original researches, peer-reviewed studies have been published regarding specific reptilian species and the effects of different anesthetic agents. The practitioner is often left extrapolating these results to clinical cases. Many times, the patient requiring anesthesia is not a healthy animal, and most published scientific studies are on animals that are healthy, thus confounding the situation. Ambient temperature is critical for evaluating drug metabolism in ectothermic animals, yet rarely is the clinical practitioner in a setting where such research standards are easily met. Researchers are often stumped on creating even a good pain model to evaluate analgesic efficacy. So what do practitioners do to manage pain in their patients while these studies continue to slowly be published? As seen in birds, an analgesic drug and dosage that works in one species of Amazon parrot may not do so in another Amazon parrot species, and even preliminary reptile studies suggest significant species variation. Determining whether a surgical plane of anesthesia is reached with a particular product in a particular individual of a particular species for a particular procedure creates so many variables, that one can almost be paralyzed trying to compensate for all of these factors.

The Classes Amphibia and Sauropsida (reptiles and birds, Reptilia is no longer correct) are very diverse, broad classes comprising many thousands of species, some very primitive, some very advanced. While all anatomic systems can be affected by anesthesia and analgesia, the most commonly focused on is the cardiopulmonary system. Further information on amphibians can be found in Wright and Whitaker⁷ and in Mader³ on crocodilians. Most reptiles have a three-chamber heart with two atria, one partition-able ventricle, and two aorta that empty systemically. The degree of mixing of oxygenated and deoxygenated blood in the three-chamber heart is variable depending on species and their physiological state. Under certain conditions, deoxygenated blood is shunted back to the body or oxygenated blood is shunted back to the lungs. This variation in blood flow has been hypothesized to allow more effective thermoregulation and longer diving times for aquatic species, but has not been shown to be a fitness advantage.¹ Some snake and lizard species (e.g., varanids and pythons) have three-chambered hearts that function as four-chambered hearts during contraction due to a muscular ridge incompletely subdividing the ventricle during diastole and completely during ventricular systole. Because of this ridge, some of these squamates (lizards and snakes) can producing ventricular pressure differentials equivalent to mammalian hearts.⁶ All reptiles use lungs, however, aquatic turtles have developed more permeable skin, and some species have modified their cloaca to increase the area for gas exchange. Even with these adaptations, lungs are still required for respiration. In squamates, the lungs are ventilated almost exclusively by the axial musculature that is also used for locomotion. This forces most squamates to hold their breath during intense runs. Varanids and a few other lizard species, complement their normal respiration with buccal pumping, allowing the lizard to completely fill their lungs during intense activity, and thus remain aerobically active longer. Turtles and tortoises have a variety of solutions to breathing. Some have a sheet of muscle enveloping the lungs. When it contracts, the turtle exhales; retraction of limbs into the body cavity forces air out of the lungs; protraction of the limbs, reduces lung pressure leading to inhalation. Turtle lungs are attached to the carapace, with the bottom of the lungs attached to the viscera. Box turtles (Terrapene sp.) breathe continuously during locomotion, and ventilation is not coordinated with limb movements.² They are probably using their abdominal muscles to breathe during locomotion and have also been observed to breathe while completely sealed up inside their shells. Red-eared sliders also breathe during locomotion, but had smaller breaths during locomotion than during small pauses between locomotor bouts, indicating possible mechanical interference between limb movements and breathing.

With inhalant anesthetics, two physiologic concepts need to be remembered. First, reptiles breathe due to LOW partial pressure of oxygen. So to recover an intubated reptile, room air or air with less than 100% oxygen is advised to be used, or else the reptile may take hours longer to recover than what it should. Secondly, reptiles also can bypass standard metabolism requiring oxygen. If they do not breathe a lot on their own or are regularly provided intermittent positive pressure ventilation (IPPV), reptiles can revert to utilizing the effective but energy inefficient option of anaerobic respiration. Not only does this anesthetic patient not intake oxygen, but they are also not getting anesthetic and will respond to noxious stimuli such as cutting during surgery.

Do not forget the power of local anesthesia. Lidocaine 2% can be buffered one part to one part with sodium bicarbonate then combined one part buffered lidocaine with one part bupivacaine to provide almost immediate local anesthesia/analgesia effect with longer term effect.

Amphibians

For physical restraint, one should always wear latex gloves. This protects the animal from chemicals on one's hands, but also protects the handler in the case of toxins that some amphibians emit from their skin (Bufo sp.--toads). Moisten the gloves and also unbleached paper towels to use as a procedure substrate with distilled water. For anesthesia in amphibians, I usually rely either on tricaine methanesulfonate (MS-222) or isoflurane.

