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AVA/ECVAA Meeting Leipzig 2007 69

Training Day Avian and Reptile Anaesthesia and Analgesia Avian Anatomy and Physiology Dr. Conny Gunkel, Dr.med.vet, DACVA Oregon State University, Corvallis, OR Abstract Anesthesia is an important and challenging aspect of avian medicine for clinicians. Birds have anatomical and physiological features that differ from other species and have an important impact on anesthesia. Knowledge and understanding of the characteristics of the cardiorespiratory system of birds is important for the adequate selection and administration of anesthetic drugs in order to provide safe anesthesia. Extensive reviews of avian cardiorespiratory anatomy and physiology are available in the literature. Most important anatomical and physiological differences are summarized here with emphasis on the ones with clinical relevance. Anatomical and physiological differences: Cardiovascular system: The cardiovascular system is considered a “high performance” system to meet some difficult expectations like swimming, running, diving and flying at times under harsh circumstances. The high levels of activity demand high efficiency from the cardiovascular system to provide adequate delivery of oxygen to the tissues and removal of metabolic products. The cardiovascular system also plays an important role in conserving body heat. The avian heart is four chambered, located in the cranial part of the common thoracoabdominal cavity, slightly to the right of the midline. A very thin fibrous pericardial sac encloses the heart and contains a small volume of serous fluid. This pericardium is loosely attached to the dorsal surface of the sternum and the surrounding air sacs and more firmly to the liver. The pericardial membrane is relative non-compliant, which may lead to a low tolerance towards an increase in heart size due to for example volume overload. Birds have a proportionally larger heart in comparison to mammals, with smaller species containing a bigger heart of a given body mass than larger species. The anatomy of the heart itself is very similar to the mammalian heart, with a larger right atrium and thicker left ventricle wall diameter, but differences in myocardial structures are present. The AV valves show slight differences; the left AV-valve is tricuspid and the right AV valve consists of a single spiral flap of myocardium attached oblique to the free wall of the right ventricle. This flap is apposed to a downward extension of the free wall of the RA. The mechanism of valve closure at the start of ventricular systole is unknown and is considered to be either active by contraction of the muscular flap or passive by a brief backflow of blood at the start of ventricular systole. The aortic outflow valve shows a myocardial ring around the very rigid semilunar cusps. The coronary circulation happens through the right and the left coronary artery with slight species variation in branching. The rate of perfusion of the myocardium is much higher than for example the resting skeletal muscle and is highest during diastole. A reduction in oxygen supply or an increase in myocardial demand results in compensatory increase in coronary blood flow (high altitude flight). In comparison to mammals, birds don’t seem to show coronary changes in resistance and flow at hypocapnic conditions. Cardiomyopathies have been shown in turkeys and chicken (“round heart disease”). Functionally birds have with their larger heart also a larger stroke volume, greater cardiac output, higher blood pressures, and a lower heart rate compared to mammals. The cardiac conducting system consists of the SA node, AV node, Purkinje ring, His bundle and 3 bundle branches of Purkinje cells. The Purkinje fiber system is different at least in the ventricles to allow for increased speed of electrical excitation. The Purkinje fibers are bigger sized and lack a fibrous sheath and follow the coronary arteries through the myocardium. ECG: Lead I: Connects across the thorax from the right (negative electrode) to the left (positive electrode) wing base. Lead II: Right wing base (negative) to the left thigh (positive). Lead III: left wing base (negative) to left thigh (positive). This arrangement will show a negative polarity of the ventricular contraction. The vascular system in birds is also adapted with thicker vessel walls and lower elasticity. Capillary gas exchange has to adapt especially due to high workload of the pectoral muscle during flight (increase in O2 consumption of about 5 times that at rest) and the high altitude flights of some species.

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An increase in capillary numbers per muscle fibers and a decrease in diffusion distance might be possible explanations. Capillary branches increase the area of interface, which results in the very efficient blood-tissue O2 transfer. This also compensates for unfavourable rheological properties of avian blood like relatively low red cell deformability and low capillary hematocrit. Changes in P50, oxygen-carrying capacity of blood and/or myoglobin content of muscle fibers also facilitate oxygen delivery especially at high altitude. Fluid exchange mechanisms present some unique features in birds: The ratio of protein concentration in the interstitial fluid to that in blood is much lower than in mammals and is correlated with higher arterial blood pressure. Birds also are highly tolerant to blood loss, although the tolerance becomes more apparent during prolonged bleeding, because hemodilution continues through the period of blood loss. This hemodilution is achieved by the inflow of isotonic fluid with low protein content. The restoration of blood volume results from absorption of tissue fluid across the capillary walls due to reduced capillary pressure. Avian blood differs from mammalian blood in large oval and nucleated erythrocytes, large and nucleated thrombocytes, high glucose levels and low protein levels. The presence of the nucleus in the erythrocyte may lead to a high internal viscosity resulting in a decreased deformability of the cell. The adaptation for high altitude is a balance between polycythemia (in birds that are used to low altitude, leading to higher viscosity) and changes in hemoglobin oxygen affinity (increase of hemoglobin concentration without increase in erythrocytes and changes of P50 in birds that are used to high altitudes) and is dependent on species and environmental circumstances. Circulating catecholamines have strong effects on all elements of circulation in birds. Resting levels of especially norepinephrine are present and significant increased release of catecholamines, as it occurs under stress situations, may have greater impact on birds during anesthesia. Vasoconstriction and positive inotropic and chronotropic activation is the result with norepinephrine being more effective in the heart than epinephrine. The heart is intensely innervated with adrenergic and cholinergic nerve fibers (RA > LA > ventricles). Respiratory system: Differences between the avian and mammalian respiratory system are marked. The upper respiratory system includes the oronasal cavity, larynx, trachea with syrinx and primary bronchi. Birds can breathe through nares or mouth. The nares vary significant in structure and location between the different species. A complex bilateral infraorbital sinus is present throughout the skull, including the beak, with the main function to decrease the weight of the animals head. The larynx opens from the base of the tongue into the trachea through a slit-like glottis (rima glottis). An epiglottis is not present. The trachea has complete cartilaginous tracheal rings in most avian species. Interesting exceptions are the “double trachea” in penguins and the characteristic opening on the ventral side of the trachea in emus. Other species show tracheal elongations with loops. In general the avian trachea is typically 2.7 times longer and 1.3 times wider than their mammalian counterparts, (tracheal volume = 4.5 times larger). Dead space of the respiratory system (trachea and primary bronchi) is also increased (approximately 4 times), but a larger tidal volume and a lower respiratory frequency compensate for this difference. The trachea starts midline, then passes to the right side of the neck before it enters the thoracic inlet midline again, where it bifurcates at the level of the syrinx (vocal organ), into two primary bronchi. The small avian lungs are paired, attached firmly to the dorsal ribs, do not change volume during breathing and are located dorsally in the thoracoabdominal cavity. As part of the conducting airway the primary bronchi starts as a short extrapulmonary bronchi between the syrinx and the lung. The intrapulmonary part of the primary bronchi travels the entire length of the lung. Four groups of secondary bronchi arise from the primary bronchi and are named based on the origin: mediodorsal (6-10, caudal group), medioventral (4-5, cranial group), laterodorsal and lateroventral. The mediodorsal and medioventral groups branch and form fans to cover the lung surface. The primary and secondary bronchi are conducting airways and do not participate in gas exchange. The “parabronchi” or “tertiary bronchi” originate from the secondary bronchi and are the functional gas exchanging unit of the avian lungs. Most of the parabronchi are organized as a parallel series of hundred tubes connecting the medioventral and mediodorsal secondary bronchi (= paleo-pulmonic parabronchi). All birds (except some penguins) have also additional parabronchi (neo-pulmonic parabronchi), which contribute to gas exchange but with an irregular branching pattern. These parabronchi never compose more than 25% of the paleopulmonic bronchi and show large species variations. Oxygen and carbon dioxide exchange is very efficient in birds in comparison to mammals. Despite the fact that the actual volume of air involved in gas exchange (in air capillaries) in the avian lung is considerable less than the volume in mammalian alveolar lung, the unique pattern of air-flow through the parabronchi renews this gas-exchange volume more frequently, so that a large FRC is not necessary. Most birds have 9 air sacs (4 paired, 1 unpaired). The air sacs are thin membranous and

