Clinical Anesthesia in Reptiles

TOPICS IN MEDICINE AND SURGERY CLINICAL ANESTHESIA IN REPTILES Kurt K. Sladky, MS, DVM, Dip. ACZM, and Christoph Mans, med. vet.

Abstract The clinical use of anesthetic agents in reptiles presents a number of unique challenges because of the diversity of the class Reptilia with respect to natural history, size, anatomy, and physiology. Reptiles are commonly maintained as companion animals, widely displayed in zoological institutions, and many species serve as subjects in laboratory facilities. Therefore, to become a skillful clinician, developing an understanding of anesthetic efficacy across reptile species is important. The objective of this review is to provide a current perspective on the practical application of anesthetic agents in commonly maintained pet reptile species. Copyright 2012 Elsevier Inc. All rights reserved. Key words: anesthesia; anesthetic; reptile; sedation

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istorically, managing anesthesia in reptiles has been problematic for 3 primary reasons. First, there is an enormous diversity of species across the class Reptilia, one of the most phylogenetically diverse animal classes with 4 main orders (Crocodylia, Testudines, Squamata, and Rhynchocephalia), and greater than 7800 species worldwide. Second, there is a lack of systematic research dedicated to advancing the current understanding of effective anesthetic/analgesic induction and maintenance drugs, dose-dependent effects, duration of drug efficacy, interspecies’ differences, and potentially fatal drug-related adverse effects, such as cardiopulmonary depression. Lack of funding in support of such research has hindered scientific advancement and limited the application of anesthetic/analgesic agents in reptiles, when compared with mammals. Third, the reptile literature is rife with anecdotal information, which has persisted and, in many cases, has grown to be widely accepted dogma for clinicians. The diversity of reptiles in terms of natural history, size, and anatomy and physiology presents a unique clinical challenge to the veterinarian. However, as with all nondomestic species, the application of safe and effective anesthetic techniques is essential for those veterinarians treating reptile patients, and effective anesthetic application will facilitate performing complete physical examinations, collecting appropriate and high-quality diagnostic samples, and realizing successful surgical procedures. REPTILES AS ANESTHETIC PATIENTS: ANATOMIC, PHYSIOLOGIC, AND BEHAVIORAL CONSIDERATIONS Thermoregulation Reptiles differ from mammals both physiologically and anatomically, therefore it is difficult to directly extrapolate from those methods used in mammalian anesthesia. Most importantly, reptiles are poikilothermic, meaning that their body temperature is directly dependent on environmental temperature. Changes in body tempera-

ture significantly affect metabolic rate and many physiologic processes. Therefore, the temperature at which one maintains a patient is an important factor in the successful performance of reptilian anesthesia, because of the fact that the metabolism and excretion of drugs in reptile species are directly related to environmental temperature.1-3 Consequently, it is important to maintain the reptile patient at its preferred optimal body temperature for one to better predict the physiologic effects of anesthetic drugs, as well as ensure proper drug metabolism and excretion. The pre-

From the Department of Surgical Sciences, Zoological Medicine, School of Veterinary Medicine, University of Wisconsin, Madison, WI USA. Address correspondence to: Kurt K. Sladky, MS, DVM, Dip ACZM, Department of Surgical Sciences, Zoological Medicine, School of Veterinary Medicine, University of Wisconsin, 2015 Linden Dr, Madison, WI 53706. E-mail: [email protected]. © 2012 Elsevier Inc. All rights reserved. 1557-5063/12/2101-$30.00 doi:10.1053/j.jepm.2011.11.013

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The preferred optimal temperature zone for reptiles is generally considered to be 20°C to 25°C in temperate and aquatic species, and 25°C to 35°C in tropical species.

ferred optimal temperature zone (POTZ) for reptiles is generally considered to be 20°C to 25°C in temperate and aquatic species, and 25°C to 35°C in tropical species.4 Cardiopulmonary Systems

The reptile cardiovascular system is characterized by several unique anatomic structures and physiologic functions. In the chelonians (e.g., turtles, tortoises), lizards, and snakes, the heart is 3 chambered, composed of a single ventricle and 2 atria. The 2 atria are completely separate, and the ventricle is divided longitudinally into 2 chambers by an incomplete intraventricular septum. The incomplete septum allows mixing of oxygenated and deoxygenated blood, and shunting of blood from both left to right and right to left. All reptiles have 2 aortic arches, right and left. Although cardiac shunting is considered detrimental in birds and mammals, cardiac shunting and mixing of oxygenated and deoxygenated blood may be an important physiologic mechanism in reptiles during apneic periods (e.g., diving). The functional significance of cardiac shunts in reptiles has long been a subject of debate and is beyond the scope of this clinical review.5,6 However, the clinician should be aware that cardiac shunting can play a role in inhaled anesthetic uptake and elimination, potentially causing delayed induction, and delayed or an unexpectedly rapid recovery. In addition, cardiac shunts may affect systemic blood pressure and arterial/venous oxygen concentration, which in turn will impact anesthetic monitoring (e.g., indirect blood pressure, pulse oximetry, expired gases). A unique feature of the reptile cardiovascular system is the renal portal system, which is essentially a ring of blood vessels around the kidney. The cranial portal vein (a branch of the external ischiatic vein) and caudal portal vein (venous blood from the capillary system of the limb, tail, pelvis, and caudal part of intestine and spine) form a venous plexus located just ventral to the kidney.7 Valves are located within veins to regulate venous blood flow through or around the kidney. The valves are under parasympathetic and sympathetic control, with venous blood being directed through the kidneys when valves are