With MS-222 (Finquel^®), the first challenge is where to get it. I recommend Argent Labs: http://www.argent-labs.com/argentwebsite/trandrug.htm

Store in refrigerator until needed dry. The current cost is about $20 for 5 grams, which I divide into five equal parts. The dosages recommended in Mader 3 are 0.05% to anesthetize tadpoles, 0.1-0.2% for a surgical plane in most anurans and urodeles, and 0.3% for toads. How to make this "solution" is: 1g/2L = 0.05%, 1g/L = 0.1%, 2g/L = 0.2%, and 3g/L = 0.3%. The most important step is to buffer with an equal volume of baking soda powder (sodium bicarbonate) to MS-222, and add to a small quantity of distilled water (200mL). It usually bubbles quite a bit. I add more baking soda until bubbling stops. Then add that mixture to complete one or two liter volume. This is my stock solution. Have a second "solution" of only distilled water also ready. In a small container with a lid, add the amphibian to the MS-222 water that is full enough to partially submerge the animal. Erythema (especially ventrally) is the first sign of induction. A light plane of anesthesia is indicated by the loss of righting and/or corneal reflexes. A deep plane is indicated by only a cardiac impulse, with a withdrawal reflex the last to go. Once in a deep plane, place on paper towels moistened only with distilled water and titrate anesthesia as needed by trickling MS-222 water onto the animal. Generally, you have a surgical plane for 20 minutes, and there is no cardiovascular effect except for a slight increase in heart rate. Recovery is induced by gentle showers of pure distilled water, followed by soaking in shallow distilled water with paper towels.

Isoflurane can be used when MS-222 is not immediately available. When I have to do this, I bubble 5% isoflurane (with oxygen flow rate high enough to make bubbles) into a small container (minimal turn-around ability for amphibian) with distilled water until the anesthetic indicators noted above for MS-222 are reached. It can take much longer to reach 100% isoflurane using 1cc syringe with 25 gauge needle directly on ventral skin or intracoelomically if laparotomy, until plane reattained, or rebathe in isoflurane "bubbler" if skin has not been incised. Recover with the aforementioned distilled water "showers." Direct isoflurane can irritate their skin.

For analgesia, there are few useful studies from a clinical perspective for amphibians. I use Metacam^® (Meloxicam, Boehringer Ingelheim Vetmedica, Inc., 2621 N. Belt Hwy. St. Joseph, MO 64506-2002, USA) at 0.2 mg/kg IM, SQ, or PO q24h.

Reptiles (General Reptiles)

All reptiles need to be kept warm (85-90°F) during procedures without getting thermal burns. Bair blankets provide the best thermoregulation during anesthesia in my opinion. With general anesthesia, provide intermittent positive pressure ventilation (IPPV) throughout procedures (4 deep breaths/minute), as most reptiles do not always breathe well on their own in terms of frequency and depth. Endotracheal tubes are often smaller than one would expect, so modified tomcat catheters or 14 gauge IV catheters without stylets can be helpful when 2.0 diameter tubes do not fit. For recovery from inhalant anesthesia, I connect the endotracheal tube to an ambu-bag to introduce room air into the system, as it is LOW OXYGEN that stimulates the reptile to breathe on its own. In many cases, gentle movement of the endotracheal tube within the airway, as if to extubate or introduce deeper will stimulate the animal to try to move and expel the tube. Pinching various parts of the body and reassessing the righting reflex are good indicators as to when the endotracheal tube should be pulled. Check for mucous in the tube and at the tracheal opening. If uncertain if reptile is alive, I use the rule that if the trachea closes with tube removal, there is still hope. Remember that heart beat does not equate to being alive in reptiles.

Snakes

With snakes, I tend to try to keep it simple. Assuming a non-venomous species and an adequate number of restrainers (at least one person per four feet of snake), I open the snake's mouth and apply a drop or two of lidocaine 2% buffered one part to one part with sodium bicarbonate to the tracheal region. Some snakes have a tracheal flap to produce a particular hiss, which necessitates being creative in modifying/making an endotracheal tube, always look for this in unusual species if uncertain before beginning anesthesia. After a few minutes, I gently introduce a water lubricated endotracheal tube (uncuffed or uninflated) into the trachea, usually to the full length of the tube if able to easily advance. Then administer IPPV gently with a non-rebreathing system until the entire snake is sedated to the desired level. Usually complete anesthesia is at an acceptable level once there is no righting reflex or stimulus to noxious stimuli. Sometimes this can be challenging to assess due to reflexive muscle/skin "twitching" from the stimulus of cutting tissue. Monitor heart beat with a Doppler ultrasound unit taped directly over the heart (20-30% of head to cloaca body length). For recovery, I start backing down the isoflurane level from surgical plane during skin closure, so that it is at 0% a bit before the final suture is placed. Obviously, response during this decrease will necessitate re-evaluation of levels.

Analgesia in snakes is extrapolated from other reptiles and mammals with the exception of one study that found butorphanol at a particular dosage was ineffective in corn snakes.⁴ At this time, there are no known published studies evaluating analgesia in snakes. I use metacam (Meloxicam, Boehringer Ingelheim Vetmedica, Inc., 2621 N. Belt Hwy. St. Joseph, MO 64506-2002, USA) at 0.2 mg/kg q24h IM/SQ. I do not use opioid analgesics, lacking good supporting data.