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avascular structures connected to the primary or secondary bronchi via ostia. As such they do not contribute to gas exchange, but do contribute to the effective respiration cycle by acting as bellows to ventilate the lungs, by following the changes in body volume with the respiratory activity generated by the respiratory muscles. The nine air sacs can be divided into 2 functional groups: the cranial and caudal groups. The cranial group includes the paired cervical air sac, the unpaired clavicular air sac and the paired cranial thoracic air sacs. The caudal group consists of the paired caudal thoracic and the paired abdominal air sacs. No diaphragm is present to separate thoracic from abdominal cavity. A typical change in thoracic skeleton between normal inspiration and expiration is present: during inspiration the sternum moves cranially and ventrally with the coracoids and furcula rotating at the shoulder. Simultaneously, the vertebral ribs move cranially to expand the sternal ribs and thoracoabdominal cavity laterally. During flight this movement is supported by the movement of the wings, but at rest inspiration and expiration requite active contraction of the respiratory muscles (i.e.: M. scalenus, Mm. intercostalis externi/interni, Mm costosternalis pars major/minor, Mm. levatores costarum, M. serratus, M. obliquus externus/internus abdominis, M. transversus abdominus, M. rectus abdominus, M. costoseptalis). Airflow during inspiration and expiration is consitent in most species. There is a unidirectional flow (caudal to cranial) through the paleopulmonic parabronchi. At inspiration, most of the TV goes into the caudal air sacs. From the caudal air sacs it flows in this unidirectional pattern through the paleopulmonic lungs and enters the cranial air sacs via the cranial secondary bronchi. At expiration the air leaves the cranial air sacs via the secondary bronchi emptying into the primary bronchi and leaving through the trachea. “Arodynamic valving” is responsible for the unidirectional flow, no anatomical valves are evident. This way the cranial air sacs receive air almost only from the parabronchi, which makes the PO2 and PCO2 almost similar to end-expired values. The caudal air sacs also have higher O2 and lower CO2 levels, because they contain a mixture of reinhaled dead space gas and fresh air. Also the neopulmonic parabronchi will contribute to gas exchange. In summery the respiration system in birds behaves is like a two-cycle pump: During the first inspiration, air passes almost completely into the caudal air sacs. In the first expiration, air moves into the lungs. On the second inspiration, air is drawn from the lungs into the cranial air sacs. In the second expiration, air is exhaled to the outside. Pulmonary circulation: Pulmonary arteries > interparabronchial arteries > intraparabronchial arteries > pulmonary blood capillaries > intraparabronchial veins > interparabronchial veins. All of the parabronchi are perfused along their entire length by oxygen-poor mixed venous blood and the oxygenated blood returning to the heart in the pulmonary veins is a mixture of blood draining the entire length of the parabronchi. This allows for cross-current gas exchange system. The air flow is assumed continuous through the parabronchus which is uniformly perfused along its length by mixed venous blood. At the inspiratory end of the parabronchus is a large PO2 gradient driving diffusion of O2

into the capillary blood,which raises capillary PO2 and drops parabronchial P02. As air flows along the parabronchus, the PO2 gradient decreases, while PvO2 is constant. At the end arterial PO2 is greater than the end-expired PO2. Same counts for the CO2: expired PCO2 can exceed arterial PCO2. P50: In general, the P50 in avian blood is greater than in mammalian blood. The theory is that because of the efficiency of O2 uptake in the avian lung is greater, birds may have evolved blood with low O2

affinity to maximize O2 delivery to the tissue. Birds used to high altitude seem to have a lower P50. The primary organig phonsphate in birds is Myinositol 1,3,4,5,6-pentophosphate (IPP) and not 2,3-DPG. In birds there is no independent effect of CO2 on P50, which is due to strong binding of IPP to hemoglobin in most birds, which prevents the CO2 Bohr effect. The avian lung is a complex structure consisting of hundreds of parabronchi. Mismatching of ventilation and perfusion can change the efficacy of gas exchange. Physiological dead space is also present in birds, shunting of pulmonary blood flow is very small, and V/Q mismatch can have a significant impact on the PaO2 in birds. Another difference in birds is the presence of intrapulmonary chemoreceptors (IPC) with high sensitivity to changes in PCO2. Blood gas measurements: Avian blood presents some challenges to accurate blood gas measurements. Because avian erythrocytes are nucleated an increased oxygen consumption can occur when the arterial blood is not analyzed immediately. Some discussion is present about the importance of applying a temperature correction factor and a blood gas correction factor established with a tonometer.

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Pneumatic bones: Many of the bones, including those in the skull, are pneumatized (sternum, scapula, humerus, femur (head and body), pelvis, cervical & thoracic vertebrae), which results in light, air-filled bones as an adaptation for flight. Bone marrow cells still exist in these bones, where red blood cells are produced. A bird's skeleton comprises only about 5% of its total body weight. The vertebrae of the lumbar, sacral and coccygeal spine are fused to form the “synsacrum” (with species dependent differences in numbers of vertebrates). Renal portal system: Renal portal circulation is present when venous blood that is flowing back to the heart from the legs and lower intestine of birds is entering the kidneys through a renal portal system. Within the kidneys this blood mixes with postglomerular efferent arteriolar blood in peritubular sinuses that surround all nonmedullary nephron segments and eventually flows toward the renal veins. There are 3 potential parallel shunt pathways in the renal portal circulation, depending on the parasympathetic and sympathetic innervations of the renal portal valves within the iliac veins. 1. If the renal portal valve is open (sympathetic NS activation), blood can flow through the valve into the common iliac vein leading to the vena cava and directly back to the heart, bypassing the kidney 2. If the valve is partially closed (parasympathetic stimulation) and resistance at the valve is high, blood can alternatively enter the renal portal system by flowing into the cranial or caudal portal veins, which are arranged in parallel. a. It enters the renal system via the cranial portal vein b. Or enters the renal and hepatic system via the caudal portal vein and caudal mesenteric veins The physiological significance is still poorly understood and neural, humoral and local metabolic controls may have influence on the valve resistance. Main advantage is for birds to be able to maintain total renal blood flow at lower arterial blood pressures (40-50 mmHg) and therefore preserve a wider range of renal autoregulation. This is important especially for salt loading and dehydration during the long flights or in desert environment.