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closed (cholinergic stimulation) or bypassing the kidneys and being directed into the caudal vena cava and liver when valves are open (adrenergic stimulation). The clinical importance of the reptile renal portal system remains a bit of an enigma. Therefore, the current recommendation is to avoid administration of drugs into the caudal half of the body, the hind limbs, and/or tail because of more rapid clearance and lower plasma levels compared with administration into the cranial half of the body.8,9 The respiratory system of reptiles is characterized by primary pulmonary respiration, although some aquatic species have evolved mechanisms of extra-pulmonary respiration to cope with hypoxic environments. The reptilian lung is generally considered fragile and composed of a simple, endothelial-lined sac in most reptile species. Lizards and chelonians have paired, saclike lungs. Most snakes have a vestigial left lung and a functional right lung, which ends distally as an airsac. Boid snake species tend to have functional right and left lungs, although the left tends to be smaller and also has caudal airsacs. Of the noncrocodilian reptiles, the lung is typically septate, unlike the alveolar lungs of mammals. Reptiles lack a muscular diaphragm separating the thoracic and abdominal cavities, thus a variety of skeletal and smooth muscles facilitate breathing. Chelonians use pelvic and pectoral muscles, which are functionally analogous to a mammalian diaphragm. Snakes use a combination of smooth muscle in the lung walls as well as intercostal muscles, whereas lizards use lung smooth muscle, intercostal, pectoral, and abdominal muscles for respiration. Physiologically, reptiles have a low metabolic rate, slower respiration rate, and a lower rate of oxygen consumption than mammals. Respiration in reptiles is controlled by oxygen and carbon dioxide concentrations in the blood as well as environmental temperatures.10 Reptiles tolerate hypoxic conditions, whether it is natural hypoxia or induced breathholding during anesthesia, because they are capable of metabolically converting to a state of anaerobic metabolism. This tolerance to hypoxia seems to depend on cardiovascular shunts, environmental temperature, and the ability to buffer lactic acid.7 PREANESTHETIC PATIENT ASSESSMENT As with all patients, reptiles scheduled to undergo general anesthesia should have their health status evaluated. Systemic disease processes may

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preclude general anesthesia, or may warrant a delay until the patient has received supportive care. With the exception of elective procedures or acute disease (e.g., trauma), most reptile patients present with a history of chronic problems, frequently associated with inadequate husbandry. Therefore, a thorough history and review of husbandry, including diet, is critical before the physical examination and collection of diagnostic samples. The physical examination should be systematic and include an evaluation of all organ systems. Particular attention should be paid to the hydration and cardiopulmonary status of the patient. In addition, a complete blood count, plasma or serum biochemical profile, and diagnostic imaging should be considered. Anemia, dehydration, and underlying organ dysfunction can cause significant complications during anesthesia, and may lead to delayed or nonrecovery from general anesthesia. Dehydrated patients should be adequately rehydrated, and any underlying disease processes identified and treated before an anesthetic procedure. Fluid Support Fluid support is indicated for those individuals demonstrated to be dehydrated based on the results of the preanesthetic physical examination and diagnostic procedures. Fluids should be administered at a constant rate using an intravenous (IV) or intraosseous (IO) route in moderately to severely dehydrated reptiles. Alternatively, subcutaneous (SC) or intracoelomic (ICe) administration of fluid as a bolus can be performed. Great care should be taken to avoid any iatrogenic damage of coelomic organs (i.e., mature follicles in chelonians and lizards) during ICe injection. With few data available providing evidence for the most appropriate fluid type to administer to reptiles, it is best to choose a standard balanced electrolyte solution (e.g., lactated Ringer’s, Normosol). The authors recommend a maintenance rate of 15 to 30 mL/kg/d delivered either IV/IO as constant-rate infusion, or as a bolus administered SC or ICe. Fluid deficits, secondary to dehydration, should be corrected by adding to the daily maintenance fluid requirements. CHEMICAL RESTRAINT Injectable Anesthetics and Sedatives Sedation. Procedural sedation is defined as the state of drug-induced altered consciousness,

FIGURE 1. IV propofol administration in the left brachial plexus of a sedated African spurred tortoise. Propofol was used for induction of short-term anesthesia (approximately 15 minutes) for esophageal feeding tube placement.

which allows the patient to better tolerate stressful or unpleasant procedures without depressing protective airway reflexes or having a significant cardiopulmonary depressant effect.10 Procedural sedation is commonly used in human and domestic animal veterinary medicine to facilitate diagnostic testing (e.g., imaging) and sample collection (Fig 1). Currently, diagnostic procedures are often performed in conscious animals or when they are under general anesthesia. A variety of injectable drugs are reported to be effective when administered alone or in combination for sedation of reptile patients.11-14 Preference should be given to sedative combinations that are partially or completely reversible to avoid prolonged or unpredictable recoveries. Sedative combinations should be selected based on the desired level of sedation (mild to deep), the general condition of the patient, and the diagnostic and/or therapeutic procedure to be performed. The authors prefer combinations of midazolam with an alpha-2-agonist (e.g., medetomidine or dexmedetomidine), to which either ketamine at a low dose, or an opioid, can be added for deeper sedation and/or improved analgesia. Commonly used sedative protocols are summarized in Table 1. Sedated animals should maintain spontaneous breathing and should be responsive to overtly painful stimulation. The combination of sedation, with either local or spinal anesthesia, offers the possibility of performing surgical or invasive procedures, which would otherwise only be possible under general anesthesia. By combining se-

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2 0 TABLE 1. Sedation protocols commonly used in reptiles Dosage (mg/kg) Chelonians Midazolam