Lizards

With lizards, I usually mask down with isoflurane with a tight seal, covering the eyes and gently stimulating the lizard to breathe by "tickling" or gently pinching. Once most stimulation response is gone, I open the lizard's mouth and apply a drop or two of lidocaine 2% buffered one part to one part with sodium bicarbonate to the tracheal region and replace the mask. After a few minutes, I gently introduce a water lubricated endotracheal tube (uncuffed or uninflated) into the trachea, usually to the full length of the tube if able to easily advance. Then administer IPPV gently with a non-rebreathing system until the entire lizard is sedated to the desired level. Usually complete anesthesia is at an acceptable level once there is no stimulus to noxious stimuli. Sometimes this can be challenging to assess due to reflexive muscle/skin "twitching" from the stimulus of cutting tissue. Monitor heart beat with a Doppler ultrasound unit taped directly over the carotid artery/jugular region or even the eye in large lizards and over the heart in smaller specimens. For recovery, I start backing down the isoflurane level from surgical plane during skin closure, so that it is at 0% a bit before the final suture is placed. Obviously, response during this decrease will necessitate re-evaluation of levels.

Propofol can be used for induction at 5 mg/kg IV in the tail vein, but it is important to remember that it provides no analgesia in mammals and such is also assumed in reptiles. Isoflurane or local anesthesia is generally used to supplement in these cases.

Analgesia in lizards has some preliminary studies, but most is extrapolated from other reptiles and mammals. I use metacam (Meloxicam, Boehringer Ingelheim Vetmedica, Inc., 2621 N. Belt Hwy. St. Joseph, MO 64506-2002, USA) at 0.2 mg/kg q24h IM/SQ and am currently evaluating the use of morphine at 10-20 mg/kg (study was in bearded dragons).⁴

Chelonians (Turtles and Tortoises)

With the challenges of getting to the head of chelonians, these are species that I always start with injectable anesthetics, though intubation and IPPV become critical to incorporate. Isoflurane can be used to supplement dependent upon the procedure.

I like to use a medetomidine/ ketamine chloride dose of 0.05-0.1 mg/kg and 10-15 mg/kg respectively IM or SQ, dependent on how the math works out for smaller specimens. Significant sedation or anesthesia should be expected within 15-30 minutes, assuming the chelonian is being kept warm. Once most stimulation response is gone, I open the chelonian's mouth and apply a drop or two of lidocaine 2% buffered one part to one part with sodium bicarbonate to the tracheal region. After a few minutes, I gently introduce a water lubricated endotracheal tube (uncuffed or uninflated) into the trachea, just a slight ways into the trachea as chelonians have short tracheas before bifurcation. Then administer IPPV gently with a non-rebreathing system until the entire chelonian, supplementing with isoflurane until the chelonian is sedated to the desired level. Usually complete anesthesia is at an acceptable level once there is no stimulus to noxious stimuli. Sometimes this can be challenging to assess due to reflexive muscle/skin "twitching" from the stimulus of cutting tissue. Monitor heart beat with a Doppler ultrasound unit taped directly over the left shoulder. For recovery, I start backing down the isoflurane level from surgical plane during skin closure, so that it is at 0% a bit before the final suture is placed. Obviously, response during this decrease will necessitate re-evaluation of levels.

Analgesia in chelonians has some preliminary studies, but most is extrapolated from other reptiles and mammals. I use metacam (Meloxicam, Boehringer Ingelheim Vetmedica, Inc., 2621 N. Belt Hwy. St. Joseph, MO 64506-2002, USA) at 0.2 mg/kg q24h IM/SQ and am currently evaluating the use of morphine at 1.5 mg/kg (study was in red-eared sliders)⁵, though it did cause a concerning level of respiratory depression at this dose in this species.

References

1.  Hicks, J. 2002. The physiological and evolutionary significance of cardiovascular shunting patterns in reptiles. News in Physiological Sciences 17: 241-245.

2.  Landberg, Tobias; Mailhot, Jeffrey; Brainerd, Elizabeth (2003). Lung ventilation during treadmill locomotion in a terrestrial turtle, Terrapene carolina. Journal of Experimental Biology 206 (19): 3391-3404.

3.  Mader DR (ed). 2006. Reptile Medicine and Surgery, 2nd Edition. Elsevier: St. Louis, MO.

4.  Sladky KK, Kinney ME, Johnson SM. 2008. Analgesic efficacy of butorphanol and morphine in bearded dragons and corn snakes. JAVMA 233 (2):267-273.

5.  Sladky KK, Miletic V, Paul-Murphy J, et al. 2007. Analgesic efficacy and respiratory effects of butorphanol and morphine in turtles. JAVMA 230 (9):1356-1362.

6.  Wang T, Altimiras J, Klein, W Axelsson M. 2003. Ventricular haemodynamics in Python molurus: separation of pulmonary and systemic pressures. Journal of Experimental Biology 206: 4242-4245.

7.  Wright KM & Whitaker BR. 2001. Amphibian Medicine and Captive Husbandry. Krieger: Malabar, FL.