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Avian Anaesthesia and Analgesia Dr. Conny Gunkel, Dr.med.vet, DACVA Oregon State University, Corvallis, OR Abstract: Avian patients often require anesthesia for diagnostic or therapeutic purposes. It seems that a high mortality rate makes avian anesthesia a feared procedure for any anesthetist, especially when it isn’t a routine anesthetic event. Knowledge of the avian cardiopulmonary function is important to make the anesthesia a less stressful event with a successful outcome. Knowledge about the different effects and pharmacokinetics of injectable drugs also prevent surprises. Well organized preparation, a smooth inhalation anesthesia and some analgesia remain the key to a good anesthetic and surgical outcome in birds. Pre-anesthetic evaluation Ideally, in a scheduled anesthetic procedure, a pre-anesthetic physical examination should be completed a day early. Body weight, baseline vital signs, and blood evaluation (CBC, hematocrit, total protein, glucose, uric acid, AST, calcium, phosphorus, and CPK) would provide useful information for correct dosing and drug selection. If a pre-anesthetic examination is not possible to perform, data can be obtained during the anesthetic event. Data analysis should be done in a timely manner to facilitate treatment in the anesthetized bird. Blood samples can be obtained from the jugular vein (right side is larger in psittacines), basilic vein (located on the ventral aspect of the wing, in the elbow area, or the medial metatarsal vein (especially useful in waterfowl). The jugular vein is the usual site for phlebotomy in psittacines, as it is easily visualized under a featherless area of the skin (apterium) and is less prone to hematomas compared to the basilic vein. As a general rule, the amount of blood volume taken from a bird should not exceed 1% of their body weight. This amount is usually not associated with adverse side effects in a healthy animal; however caution is advised in cases of anemia, hypovolemia or dehydration. In these conditions, it is safer to collect a maximum of 0.5% of the bird’s body weight. The method of blood collection and sample handling are important for obtaining reliable results, with hemolysis, clotting, and over-dilution with heparin being the most common problems. Fasting Opinions on the fasting time in birds are varied and controversial, and depend partly on the clinical status, size, and species of the bird to be anesthetized. Main indications for fasting include an increased risk for regurgitation and aspiration with a full crop, and a decreased efficiency of ventilation due to a full gastrointestinal tract impeding movement of air through the air sacs. If needed, a crop flush can be done to decrease the crop volume. The procedure involves passing a red rubber catheter through the esophagus into the crop for infusion and aspiration of warmed saline as the crop is gently massaged to soften the food material present. Due to the high metabolic rate and poor hepatic glycogen storage, a higher risk of hypoglycemia exists in smaller birds. For these reasons, fasting time should not exceed 6 hours. Often a fasting time between 2 and 4 hours is recommended in medium-sized species, while birds under 200 g may not need fasting. Anesthesia All phases of anesthesia (premedication, induction, maintenance, and recovery) are very critical in bird anesthesia. Complications during these phases are common and prevention is important to avoid mishaps. Premedication The restraint of the bird requires some expertise to ensure a safe and stress-reduced experience for the bird, handler and clinician. Premedication is rarely used in avian medicine to avoid repeated episodes of manual restraint. Nevertheless, use of premedication drugs for sedation can be advantageous in anxious, frightened, or excited birds. In addition, these drugs may result in a sparing effect on the amount of inhalant anesthetic drugs needed, hence reducing the depressive cardiovascular side effects (arrhythmogenicity, hypotension) of inhalant anesthetics. Benzodiazepines (midazolam, diazepam) and opioids (butorphanol) are most commonly used. Parasympatholytics (atropine, glycopyrrolate) are only used in patients with a history of bradyarrhymias. The routine use of parasympatholitics may result in thickening of tracheobronchial secretions and saliva, which leads to an increased risk of airway obstruction, especially in smaller bird species and is therefore not recommended.

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Induction In preparation for general anesthesia, anesthetic equipment and heating devices must be prepared. Drug doses for emergency drugs should be calculated before starting the procedure and ideally drawn up in advance. The high level of excitement of the patient associated with restraint during induction can predispose to cardiac arrhythmias (tachyarrhythmias) during this critical period. After drug administration, bradycardia and apnea can be common and lead to respiratory arrest and subsequently cardiac arrest. Induction should be done in a quiet and light-subdued environment. Since birds lack a diaphragm, respiration should not be compromised by forceful restraint that prevents the bird from having normal thoracic movement for adequate ventilation and oxygenation. Experience with bird holding is necessary to avoid trauma, bites and excessive stress on the bird. Catching the bird with a towel or a net is a commonly used technique, after which the bird can be held through the towel or with bare hands for induction. At all times, the holder should have control of the head, wings and feet. Maintenance During maintenance it is important to position the bird adequately for the surgical or diagnostic procedure, to avoid impairing cardiorespiratory function and to facilitate monitoring. Normothermia should be maintained as hypothermia induces bradycardia and hypotension. In addition, it delays drug metabolism and clearance. Most anesthetic complications occur following prolonged anesthetic times. Maintenance of anesthesia is usually accomplished with use of inhalation anesthetics. For short procedures, use of injectable drugs is feasible using additional doses of the induction drugs. However, concerns on drug accumulation and rougher recoveries make inhalation anesthetics the preferred method. For some injectable drugs, the use of antagonists may represent an alternative to overcome prolonged recoveries. Anesthetic monitoring follows the principles of mammalian anesthesia. Recovery Recovery is a critical period in avian anesthesia. As for induction and maintenance phases, monitoring is important to prevent cardiorespiratory depression from the restraint. Supplemental oxygen can be given to the recovering bird through the ET tube or by placing the face mask directly in front of the bird’s face. Positioning of the patient is very important, and the clinician must verify that the bird can breathe without difficulty. It is common practice for some practitioners to hasten recovery by side-to-side rocking; however, these fast turns may lead to hemodynamic changes, which may be detrimental and are therefore not recommended. Extubation should occur when the bird is fully awake, breathing well and able to swallow. Examination of the glottis prior to and immediately after extubation is crucial to avoid obstruction from secretions. If regurgitation occurs, the bird’s head should be lowered while the secretions are removed with cotton-tip applicators or gauzes. Use of suction is rarely necessary. The avian patient is held until it is mildly sedated, at which point it is returned to a cage or kennel and in a quiet and warm environment if possible. Small padded cages or incubator and towels wrapped around the bird assure this and might help keep then temperature in normal ranges. Hypothermia and hyperthermia should be avoided, as well as drastic changes in body temperature while warming the patient. Due to the risk of hypoglycemia it is necessary to ensure that patients eat soon after recovery, especially with the smaller species. Oral administration of a few drops of 50% dextrose can be performed during recovery if hypoglycemia is suspected. Adequate pain management will likely facilitate the recovery period. Anesthetic drugs: Inhalation anesthetics Use of inhalation anesthetics is the preferred method for induction and maintenance of anesthesia in birds. Advantages of using inhalation anesthetics for these phases include rapid induction and recovery, ability to make rapid and frequent adjustments in anesthetic depth, minimal biotransformation, minimal cardiorespiratory side effects or organ toxicity at clinical useful doses. These qualities make inhalation anesthetics ideal for birds with liver and/or kidney-altered function. Values for MAC are similar in birds and mammals. The cardiorespiratory depression of inhalation anesthetics is dose-dependant. Respiratory depression seems to be more significant in birds than in mammals. A decrease in respiratory rate and/or apnea is observed earlier in birds than in mammals. Because birds rely on thoracic muscles for ventilation, during anesthesia the relaxed muscles are not as efficient, resulting in a decrease in tidal volume and less efficient CO2 elimination. Birds possess a highly efficient gas exchange mechanism and their anesthetic depth can rapidly vary with changes in the anesthetic concentration. The concentration of inhalation anesthetics at which apnea ensues is known as the anesthetic index (AI). Because MAC and AI lay in a close range, assisted or controlled ventilation is often recommended. It also emphasizes the need for close anesthetic depth monitoring.