Route

Species

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2

SC

RES

Dexmed ⫹ midaz

0.1 ⫹ 1.0

SC

Dexmed ⫹ midaz ⫹ ketamine

0.1 ⫹ 1.0 ⫹ 2.0

SC

RES NA box turtle RES

Dexmed ⫹ midaz ⫹ ket

0.07 to 0.1 ⫹ 1.0 ⫹ 2.5 to 5.0

SC

African spurred tortoise

Med ⫹ ket

0.1 ⫹ 5.0

IM

RES

Med ⫹ ket ⫹ morphine

0.15 ⫹ 2.5 ⫹ 1.0

SC

African spurred tortoise

Med ⫹ midaz ⫹ morphine

0.2 ⫹ 2.0 ⫹ 1.0

SC

African spurred tortoise

2.0 to 5.0

IV

0.1 ⫹ 1.0

SC

0.05 – 0.1 ⫹ 1.0 ⫹ 3.0

SC, IM

3.0 to 5.0

IV, IO

1.0 2.0 5.0 3.0

SC, IM SC, IM SC, IM IV

Propofol Lizards Dexmed ⫹ midaz Dexmed ⫹ midaz ⫹ ket

Propofol Snakes Midaz Telazol Ket Propofol

to to to to

2.0 5.0 10.0 5.0

Green iguana Bearded dragon Bearded dragon Uromastyx

Large snakes

Comments Mild sedation, inconsistent effects Completely reversible; rapid recovery after reversal Mild to moderate sedation Completely reversible; rapid recovery after reversal Moderate to deep sedation Partially reversible; rapid recovery after reversal Suitable for intrathecal injection Moderate to deep sedation Partially reversible; rapid recovery after reversal Suitable for cloacoscopy Deep sedation Suitable for intubation and nonpainful procedures Deep sedation Partially reversible; rapid recovery after reversal Suitable for cloacoscopy Moderate to deep sedation Completely reversible; rapid recovery after reversal Suitable for cloacoscopy Moderate sedation—light anesthesia Moderate sedation Completely reversible; rapid recovery after reversal Deep sedation, suitable for intraoral examination, endotracheal intubation, and minor surgery if used together with local anesthesia, partially reversible Deep sedation-light anesthesia Mild sedation, inconsistent effects Mild to moderate sedation, endotracheal intubation Mild to moderate sedation, endotracheal intubation Moderate sedation—light anesthesia

Reference 12 *

19,20 27 18 27 27 16 * *

16 16 21 21

16,17

Abbreviations: RES, Red-eared slider turtle; Dexmed, dexmedetomidine; Ket, ketamine; Med, medetomidine; Midaz, midazolam; SC, subcutaneous; IM, intramuscular; IV, intravenous; IO, intraosseous; NA, North American. *Mans et al, unpublished. Atipamazole was used to antagonize medetomidine (1 mg/ml) and dexmedetomidine (0.5 mg/ml) in 1:1 volume, SC. Flumazenil was used to antagonize midazolam: 0.05 mg/kg SC or 13:1 midazolam:flumazenil (mg). Nalaxone was used to antagonize morphine: 0.04 mg/kg SC.

dation with local or spinal anesthesia, the dose of a general anesthetic drug can be reduced, which will lead to reduced cardiovascular depression, more rapid recoveries, and potentially fewer anesthetic complications compared with general anesthesia alone. Anesthesia. A variety of injectable anesthetic protocols have been reported in different reptile species. However, the published dosages vary widely for many drugs, which is partially because different species of reptiles require different dosages to achieve the same effect. In general, we recommend the avoidance of administering high doses of a single anesthetic agent, and instead consider protocols in which multiple drugs are combined with synergistic actions, thereby requiring lower dosages for each drug. Additionally, using readily reversible drug protocols will result in more rapid recoveries. Because most deleterious side effects (e.g., prolonged recovery, cardiopulmonary depression) associated with anesthetic and sedative drug administration are dose dependent, individual drug dosage reduction and reversibility will result in fewer complications and improved recoveries. Table 2 summarizes commonly used anesthetic protocols in reptile species. One must keep in mind that many factors (e.g., site of injection, underlying disease, dehydration, body temperature, gravidity) may affect the onset, recovery, and efficacy of anesthetic drugs.1-3,15 Therefore, an individualized approach to choosing appropriate drug combinations and dosages should be considered. Routes of Parenteral Drug Administration. Historically and anecdotally, IM administration of anesthetic agents was the preferred route in reptiles.5,16 Many veterinarians believed that the SC space of reptiles was not well vascularized and therefore drug absorption would be prolonged and incomplete. However, advantages associated with SC drug administration include access to a variety of SC sites requiring minimal manual patient restraint or manipulation and, consequently, increased safety and the capability of administering larger volumes in a single location. Results from our laboratory indicate that drug effects can be measured rapidly after SC administration.17 Overall, although the induction times after SC administration of ketamine-dexmedetomidine in red-eared slider turtles (Trachemys scripta elegans) were significantly longer and more variable compared with IM injection, the depth of anesthesia achieved within 45 minutes was

not significantly different between IM or SC injection.15 Therefore, the authors routinely administer anesthetic drugs and their antagonists SC in chelonians, lizards, and snakes (Fig 2), particularly if larger volumes of anesthetic drugs need to be injected. Anesthetic drugs administered into the caudal half of the body or tail of reptiles have traditionally been avoided because of the presence of a renal-portal system. Recently, we compared induction times of ketamine-dexmedetomidine (SC) administered in the forelimbs and hind limbs of red-eared slider turtles and found that anesthetic induction was achieved after forelimb administration, but not after hind limb administration.15 Therefore, the authors support the current recommendation that all anesthetic drugs be administered in the cranial half of the body. Table 3 summarizes preferred drug administration sites in reptiles.

Overall, although the induction times after SC administration of ketaminedexmedetomidine in red-eared slider turtles (Trachemys scripta elegans) were significantly longer and more variable compared with IM injection, the depth of anesthesia achieved within 45 minutes was not significantly different between IM or SC injection.