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Isoflurane and sevoflurane are the most common inhalation anesthetics used for bird anesthesia. Halothane is rarely used nowadays, due to its higher lipid solubility, biotransformation, and cardiorespiratory depression. For example, in Galahs, halothane was shown to cause more hypothermia, hypercapnia and electrocardiographic abnormalities than isoflurane. Similarly in ducks, increasing MAC multiples of halothane cause more cardiorespiratory depression than isoflurane. Sevoflurane was evaluated in chickens, pigeons and psittacines. In pigeons, induction and recovery times were shorter with sevoflurane compared to isoflurane. A study comparing sevoflurane to isoflurane in psittacines showed no differences between heart rate and respiratory rate during anesthesia and recovery time between the two groups; however, the birds anesthetized with sevoflurane became alert sooner and had less ataxia than the ones anesthetized with isoflurane. Sevoflurane it thought to have less irritant and repulsive odor in comparison to isoflurane and was shown to have less breath withholding (and thus a quicker induction time) in humans, other mammals and reptiles. It is unknown if this has any clinical impact in birds. The more controlled recovery exhibit with sevoflurane may give some advantage to this drug. No studies reporting the effects of desflurane in birds could be found in the literature. Anesthetic drugs: injectable drugs Injectable drugs are preferred in conditions of field work or where inhalation anesthesia is not readily available. The advantages of using injectable drugs in the avian patient however, rarely outweigh their disadvantages. These include dose-dependant cardiovascular side effects, interand intra-species effect variability, inability to reverse the drug and thus dependence on drug biotransformation and clearance, some potential prolonged and rough recoveries, need for an accurate weight (which may not be possible to obtain in field settings), and the narrow margin of safety for an appropriate anesthetic concentration. Injectable drugs reportedly used in birds include propofol, ketamine, and ketamine combinations with benzodiazepines, opioids or alpha-2 agonists. Tiletamine-zolazepam and alphaxalonealphadolone have been used in avian species with poor results. Propofol Propofol has been evaluated in several avian species including pigeons, barn owls, turkeys, chickens, mallard ducks, Canvasback ducks, ostriches, red-tailed hawks and Great horned owls, and Hispanolian parrots. Propofol was given IV between 3-15 mg/kg depending on the birds’ species and provided a smooth and rapid induction. Significant cardiopulmonary effects such as respiratory depression and hypotension can occur. Ventilation is therefore strongly recommended in birds anesthetized with propofol. Prolonged and/or stormy recoveries were reported in most studies, and were more significant in studies where CRI was used. Dosage of propofol during CRI was higher compared to that used in mammals (0.8- 1mg/kg/min in Hispaniolan parrots and Canvasback ducks versus 0.15 – 0.4 mg/kg/min in mammals). The observed longer recovery time suggests that some differences in phenolic metabolism may exist in birds as compared to mammals, and may allow for drug accumulation. The need of ventilation as well as prolonged and agitated recoveries decreases the benefits of using propofol as a sole anesthetic drug. As in mammals, the excitatory phase may be reduced by using a combination protocol of propofol plus a sedative and/or a muscle relaxant drug. This may provide some drug-sparing effect at induction and/or with a CRI. Dissociative drugs Ketamine is a phencyclidine anesthetic that causes dissociative anesthesia, and has been shown to have some analgesic properties in mammals. In birds, ketamine anesthesia requires a high dosage and is associated with poor muscle relaxation, muscle tremors, myotonic contractions, opisthotonus, and rough and prolonged recoveries. Moreover, in certain studies, even high dosages of ketamine were not sufficient to reach a surgical plane of anesthesia. For these reasons, it is not recommended to use ketamine as the sole anesthetic drug in birds. Another major disadvantage of using ketamine is its non-reversibility, which may result in prolonged recoveries, ranging from 40-100 minutes. Combinations using ketamine with benzodiazepines or alpha2-adrenergic agonists were used in attempt to improve relaxation, depth of anesthesia, and quality of recovery. The use of tiletamine-zolazepam has been reported in ducks, raptors and ostriches. It was primarily used in zoo and free ranging birds, or birds difficult to handle, however due to unacceptable prolonged and stormy recoveries (2-4 hours), its use is rare.

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Benzodiazepines Benzodiazepines have a sedative, anxiolytic, muscle relaxant, and MAC sparing effects. Their minimal cardiovascular effects and reversibility make the benzodiazepines ideal adjunct drugs to reduce the main induction and/or maintenance drug dosages. Diazepam is insoluble in water and is best administered IV only, to avoid pain or unreliable absorption when administered IM due to its carrier, propylene glycol. Midazolam, a water-soluble benzodiazepine, is preferred for IM administration. Flumazenil is benzodiazepine antagonist that can be administered in cases of overdose or prolonged recoveries. Flumazenil can be administered as a full bolus or titrated to effect; the latter method is preferred, to avoid reversing beneficial effects, such as anxiolysis, while muscle relaxation and sedation are reversed. Alpha-2 agonists Alpha-2 agonists were commonly used in combination with ketamine in birds before inhalation anesthesia was widely available. Xylazine was evaluated in avian patients but was found to provide unreliable anesthesia with severe cardiorespiratory depressant effects. Medetomidine, one of the most recent alpha-2 agonists, has the great advantage of providing profound analgesia and sedation in mammals and is reversible with atipamezole. Medetomidine has been evaluated in pigeons and Amazon parrots with and without midazolam or ketamine. The result of these studies showed that alpha2-agonists are not recommended as a short term anesthetics in these avian species due to their unreliable sedative effect, even at high doses, their inability to provide immobilization, their profound cardiovascular and respiratory side effects, and the fact that general excitement can effectively override their sedative effects. Induction: Face mask vs. intubation Face mask For a smooth induction with inhalation anesthetics and less risk of ambient pollution, a broad variety of masks with tight fits are needed, depending on the shape and size of the birds’ beaks. Due to the lack of commercially available masks of unusual shapes, custom made masks can be fabricated out of plastic water bottles and syringe cases to fit over the different shapes and length of beaks, making sure that the nares of the bird are included into the mask. In smaller species, the entire head of the bird can be placed into the face mask. A short time of pre-oxygenation is ideal, but the risks associated with prolonged time of physical restraint and excitement before being anesthetized often outweigh the benefits, such that preoxygenation is mainly used in respiratory compromised animals. Priming of the anesthetic circuit with the inhalant should be avoided because of the high exposure of gas waste pollution. There are two methods of anesthetic induction using inhalation anesthetics. One method involves incremental increases of the inhalation anesthetic over time (low-to-high-protocol). This method has the advantage of a reduced risk of overdose by allowing for rapid reversal, but has the disadvantage of a longer induction and excitement phase, which can be detrimental in a stressed and debilitated bird. However, a premedicated (hence sedated) bird may accept this technique well. The second method (high-to-low-protocol) is often the preferred method. This technique starts with a high percentage of inhalant (4-5% isoflurane or 6-8% sevoflurane in 1-2 L/min of oxygen) for induction, followed by maintenance of anesthesia (2-3% isoflurane or 4-5% sevoflurane), depending on the status of the animal and if any premedication was given. This technique requires a close observation of the animal and a timely decrease in anesthetic concentration delivered to avoid overdosing. The excitement phase with this method is much shorter, which generally makes it a ‘safe’ induction protocol, even for debilitated animals. When a light surgical plane of anesthesia is reached, the mask is removed and intubation with an endotracheal tube of appropriate size can be performed. Intubation Although some birds can be maintained on a face mask for short anesthetic procedures, in most cases intubation is recommended and is relatively easy to perform in birds. The avian glottis is located at the base of the tongue, and is usually easily visualized, due to the absence of an epiglottis. The beak is carefully opened with both hands or via gauze strips. A cotton-tip applicator may help exteriorize the tongue and give better access to the glottis. A mouth gag made of rolled gauzes and tape can be used to prevent damage to the tube. The advantages of intubation, even for short procedures, include the ability to provide manual ventilation, better control of anesthetic depth, and prevention of aspiration from food reflux. Because respiratory arrest can be followed quickly by cardiac arrest, it is easier to ventilate an already intubated bird. Moreover, endotracheal tubes allow the use of a capnograph as a monitoring tool during anesthesia. ndotracheal tubes used for intubation in birds should be uncuffed, due to the presence of complete tracheal rings and the fragility of the avian tracheal mucosa. Damage to the tracheal mucosa may be

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followed by a fibrotic healing process, which can narrow the tracheal lumen and cause respiratory complications. This may not become evident until 3-7 days post intubation. Uncuffed ET tubes come in a wide variety of sizes, the smallest being 1.0 mm internal diameter. Small tubes may need a stylet to help with intubation, while some other brands have a metal coil incorporated into the tube that provides rigidity and prevents kinking. For small birds, a very small ET tube, or alternatively a catheter sheath, rubber feeding tube, or urinary catheter can be used for intubation. However, birds weighing less than 80 g are usually not intubated, due to the fact that any of these small tubes can be easily clogged with mucous. This will decrease the size of the internal diameter even further and create a significant resistance to airflow. The use of cuffed ET and very minimal inflation of cuffs can be used in larger birds where an effective seal for assisted or controlled ventilation is important, or to prevent aspiration when a crop lavage is done. Maintenance: flow rate and ventilation Assisted or controlled ventilation is recommended in most birds, due to the respiratory depressant effects of most inhalation anesthetics. Respiratory rate of 10-25 breaths/min in larger species and 30-40 breaths/minute for smaller birds are recommended. Adequate ventilation is monitored by visualizing chest excursions and/or a capnograph. Peak inspiratory pressure should not exceed 5-15 cm of H2O, depending on size of the bird, to prevent trauma to the air sacs. Pediatric ventilators can be used in avian patients, but an alarm system for early detection of obstructions or changes in resistance is mandatory. Non-rebreathing circuits, like the Ayre’s T-piece or Bain’s coaxial are most commonly used in birds weighing less than 5-7 kg. Advantages of these systems include faster changes in the anesthetic gas concentration, lower dead space, less resistance, and their convenient lightweight. Disadvantages include a constant high use of oxygen and anesthetic gas, and cooling of the patient with the cold and dry fresh gas flow. The T-piece has an inconvenient position of the bag and pop-off valve and there is an increased risk of over inflation from accidentally closing the valve. This can lead to an increase in intracoelomic pressure and decreased venous return and consequently cardiac arrest. In birds weighing over 7 kg, a pediatric rebreathing circle system could be used, which may have the advantage of a better heat preservation compared to a non-rebreathing circuit, especially when low-medium flow rates are used (economical, ecological and heat-preserving advantages, but do require the monitoring of end-tidal CO2 ). Adding monitoring equipment, such as capnographs and respiration monitors, or long ETtubes will increase dead space, especially in smaller birds. The ET tube can either be cut shorter to fit the bird beak length or the capnograph could be placed intermittently. False low readings from the capnograph may be present due to a low TV, especially when a side-stream capnograph is used. Positioning The respiratory function in anesthetized birds can be significantly altered as a result of direct respiratory depressant effects of the anesthetic drug, muscle relaxation, increased dead space and positioning. Lacking a diaphragm, an enlarged gastro-intestinal tract or physical pressure from the surgeon’s hand can impede on the ventilation of the bird during anesthesia. Positioning of the bird will depend on the procedure to be performed, but if possible the bird should lay in lateral recumbency. Dorsal or ventral recumbency can impact the movement of the sternum and compress the abdominal air sacs, which can lead to a decrease of effective ventilation. Air sac intubation Air sac intubation is used as an emergency procedure in cases of upper airway obstruction, for example in tracheal foreign body, masses, and fungal granulomas. It can also be used during anesthesia as an alternative to endotracheal intubation, when free access to the head and upper respiratory tract is needed, for example during tracheoscopy, tracheotomy or ophthalmic procedures. Purpose-made avian air sac cannulas are available (Air sac surgical catheters, Global Veterinary Products, Inc.). These short tubes (2.5-3.6 cm long) have multiple ventilation holes and a convenient silicone disk for easy skin fixation using suture material. They also have a removable proximal fitting that accepts conventional anesthesia tubing. Their diameter is 14 or 20 French (3 and 4 mm internal diameter), and the tubes can be cuffed or uncuffed. Alternatively, an air sac cannula can be made of modified ET by cutting it shorter and creating supplemental holes in the tube in order to reduce chances of obstruction. Air sac cannulation can be performed on either side of the bird, in the caudal thoracic, abdominal, or cervical air sacs. The left caudal thoracic air sac is usually preferred because it is greater size and its approach is familiar to most clinicians who perform coelioscopy for gender determination. Clavicular air sac cannulation failed to provide effective ventilation or maintain anesthesia in Sulphur-crested cockatoos and is therefore not recommended.