Parenteral Sedative and Anesthetic Drugs. Ketamine, a dissociative agent with anesthetic and analgesic properties, is frequently used in reptile anesthesia. However, when administered alone, muscle relaxation is considered inadequate and recoveries excessively prolonged, especially when high dosages are used. Very high dosages of ketamine (up to 90 mg/kg) have been published, which are commonly used alone or in combination with alpha-2-adrenergic agonists or benzodiazepines.5,16,17 Preferable to administering ketamine alone at high dosages, is its combination with an alpha-2-adrenergic agonists, which provides surgical anesthesia in many reptile species.17,18 Administering alpha-2-adrenergic antagonists allows partial reversibility of these protocols, leading to more rapid recoveries and increased safety. Even at low dosages, ketamine (⬍ 5 mg/kg) can provide additional sedation and analgesia if combined with other anesthetic drugs. When combined with midazolam-dexmedetomidine (1 mg/kg ⫹ 0.1 mg/kg), ketamine (2 mg/kg) provided deep sedation in red-eared

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2 2 TABLE 2. Anesthesia protocols commonly used in reptiles Dosage (mg/kg) Chelonians Med ⫹ midaz ⫹ ket ⫹ hydromorphone

Route

Species

0.15 ⫹ 0.5 ⫹ 10 ⫹ 0.5

SC

African spurred tortoise

Med ⫹ ket

0.2 ⫹ 10.0

IM

RES

Med ⫹ ket

0.1 ⫹ 10.0

IV

Med ⫹ ket ⫹ morphine Sladky and Mans/Journal of Exotic Pet Medicine 21 (2012), pp 17–31

Propofol Propofol Isoflurane Sevoflurane Lizards Propofol

Alfaxalone Alfaxalone Isoflurane Sevoflurane Snakes Alfaxalone Propofol Telazol Isoflurane Sevoflurane

0.1 ⫹ 10.0 ⫹ 1.5

IM

Galapagos tortoise Map turtle

2.0 to 10.0 10.0 to 20.0

IV IV

RES

2 to 5% 2.5 to 8% 5.0 to 10.0

IV,IO

Green iguana

10.0 9.0 2.0 to 5.0% 2.5 to 8.0%

IV IV

Green iguana

6.0 to 15.0 3.0 to 5.0 2.0 to 6.0 2.0 to 5.0% 2.5 to 8.0%

IV IV IM

Comments Surgical anesthesia. Deepen with gaseous anesthetic if required Partially reversible; rapid recovery after reversal Surgical anesthesia Partially reversible Surgical anesthesia Partially reversible Surgical anesthesia. Partially reversible; rapid recovery after reversal; administration of nalaxone prevents renarcotization Induction agent; use lower dose in large tortoises Induction agent; light anesthesia maintained for 60 min (10 mg/kg) and approximately 90 min (20 mg/kg) Induction 5.0%, maintenance 2.0% to 3.0% Induction 7.0% to 8.0%, maintenance 2.5% to 4.5% Induction agent; duration of action approximately 20 to 30 min (5 mg/kg) Maintain with gaseous anesthetic Induction agent; maintain with gaseous anesthetic Induction agent; maintain with gaseous anesthetic Induction 5.0%, maintenance 2.0% to 3.0% Induction 7.0% to 8.0%, maintenance 2.5% to 4.5% Induction Induction Induction Induction Induction

agent; maintain with gaseous anesthetic agent; maintain with gaseous anesthetic agent; maintain with gaseous anesthetic 5.0%, maintenance 2.0% to 3.0% 7.0% to 8.0%, maintenance 2.5 to 4.5%

Reference *

18 26 23

16,17 28

16,17 16,17 17,50 36 25

16,17 16,17 35 16 16

16,17 16,17

Abbreviations: RES, Red-eared slider turtle; Dexmed, dexmedetomidine; Ket, ketamine; Med, medetomidine; Midaz, midazolam; SC, subcutaneous; IM, intramuscular; IV, intravenous; IO, intraosseous. *Mans et al, unpublished.

TABLE 3. Preferred sedative and anesthetic injection sites in reptiles

FIGURE 2. Coelomic ultrasonography in an African spurred tortoise (Geochelone sulcata) after sedation using a combination of dexmedetomidine (0.075 mg/kg), midazolam (1 mg/kg), and ketamine (2.5 mg/kg). Moderate sedation was achieved within 30 minutes after SC injection. Atipamazole (0.75 mg/kg) and flumazenil (0.05 mg/kg) were administered SC for reversal.

slider turtles suitable for intrathecal injections and endotracheal intubation.19,20 Tiletamine/zolazepam (Telazol, Fort Dodge Laboratories, Fort Dodge, IA USA) is a commercially available drug combination composed of a dissociative anesthetic (tiletamine) and a benzodiazepine (zolazepam). Dosages of 2 to 10 mg/kg have been recommended for sedation, induction of general anesthesia, or facilitation of intubation.16,17,21 The anesthetic efficacy of tiletamine/zolazepam can be unpredictable and recovery is always prolonged. In dehydrated patients, or those with underlying renal or metabolic disorders, the use of this drug combination is contraindicated.17 Tiletamine/zolazepam may be most useful in large or dangerous species (e.g., crocodilians, large monitor lizards), but is not recommended by the authors for routine use in common pet reptile species. Alpha-2-adrenergic agonists, such as medetomidine and dexmedetomidine, provide sedation, muscle relaxation, and possibly analgesia in reptiles. However, dose-dependent cardiovascular depression has been documented in reptiles after medetomidine administration.14,22 Alpha-2-adrenergic agonists are commonly used in combination with ketamine, especially in chelonian species, for safe, reliable, and reversible anesthesia, but can also be combined with benzodiazepines for procedural sedation (Fig 3).18,23-27 Combining ketamine with medetomidine or dexmedetomidine allows for reduction of both drug dosages, and reversibility of the alpha-2-adrenergic agonists with atipamezole will lead to faster and more predictable recoveries.

Chelonians ● SC drug administration sites include the skin between the neck and forelimbs. ● IM drug administration sites include the forelimb muscles. ● IV access includes the supravertebral (subcarapacial) sinus, jugular vein, brachial vein, or dorsal coccygeal venous sinus. Lizards ● SC drug administration sites include the dorsum, overlying the epaxial muscles. Stay in the cranial half of the body. ● IM drug administration sites include the forelimb and epaxial muscles in the cranial half of the body. ● IV access includes the ventral coccygeal vein (i.e., ventral tail vein), jugular vein, or the palatine veins in the oral cavity of larger species. ● IO administration requires catheterization of the distal femur or proximal tibia. Snakes ● SC drug administration sites include the dorsum, overlying the epaxial muscles. Stay in the cranial half of the body. ● IM drug administration sites include the epaxial muscles in the cranial half of the body. ● IV access includes the ventral coccygeal vein (i.e., ventral tail vein), or the palatine veins in the oral cavity of larger species. Abbreviations: SC, Subcutaneous; IM, intramuscular; IV, intravenous; IO, intraosseous.