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The procedure is best performed with the bird under anesthesia (except when used as an emergency procedure). Pain medication should be used and/or a local block performed. The bird is placed in right lateral recumbency, the left leg is pulled caudally, and the wings are pulled upwards as to reveal the left paralumbar fossa. This area is defined as a triangular area bordered caudally by the cranial thigh (femur), cranially by the last 2 ribs, and dorsally by the synsacrum. The area is plucked and aseptically prepared. A small incision (0.5-1.0 cm, long enough to pass the cannula) is made through the skin caudally to the last rib (or in certain cases between the last two ribs). A sterile hemostat is used to dissect the soft tissue, and with a quick and controlled stab, the coelomic cavity is bluntly penetrated, in a similar technique as used for a laparoscopy procedure. A loud popping noise is often heard upon penetration of the coelom. The hemostats are opened and the cannula is inserted into the underlying abdominal air sac (1-2 cm deep). The short tube is secured to the skin with non-absorbable sutures, by suturing the silicon disk present in purpose-made air sac cannulas or by using butterfly tape and sutures on the homemade ones. Purse-string sutures and a Chinese finger trap technique can also be used. Confirmation of proper placement and function can be done by observing condensation into the tube during breathing, or by placing a down feather in front of the tube and looking for its movement during breathing. The air sac tube can be used for inhalation anesthesia maintenance, using positive pressure to ventilate the bird. Due to the unusual location of the endotracheal tube, chest excursions are more difficult to observe. After recovery, the tube may be left for several days if necessary; its patency must be verified frequently. Before removing the tube, it is wise to first obliterate it and verify that the bird can still breathe well. Most birds do not tolerate their air sac cannulas and require an Elizabethan collar to prevent destruction or removal of the tube. Analgesia The difficult part about avian analgesia is the recognition of pain, appropriate treatment of the individual patient and evaluation of the effects of the treatment. Unfortunately only few studies have looked at the efficacy of the different analgesic agents in the various species of birds, mainly due to a lack of understanding of “pain behavior” in these different avian species. Nevertheless pain relief should always be provided from an ethical standpoint (birds have a very similar pain pathway than mammals), and to prevent negative outcomes due to increased anxiety and stress responses and decreased homeostasis and healing. Also the reduction of required anesthetics used by providing analgesia decreases the risk of cardiopulmonary side effects of the anesthetic agents. The principles of pain management apply in birds as they do in mammals. Preemptive and balanced analgesia are the main components to provide adequate analgesia and prevent peripheral and central sensitization. Opioids are frequently used in birds, but unfortunately inconsistent experiences and species specific differences in studies lead to controversial opinions about their use, effects and doses. A difference in opioid receptor population and distribution between birds and mammals may contribute to the difference in response. Studies have been done on butorphanol, morphine, buprenorphine and fentanyl with various results. Butorphanol and Buprenorphine remain the most studied opioids in birds and clinically the most popular analgesic agents for surgical procedures. NSAIDs are also commonly used in birds, often in combination with an opioid. The less obvious side effects (respiratory depression, behavior changes, sedation), make this group of drugs a good choice for field work (transmitter placement etc). Analgesic studies have been performed on chicken treated with carprofen and ketoprofen. More recently meloxicam is clinically used in birds, but still lacking more efficacy, safety and pharmacokinetic studies in the different avian species. Flunixin is considered nephrotoxic especially in cranes. The alpha2-adrenergic agonists show undesirable side effects in birds (muscle tremor, respiratory depression, auditory sensitivity etc), and are rarely used as an analgesic agent. Local anesthetics (lidocaine, bupivacaine) are an easy and efficient option to prevent the activation of the nociceptive pathway. If the anatomy allows for a local block (beak amputation, extremity surgery etc), a nerve-block should be performed, but accurate dosing is important especially in smaller birds. Birds may be more sensitive to the local anesthetics and an overdose can lead to cardiac arrest and seizures. Commonly a max dose of 4 mg/kg of lidocaine and 2 mg/kg of bupivacaine is recommended. Further clinical investigation and clinical assessment is needed to evaluate and improve analgesia options in the various avian species. In the mean time a progressive pain management approach in avian medicine is an important goal to improve patient welfare and clinical outcome as well as gain clinical experience with these drugs in the different avian species.

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Supportive therapy Fluid therapy- Crystalloids All birds that undergo a long anesthetic procedure such as surgery or diagnostic procedures should receive fluids. The fluids should be warmed to body temperature. Replacement fluids such as lactated ringers solution, plasmalyte 148, normosol during anesthesia for a surgery are given IV or intraosseously at 10-20 ml/kg/hr. This small amount of fluid can be given continuously using a syringe pump or in slow boluses every 5 to 10 minutes. Alternatively, subcutaneous fluid can be administered at 30 ml/kg once. This is a quick, easy, and helpful method and appears quite effective even in debilitated in birds. Intraosseous catheter The placement of an intraosseous (IO) catheter is an option for administration of fluids in dehydrated or hypovolemic patients, where venous access is difficult. Any types of fluids, medication or emergency drugs that can be given IV can be given IO and are absorbed as quickly. The most common sites for placing IO catheters include the ulna (proximal or distal), and proximal tibiotarsus. The avian humerus and femur are often pneumatized and connected to the respiratory system; therefore they are contraindicated locations for IO catheter placements. Drugs injected into the IO space are absorbed by sinusoids and drain into veins that connect into the systemic circulation. Purpose-made IO needles are available (Global Veterinary Products, Inc.). They range from 14 – 20 ga and are 3 cm in length. These needles have a metal stylet, a convenient handle to facilitate the introduction of the needle into the bone, and a plastic fixation device that can be sutured to the skin and help stabilizing the IO catheter. Alternatively, regular needles or spinal needles can be used. An appropriate size and length of needle is chosen for the size of the avian patient (18- 25 ga). A disadvantage of using regular needles is the possibility of a bone plug blockage, although this problem is rarely encountered. In small patients, regular needles are actually easier to use and more versatile in size than an IO needle, and can easily penetrate the thin cortex of the avian bones. To place a proximal ulnar IO catheter, the dorsal aspect of the ulna is plucked and the area is aseptically prepared. The ulna is held in one hand (usually the left for a right handed person), and the needle is held in the other hand (right) and positioned ventral to the condylar ridge of the distal ulna. With a firm and slight rotating movement similar to that of a retrograde pin placement, the needle is gently driven into the ulnar bone at a 45°-70° angle. Once the ulna is penetrate d, the clinician should feel a markedly reduced resistance. At that point, the needle angle is reduced as to be as parallel to the bone as possible and with a gentle rotating movement, the needle is completely driven into the medullary cavity of the bone. Correct placement of the IO catheter can be confirmed by taking a radiograph. If the needle is properly positioned, fluids will easily flow through the catheter and can be visualized passing through the ulnar vein. The catheter is capped with a luer-lock injection port and secured to the wing with tape and/or sutures. Gauzes can be used to protect the injection port, and the wing is then wrapped with a standard figure-of-8 bandage. When placing a proximal ulnar catheter, the point of entry should be three to four flight feathers proximal to the elbow. The ulna is penetrated at an acute angle with a rotating motion. Once the needle is inserted into the bone, the angle is reduced, and the needle is fully inserted into the ulnar medullary cavity with a gentle rotating movement. If placing a proximal tibiotarsus IO catheter, the stifle is flexed and the needle is inserted into the trochanteric fossa. As for an IV catheter, fluids can be given continuously or via boluses. The catheter can be left in place for several days. The choice of fluid type is determined by the status of the patient (PCV, TP, dehydration status) and the suspected blood loss during surgery. Due to the bird’s high metabolic rate and low glycogenstorage capacity, a regular assessment of glucose levels is recommended, but unfortunately rarely performed in clinical settings due to the small size of most avian patients. Intra-operative measurement of glucose should become a routine monitoring procedure for prolonged procedures, and could be performed with a glucometer. These devices require a minimal amount of blood, which would be possible to get even in small birds. Lactated ringers solution with added 2.5-5% dextrose is commonly used for replacement fluids in birds. Reference biochemistry values for the specific avian species are important for fluid choice. Isotonic and normoelectrolyte fluid therapy can be provided. Interspecies differences in electrolytes exists, and birds have generally lower potassium and higher sodium values compared to mammals. This difference may have an impact on physiological balances (fluid dynamics in the body, cardiac cycle) when using fluids that are formulated to be isotonic for mammals. This needs to be considered when choosing the type of fluid for fluid therapy. Careful monitoring of electrolytes in a sense of taking blood gas samples would be ideal, but often not clinically applicable.