FIGURE 3. Red-eared slider turtle (Trachemys scripta elegans) demonstrating injection sites in the front half of the body. The SC site highlights the skin between the fore limb and neck. The IM site highlights the muscle bellies of the forelimb.

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Benzodiazepines, such as midazolam and diazepam, have sedative, anxiolytic, and muscle relaxant properties. Midazolam is water soluble, can be administered SC, IM, and IV, and is considered a more appropriate choice than diazepam, which is not recommended for IM or SC injection.5 Midazolam, used alone, provides mild, but highly variable sedation, which may be sufficient for minor clinical procedures.11,12 Midazolam, combined with ketamine or an alpha-2adrenergic agonist, reduces all drug dosages and attenuates the dose-dependent cardiovascular depressant effects and prolonged recoveries commonly observed with high dosages of ketamine. The effects of benzodiazepines can be antagonized with flumazenil (0.05 mg/kg, SC), which shortens recovery from sedation or anesthesia.27 However, the expense of flumazenil may preclude its use in larger animals. Propofol is one of the most commonly used induction agents across all species of reptiles. Propofol administered IV or IO results in smooth inductions and reliable chemical immobilization. However, intravascular access is required and induction times can be prolonged in reptiles compared with mammals.17 The depth and duration of anesthesia, as well as cardiopulmonary depressant effects, are dose dependent.28 Respiratory depression is also more profound when propofol is rapidly administered.17,28 Because propofol does not accumulate in tissues and is rapidly metabolized, recovery can be expected with assisted ventilation in cases of overdose. Propofol has been used as an induction agent to facilitate endotracheal intubation and maintenance on an inhalant anesthetic agent. Alternatively, propofol can be used as the sole anesthetic for short surgical procedures, such as esophageal feeding tube placement, aural abscess treatment, or hemipenile/phallus amputation. Because propofol is considered to have no analgesic properties, appropriate analgesic administration should be included in the anesthetic protocol if painful procedures are to be performed. Propofol requires intravascular administration, which can be challenging in some species. Recently, published reports have described complications secondary to accidental extravascular injection of propofol intrathecally and into the subcarapacial sinus of chelonian species.29,30 Complications associated with intrathecal propofol injection included forelimb and hind limb paralysis, coma, and spinal necrosis. Therefore, the current recommendation is to use other available venous access sites (e.g., jugular vein, brachial plexus) (Fig 3) before at2 4

tempting propofol administration into the subcarapacial sinus of chelonians. Alfaxalone is a neuroactive steroid anesthetic, which has been licensed for use in dogs and cats outside North America. It is a commonly used alternative to propofol in reptiles in Europe, Australia, and New Zealand. Unlike propofol, one major advantage of afaxalone is that it can be administered IM as well as IV.31 Afaxalone is rapidly cleared and its metabolism is independent of organ function. Similar to propofol, afaxalone administration is associated with smooth and rapid recoveries and minimal dose-dependent cardiovascular and respiratory depression in mammals.31-33 However, information on the use of afaxalone in reptiles is limited.34-36 In a recent preliminary study in native Australian lizards and snake species, the effects of afaxalone (9 mg/kg, IV) resulted in rapid induction in most investigated species, and spontaneous breathing was maintained in all animals.35 Although a surgical plane of anesthesia was not achieved, endotracheal intubation was possible. In green iguanas (Iguana iguana) afaxalone (10 mg/kg IV) resulted in rapid induction of anesthesia, which allowed endotracheal intubation.36 For IM administration, a dosage of up to 18 mg/kg has been recommended in reptiles.17,31 Opioid drugs administered alone do not have significant sedative effects in reptiles. Premedication of green iguanas with butorphanol (2 mg/kg IM) had no effect on induction times associated with isoflurane-induced anesthesia, and no isoflurane-sparing effects.37,38 Local Anesthesia Local anesthesia is currently underutilized in reptile medicine, but offers significant benefits, such as the ability to reduce the depth of general anesthesia or to use sedation for certain surgical procedures. In some cases, local anesthesia can be used solely in manually restrained animals for minor surgical procedures.17 Common indications for the use of local anesthetics as part of an anesthetic protocol in reptiles include distal limb surgery, tracheal or lung washes, or infiltration of incision sites before coeliotomy. Lidocaine and bupivacaine are readily available in most veterinary clinics, and both are effective local anesthetics in reptiles. The toxic dosages for local anesthetics have not been determined in reptiles. In mammals, toxic dosages for lidocaine have been reported to range from 5 to 20 mg/kg.17 Although toxic dosages in reptiles are unknown, care should be taken not to exceed those dosages

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FIGURE 4. Intrathecal lidocaine injection in the proximal coccygeal region of a red-eared slider turtle. A 28gauge, 1/2-inch needle attached to a 0.5-mL insulin syringe was used for intrathecal injection. The animal was sedated with dexmedetomidine-ketamine-midazolam before intrathecal injection at the proximal coccygeal level.