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Colloids (Blood transfusion, Oxyglobin and Hetastarch) If severe blood loss occurs during surgery, blood transfusions, colloid solutions or hemoglobinbased products should be considered to expand the vascular space and improve the cardiac output. The pathophysiology of hemorrhagic shock in birds is still not fully understood, but signs of hypotension and tachycardia have been described in response to acute blood loss. Birds seem to be more tolerant to acute blood loss than mammals, due to their ability for prolonged hemodilution. Estimated circulating blood volumes in different avian species vary considerably from 5% of body weight in ring-necked pheasants up to 20% in racing pigeons. The acute blood loss of 60% of assumed total blood volume in ducks has been described as the LD50, compared to mammals, where the LD50 was reached with 40-50% loss of total blood volume. The administration of whole blood is optimal to restore tissue perfusion and oxygenation, but has some limitations due to the availability of donors. Blood types are currently unknown in birds, so cross matching is generally not performed. Homologous (of the same species) transfusions have the longest erythrocyte survival time, but heterologous (between different species) blood transfusion in avian species is often more feasible and has been documented without any significant side effects. Anticoagulants include acid citrate dextrose, sodium citrate, or citrate phosphate dextrose at 1-1.5 ml per 10 ml of blood. Heparin can also be used at 0.25 ml / 10 ml of blood. Another problem with transfusion of whole blood is the fact that the volume of blood to be given is often dictated by the donor bird rather than the recipient. The blood can be given IV or IO through a blood filter at a continuous rate of 2 ml/minute, or through multiple boluses over a few hours. Due to the limited availability of avian blood products, the use of other colloids are often the only source to treat acute hemorrhagic shock in birds and has been used in clinical and research settings. In comparison to mammals, avian total protein concentration is substantially lower (21- 45 g/L [2.1-4.5 g/dl]). Proteins are the major determinant of colloid osmotic pressure (COP) and may indirectly influence blood pressure. COP in birds is substantially lower compared to mammals (11 mmHg for chicken and 8.1 mmHg in doves, compared to 25 mmHg in mammals), but the ratio of protein concentration in the interstitial fluid to that in the blood is much lower as well. This lower ratio in birds may be correlated with a higher arterial blood pressure. Resting blood pressure values in awake birds has been reported to be higher than that of mammals and mean arterial blood pressure may exceed 150 mmHg in some species. The generally lower total protein value in birds is a point of consideration in the choice of fluid type and volume to administer, since most colloids COP is 20-25 mmHg. This may exceed the avian COP and may lead to fluid movement from the extravascular space to the intravascular space. This beneficial effect of volume expansion of the vessels could cause fluid overload. In addition, the hydration of the interstitial space needs to be guaranteed with the concurrent administration of crystalloids. The clinical relevance of this concern is not known. The use of hetastarch (HES, Abbott Laboratories, North Chicago, IL) for the management of hypoproteinemia and hypovolemia has been evaluated in birds. IV boluses of 10 ml/kg can be given to birds, but commonly colloids are administered over several minutes / hours via a syringe pump, in conjunction with crystalloids. In a study on cockatiels, hetastarch was given as an IV bolus with crystalloids between 1 and 15 ml/kg. The effect of hetastarch on platelet aggregation in birds has not been investigated. Oxyglobin is a purified polymerized bovine hemoglobin and has the advantage of being a potent colloid as well as having a major oxygen carrying capacity to provide delivery of oxygen to the tissues. Because it contains no antigen, cross matching is not required and no filters are needed for the administration. In birds, oxyglobin has been given as a rapid bolus over a few minutes with crystalloids between 1 and 15 ml/kg. Anesthetic complications With avian patients, it is wise to have common emergency drugs (e.g. epinephrine and atropine) drawn up and ready for injection, to counteract anesthetic complications. Respiratory arrest followed by cardiac arrest is the most common anesthetic avian complication. Respiratory arrest is often treatable if detected early. The first step includes reducing or turning off the inhalant anesthetic and/or using reversal drugs, and providing manual ventilation until return of spontaneous respiration. If the respiratory arrest progresses to a cardiac arrest, cardiac compressions can be attempted, but are difficult to perform in birds due to their sternum preventing access to the heart. Epinephrine can be given IV, IO, or intra-tracheally. Unfortunately, the success rate for the return of cardiac function in birds is low. Other anesthetic complications include cardiac arrhythmias, which can be observed in combination with pain, inappropriate depth of anesthesia, changes of plasma glucose levels, temperature, blood gas or electrolyte values, and blood loss. Hypothermia and blood loss can be important intraoperative complications. Hypothermia can lead to bradyarrythmias, which may be resistant to treatment with glycopyrrolate or atropine. Warm-forced air blankets are the most successful heating devices for preventing hypothermia in very small animals.