used in mammals, particularly in small reptile patients, to avoid systemic side effects. Spinal Anesthesia The application of segmental spinal anesthesia (also known as intrathecal anesthesia), in reptiles is currently in its infancy and has only been reported in tortoises and turtles.19,39 Most reptiles lack an epidural space, and therefore epidural anesthesia is not possible.40 Instead, intrathecal anesthesia is a feasible alternative, particularly in chelonian species.19,39 Intrathecal anesthesia involves the injection of anesthetic and analgesic drugs into the intrathecal space, which surrounds the spinal cord and is filled with cerebrospinal fluid. Because of the presence of the carapace and the fusion of the vertebral column to the carapace, access to the intrathecal space is limited in chelonian species, with the exception of the cervical and coccygeal vertebrae; intrathecal injections are most often performed in the coccygeal area.19,39 The sacral plexus arises from spinal nerves XVII to XXI located at the last dorsal and sacral vertebrae.41,42 Within the sacral plexus, the nerves interconnect and subdivide several times before terminating in the inguinal, pelvic, and hind leg muscles.43 The cloaca and genitalia are innervated by caudal branches of the sacral plexus and coccygeal nerves.44,45 Therefore, the temporary desensitization of these nerves, by intrathecal injection of local anesthetics and analgesics, should provide local anesthesia and analgesia sufficient for a multitude of surgical procedures involving the cloaca, urinary bladder, genitalia, and hind limbs. Spinal anesthesia using lidocaine at 1 mg/kg has been used successfully

for phallectomy in male Galapagos Local anesthesia tortoises (body weight 32-97 kg).39 is currently Recent studies performed in our laboratory evaluated the efficacy of underutilized in lidocaine and bupivacaine after reptile medicine, intrathecal injection in sedated male and female red-eared slider but offers turtles with a minimum body significant weight of 0.5 kg.19 Both intrathecal lidocaine (4 mg/kg) and bupivabenefits, such as caine (1 mg/kg) provided motor the ability to and sensory block of varying duration (Figs 4 and 5). Our prelimreduce the depth inary results indicated that intraof general thecal anesthesia is an effective and readily applicable clinical anesthesia or to technique in red-eared slider turuse sedation for tles with body weights as low as 19 0.5 kg. Sedation before intrathecertain surgical cal drug administration is recomprocedures. mended and only preservative-free preparations of anesthetic and analgesic drugs should be administered intrathecally to avoid spinal toxicity and associated neurologic complications. INHALANT ANESTHESIA Intubation All reptiles undergoing general anesthesia for longer than 15 minutes should be intubated to provide assisted ventilation and/or to deliver inhalant anesthetic gases for maintenance of general anesthesia. Intubation of reptiles is relatively straightfor-

FIGURE 5. Complete motor and sensory block of the hind limbs, prefemoral fossae, tail, cloaca, and penis in a male red-eared slider turtle after intrathecal injection of bupivacaine at the mid-coccygeal level.

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Using isoflurane or sevoflurane for maintenance of anesthesia requires endotracheal intubation and IPPV, because most reptiles develop apnea, or marked bradypnea and hypoventilation, when anesthetized.

ward. In contrast to mammals, reptiles do not have an epiglottis, and the glottal folds are maintained in a closed position, opening only during a respiratory cycle. Therefore, the application of lidocaine on the glottis can facilitate intubation in conscious or sedated animals. The glottis in snakes and lizards is located at the base of the tongue, and the larynx is highly mobile which makes intubation quite easy (Figs 6 and 7). In chelonians, the glottis is also at the base of the tongue, but those with fleshy, muscular tongues provide more of a challenge to intubate. Reptiles have a tremendous capacity for anaerobic metabolism and can hold their breath for extended periods of time. Mask, chamber, or snake tube inductions can be prolonged and/or may be totally unsuccessful depending on the species. Once intubated, caution should be exercised when considering endotracheal tube cuff inflation. It is easy to damage the fragile trachea with excessive inflation. Unlike snakes and lizards with incomplete tracheal rings, chelonians have complete tracheal rings and, therefore, cuffed endotracheal tubes should not be used. Additionally, in chelonians, the trachea can be short and it bifurcates cranial to the thoracic inlet, so caution should be used when advancing the endotracheal tube to avoid unilateral lung intubation.

FIGURE 6. Oropharynx of a green iguana (Iguana iguana) showing a prominent open glottis at the base of the tongue.

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FIGURE 7. An intubated green iguana being maintained on isoflurane gas.

Inhalant Anesthetic Agents Isoflurane and sevoflurane are the most commonly used inhalant anesthetics in reptiles. Inhalant anesthetics undergo minimal metabolism before elimination by the lungs. Therefore, underlying kidney or liver dysfunction does not affect the clearance of inhalant anesthetics, making these compounds preferred maintenance anesthetic agents in mammals and birds. However, because of right-to-left cardiac shunting in reptiles, reduced lung perfusion can occur; therefore concentrations of inhalant anesthetic gases in the lungs do not necessarily reflect the concentrations in the blood or brain.5 Sudden changes in shunting directions can lead to sudden changes in inhalant anesthetic blood concentration, which can lead to significant changes in anesthetic depth, evidenced by slow induction, sudden arousal from anesthesia, or prolonged recovery from anesthesia. In lizards and snakes, changes in gas concentration can be used to control anesthetic depth, but in chelonians, particularly aquatic turtles, a change in gas concentration may not lead to measurable changes in anesthetic depth. Nonrebreathing systems are recommended for reptile patients weighing less than 10 kg.16 It is preferable to anesthetically induce reptiles with injectable agents (Table 1 and 2), and then maintain anesthesia with inhalant gases. Snakes and lizards can be induced with gaseous anesthesia delivered either by mask, in a snake tube, or via an induction chamber (Figs 8, 9, 10). However, such inductions may require high concentrations of gas anesthetic, which are associated with environmental contamination and can contribute to unpredictable and prolonged induction times. Medium to large snakes can be intubated consciously (Fig 11) and in-

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FIGURE 10. Isoflurane anesthesia chamber induction of a snake.

FIGURE 8. Isoflurane gas anesthesia induction in a corn snake (Pantherophis guttata) using a standard large-sized mask.

should be performed cautiously. Limiting peak inspiratory pressure to ⬍10 cm H2O is recommended. In green iguanas the minimum anesthetic concentration (MAC) of isoflurane has been reported

duced with intermittent positive-pressure ventilation (IPPV). Venomous or aggressive snake species can be induced with a snake tube (Fig 12). Using isoflurane or sevoflurane for maintenance of anesthesia requires endotracheal intubation and IPPV, because most reptiles develop apnea, or marked bradypnea and hypoventilation, when anesthetized. Therefore, constant ventilatory support (approximately 2– 4 breaths per minute) should be provided throughout the anesthetic procedure. It is important to remember that the thin reptile lung and diminished exterior muscular support can lead to damage because of overexpansion/inflation during inhalant anesthetic administration and ventilation. Thus, IPPV

FIGURE 9. Isoflurane gas anesthesia induction in a green iguana using a standard medium-sized mask.

FIGURE 11. Conscious intubation of a snake. Once the snake was intubated, intermittent positive-pressure ventilation was used to deliver isoflurane gas for anesthetic induction.

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FIGURE 12. Isoflurane gas anesthesia induction in a snake using a snake tube.

to be 1.8% to 2.1%, and 3.1% ⫾ 1.0% for sevoflurane.38,46 For maintenance of general anesthesia, most reptiles require concentrations of 2% to 3% of isoflurane and 3.5% to 4.5% of sevoflurane.16 Debilitated reptile patients may need to be maintained on lower anesthetic gas concentrations to minimize the risk of cardiovascular depression.16 Isoflurane has been shown to significantly reduce blood pressure in green iguanas and ball pythons in a dose-dependent manner.43,46 Therefore, significant cardiovascular depression can occur if excessive concentrations of isoflurane, above the MAC, are delivered. No significant differences were detected between sevoflurane and isoflurane in cardiopulmonary function of green iguanas.44 However, use of sevoflurane resulted in faster induction and more rapid recovery compared with isoflurane.44 The authors recommend gradually reducing the concentration of inhalant anesthetic as the procedure is concluding, and discontinuing the inhalant anesthetic at the end of surgical procedures (i.e., at the time of SC

FIGURE 13. Green iguana in dorsal recumbency positioned for an ovariectomy. Note the ultrasonic Doppler flat probe is placed over the cranial chest between the forelimbs to monitor pulse/heart rate during anesthesia. With the Doppler probe in this location, the great vessels of the heart, or the heart itself, can be easily detected.

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FIGURE 14. Common map turtle (Graptemys spp.) in dorsal recumbency positioned for a prefemoral ovariectomy. Note the ultrasonic Doppler pencil probe placed in the thoracic inlet over the right carotid artery to monitor pulse rate during anesthesia.

tissue or skin closure). This will shorten the postprocedure anesthetic recovery period. MONITORING Monitoring of anesthetic depth and cardiopulmonary function is a continual clinical challenge in reptiles. Care should be taken to avoid noncritical application of anesthetic monitoring principles and devices used in mammals. Reptiles dramatically differ in their anatomy and physiology and are capable of maintaining physiological processes under extreme environmental circumstances, such as hypoxia and hypothermia. Monitoring of Anesthetic Depth A variety of physiological reflexes, as well as muscle tone, can be evaluated in reptiles to assess the depth of sedation or anesthesia. However, significant anatomic differences exist between snakes, chelonians, and lizards. The corneal and palpebral reflex is available for assessment in many reptiles, but cannot be appraised in snakes and most geckos (one exception being leopard geckos), because of the lack of eyelids and presence of spectacles. The righting reflex should be absent in snakes and lizards during a surgical plane of anesthesia, but is not as useful in turtles and tortoises.16 Assessment of head withdrawal and neck tone may provide useful information in turtles and tortoises. Snakes are likely to lose muscle tone from head to tail during induction, and regain muscle tone from tail to head during recovery.17 Deep or surgical planes of anesthesia have been associated with complete lack of movement, total muscle relaxation, and no response to painful stimuli (e.g., incision, hypodermic needle puncture).45

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FIGURE 15. Russian tortoise (Testudo horsfieldi) recovering from propofol-induced anesthesia for penile amputation. An Ambu bag is used to perform intermittent positive-pressure ventilation every 5 minutes during the recovery phase until the animal resumes spontaneous breathing. A flat ultrasonic Doppler probe is placed in the left thoracic inlet to monitor the pulse rate.

Cardiovascular Monitoring The heart rate in reptiles can be influenced by a variety of factors, including environmental temperature, metabolism, and presence of noxious stimulation.47 In addition to auscultation, the heart or pulse rate can be quantified in most reptile species by use of an ultrasonic Doppler device or sonography. The ultrasonic Doppler device is extremely useful and a necessary piece of equipment for cardiovascular monitoring of reptile patients. Flat and pencil probes are available. The ultrasonic Doppler probe can be placed over the heart in snakes and lizards, or the carotid arteries in lizards and chelonians (Figs 13 and 14). In lizards, the heart is located cranially between the forelimbs (except in monitor lizards), and the probe can either be positioned ventrally on the chest or in the axillary region. In chelonians the placement of a Doppler probe for heart rate measurement can be more challenging. Placing a flat or pencil probe on either side of the neck, within the thoracic inlet, can be effective in detecting the pulse rate via the carotid artery (Figs 14 and 15). If a Doppler device is not available, echocardiography, using an appropriate-sized probe, can serve as an effective way to monitor cardiac function and measure heart rate. Indirect blood pressure measurements can be measured using small cuffs wrapped around tails or limbs. However, indirect blood pressure values correlate poorly with direct blood pressure measurements in reptiles.48 Electrocardiography can be used to measure heart rate and assess rhythm, but does not provide information

about cardiac output; therefore it should not be used alone to assess cardiac function in anesthetized reptiles.49 A pulse oximeter indirectly estimates arterial oxygen-hemoglobin saturation and is calibrated based on the oxygenhemoglobin dissociation curve in humans. Reflectance probes (e.g., mammalian rectal probe) are more reliable in reptiles and can be placed within the esophagus or cloaca. In green iguanas, a pulse oximeter placed in the esophagus provided accurate oxygen hemoglobin saturation and pulse rate.43 Therefore, although data generated from pulse oximeters in reptiles should be evaluated with caution, trends in oxygen saturation are useful. Respiratory Monitoring Respiratory movements of the body wall in snakes and lizards can be easily visualized and measured in breaths/minute. In chelonians, movement of the skin overlying the prefemoral fossae and at each side of the thoracic inlet can be observed and measured during the respiratory cycle. Blood gas analysis provides information regarding oxygenation (PaO2), acid-base status (pH), and ventilatory status (PaCO2). Arterial blood gas analysis is not practical, because a cut-down procedure is required to access any artery (e.g., carotid, femoral). The introduction of the iSTAT, a portable clinical analyzer (Heska, Waukesha, WI USA), has made blood gas evaluation more practical for clinicians.50 As with pulse oximeters, these point-of-care analyzers have been validated for humans at standard body temperature; therefore absolute values in a nonendotherm may not be accurate, but trends are of use to the clinician. Venous blood gases can be useful if measured at several time points during a procedure to measure trends. However, a single venous PO2 is obviously going to be low and not reflective of actual arterial PO2. Increases in PCO2 do not necessarily reflect insufficient ventilation, but can also be the result of reduced tissue perfusion or increased metabolism.17 Capnographs estimate CO2 concentrations in expired gases and can be quite useful in mammals. However, multiple sampling problems exist when applied to reptiles, and capnography is still considered inaccurate in reptile species because reptiles can develop cardiac shunts. In green iguanas, there was no correlation between end-tidal CO2 concentrations and arterial PCO2 values.44 Monitoring Body Temperature Body temperature can be measured with a standard rectal thermometer placed in the cloaca or by

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CONCLUSION

FIGURE 16. Red-eared slider turtle recovering from anesthesia after surgical shell fracture stabilization and esophageal feeding tube placement. Note the IV catheter placed in the left jugular vein, which permitted peri-anesthetic fluid therapy in this hemodynamically compromised patient.

means of an esophageal temperature probe. Hypothermia will significantly delay recovery from anesthesia, and therefore reptiles should be maintained at a species-specific preferred optimum body temperature during the peri-anesthetic period. Recovery To facilitate recovery from general anesthesia, the concentration of the gas anesthetic agent should be gradually reduced toward the end of a surgical procedure and one should consider antagonizing any benzodiazepines and alpha-2-adrenergic agonists. Recovery from anesthesia can be best accomplished by placing the reptile in a temperature-controlled environment, such as an incubator, and periodically assessing muscle tone and reflexes. Spontaneous breathing is a reliable indicator of anesthetic recovery. Reptiles should be extubated once spontaneous breathing is observed and oropharyngeal reflexes (e.g., jaw tone, tongue movement) have returned.16 If spontaneous breathing has not returned, an Ambu bag should be attached to the endotracheal tube and IPPV performed (Fig 15). Room air for ventilation during recovery is preferred over oxygen, because high oxygen concentrations in the lungs have been shown to significantly delay the return of spontaneous respiration in recovering green iguanas.16 Fluid therapy and pain management should be continued during the recovery phase (Fig 16). 3 0

Although our understanding of reptile anesthesia has advanced significantly during the past decade, our ability to extrapolate across orders and species remains a major limitation. In other words, an effective anesthetic drug administered to a red-eared slider turtle may not have the same efficacy in a corn snake. Our inability to have a “one-size-fitsall” model for reptile anesthetics stems from the fact that the class Reptilia is incredibly diverse. Continued research focusing on effective anesthetic induction and maintenance drugs, dose-dependent effects, duration of drug efficacy, interspecies’ differences, and potentially detrimental drug-related adverse effects will advance the field and begin to reinforce, or abolish, the perpetuation of anecdotal information, which remains pervasive in the reptile clinical medicine literature. REFERENCES 1. Preston DL, Mosley CAE, Mason RT: Sources of variability in recovery time from methohexital sodium anesthesia in snakes. Copeia 2010:496-501, 2010 2. Carregaro AB, Cruz ML, Cherubini AL, et al: Influence of body temperature on rattlesnakes (Crotalus durissus) anesthetized with ketamine. Pesqui Vet Bras 29:969-973, 2009 3. Stirl R, Bonath KH: Anesthesia of tropical boids (Boa constrictor)—influences of the ambient temperature? Kleintierpraxis 43:839-845, 1998 4. Varga M: Captive maintenance and welfare, in Girling SJ, Raiti P (eds): BSAVA Manual of Reptiles (ed 2). Glouchester, UK, BSAVA Press, pp 6-17, 2004 5. Mosley CA: Anesthesia and analgesia in reptiles. Semin Avian Exot Pet Med 14:243-262, 2005 6. Hicks JW: The physiological and evolutionary significance of cardiovascular shunting patterns in reptiles. News Physiol Sci 17:241-245, 2002 7. O’Malley B: General anatomy and physiology of reptiles, in O’Malley B (ed): Clinical Anatomy and Physiology of Exotic Species: Structure and Function of Mammals, Birds, Reptiles and Amphibians. London, UK, Elsevier, pp 17-39, 2005 8. Holz P, Barker IK, Burger JP, et al: The effect of the renal portal system on pharmacokinetic parameters in the redfared slider (Trachemys scripta elegans). J Zoo Wildl Med 28:386-393, 1997 9. Holz P, Barker IK, Crawshaw GJ, et al: The anatomy and perfusion of the renal portal system in the red-eared slider (Trachemys scripta elegans). J Zoo Wildl Med 28:378-385, 1997 10. Coulthard P: Conscious sedation guidance. Evid Based Dent 7:90-91, 2006 11. Bienzle D, Boyd CJ: Sedative effects of ketamine and midazolam in snapping turtles (Chelydra serpentina). J Zoo Wildl Med 23:201-204, 1992 12. Oppenheim YC, Moon PF: Sedative effects of midazolam in red-eared slider turtles (Trachemys scripta elegans). J Zoo Wildl Med 26:409-413, 1995 13. Quagliatto Santos AL, Luz Hirano LQ, Pereira PC, et al: Anaesthesia of geoffroy’s side-necked turtle Phrynops geoffroanus Schweigger, 1812 (Testudines) with the association

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