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Avian Reference List Anatomy and Physiology Dr. Conny Gunkel, Dr.med.vet, DACVA Oregon State University, Corvallis, OR 1. Abou-Madi N: Avian Anesthesia. Vet Clin North Am (Exotic Anim Pract) Analgesia and Anesthesia, 4:147- 167, 2001 2. Altman RB, Clubb SL, Dorrestein GM, et al (eds): Avian Medicine and Surgery, Philadelphia, PA, Saunders, 1997 3. Atalan G, Uzun M, Demerkan I, et al: Effect of medetomidine-butorphanol-ketamine anaesthesia and atipamezole on heart and respiratory rate and cloacal temperature of domestic pigeons. J Vet Med Assoc 49:281-285, 2002 4. Beart K, de Backer P: comparative pharmacokinetics of three non-steroidal antiinflammatory drugs in five bird species. Comp Biochem Physiol Part C 134:25-33, 2002 5. Campbell TW: Avian hematology, in Campbell TW (ed): Avian hematology and cytology. Ames, IA, Iowa State University Press, 1995. pp 54-59 6. Carp NZ, Saputelli J, Halbherr TC, et al: Technique for liver biopsy performed in Pekin ducks using anesthesia with Telazol. Lab Anim Sci 41:474-475, 1991 7. Carpenter JW, Mashima TY, Rupiper DJ (eds): Exotic animal formulary, (ed 2). Philadelphia, PA, Saunders, 2001 8. Christensen J, Fosse RT, Halvorsen OJ, et al: Comparison of various anesthetic regimens in the domestic fowl. Am J Vet Res 48:1649-1657, 1987 9. Concannon KT, Dodam JR, Hellyer PW: Influence of a mu- and kappa-opioid agonist on isoflurane minimal anesthetic concentration in chicken. Am J Vet Res 56:806-811, 1995 10. Corr SA, McCorquodale C, McDonald J, et al.: A force plate study of avian gait. J Biomech. 40(9):2037- 2043. 2007 11. Costello MF: Principles of cardiopulmonary cerebral resuscitation in Special species. Seminar in avian and exotic pet medicine, 13:132-141, 2004 12. Curro TG, Brunson DB, Paul-Murphy J: Determination of the ED50 of isoflurane and evaluation of the isoflurane-sparing effect of butorphanol in cockatoos. Vet Anesth 23:429-433, 1994 13. Curro TG: Anesthesia of pet birds. Seminar in avian and exotic pet medicine 7:10-21, 1998 14. Danbury TC, Chambers JP, Weeks CA, et al.: Self-selection of analgesic drugs by broiler chickens. Anim Choices 20: 126-127, 1997 15. Degernes LA, Crosier ML, Harrison LD, et al: Autologous, homologous, and heterologous red blood cell transfusions in cockatiels (Nymphicus hollandicus). J Avian Med Surg 13:2- 9, 1999 16. Degernes LA, Harrison LD, Smith DW, et al: Autologous, homologous, and heterologous red blood cell transfusions in conures of the genus Aratinga. J Avian Med Surg 13:10-14, 1999 17. De Lucas JJ, Rodriguez C, Marin M, et al.: Pharmacokinetics of intramuscular ketamine in young ostriches premedicated with romifidine. J Vet Med A Physiol Pathol Clin Med; 54(1):48-50, 2007. 18. Edling TM, Degernes LA, Flammer K, et al.: Capnographic monitoring of anesthetized African grey parrots receiving intermittent positive pressure ventilation. J Am Vet Med Assoc. 15;219(12):1714-8. 2001 19. Fowler ME, Miller RE (eds): Zoo and Wild animal medicine (ed 5). Philadelphia, PA, Saunders, 2003 20. Fitzgerald G, Cooper JE: Prelimiary studies on the use of propofol in the domestic pigeon (Columbia livia). Res Vet Sci 49:334-338, 1990 21. Gentle MJ: Pain in birds. Anim Welfare 1:235-247, 1992 22. Glatz PC, Murphy BL, Preston AP: Analgesia therapy of beak-trimmed chickens. Aust Vet J 69:18, 1992 23. Gleed RD, Ludders JW (eds): Recent Advances in Veterinary Anesthesia and Analgesia: Companion Animals, International Veterinary Information Service. Ithaca, NY, 2001 24. Harr KE: Clinical Chemistry of Companion Avian Species: A Review. Vet Clin Path 31:140-151, 2002 25. Hawkins MG, Wright BD, Pascoe PJ: Pharmacokinetics and anesthetic and cardiopulmonary effects of propofol in red-tailed hawks (Buteo jamaicensis) and great horned owls (Bubo birginiansus). Am J Vet Res 64:677-83, 2003 26. Heatley JJ, Marks S, Mitchell M, et al: Raptor emergency and critical care: Therapy and techniques. Compendium 23:561-570, 2001 27. Hocking PM, Bernard R, Maxwell MH: Assessment of pain during locomotion and the welfare of adult male turkeys with destructive cartilage loss of the hip joint. Br Poult Sci 40: 30-34, 1999 28. Hocking PM, Robertson GW, Gentle MJ: effects of anti-inflammatory steroid drugs on pain in domestic fowl. Res Vet Sci 71: 161-166, 2001

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29. Hocking PM, Robertson GW, Gentle MJ: Effects of non-steroidal anti-inflammatory drugs on pain-related behaviour in a model of articular pain in domestic fowl. Res vet Sci 78: 69-75, 2004 30. Hoppes S, Flammer K, Hoersch K, et al.: Disposition and analgesic effects of Fentanyl in white cockatoos. J Avian Med Surg 17: 124-130, 2003 31. Jaensch SM, Cullen L, Raidal SR: Comparative cardiopulmonary effects of halothane and isoflurane in Galahs (Eolophus rosweicapillus) J Avian Med Surg 13:15-55, 1999 32. Jaensch, SM, Cullen L, Raidal SR: Comparison of endotracheal, caudal thoracic air sac and clavicular air sac administration of isoflurane in Sulfur-crested cockatoos (Cacatua galerita). J Avian Med Surg 15:170-171, 2001 33. Jenkins JR: Post-opertaive care of the avian payient. Sem Avian Exotic Pet Med 2:97- 102,1993 34. Korbel R: Comparative investigations on inhalation anesthesia with isoflurane and sevoflurane in racing pigeons (Columba livia Gmel., 1789, var. domestica) and presentation of a reference anesthesia protocol for birds. Tierarztl Prax Ausg K Klientiere Heimtiere. 26:211-23, 1998 35. Kreeger TJ, Degernes LA, Kreeger JS, et al: Immobilization of raptors with tiletamine and zolazepam (telazol), in Redig PT, Cooper JE, Remple JD, et al (eds): Raptor biomedicine. Minneapolis, MN, University of Minnesota Press, 1993; pp 141-144 36. Langlan JN, Ramsay EC, Blackford JT, et al: Cardiopulmonary and sedative effects of intramuscular medetomidine-ketamine and intravenous propofol in ostriches (Strutio camelus) J Avian Med Surg 14:2- 7, 2000 37. Langlois I, Harvey CR, Jones MP, et al: Cardiopulmonary and anesthetic effects of isoflurane and propofol in Hispaniolan Amazon Parrots (Amazona ventralis). J Avian Med Surg 17:4-10, 2003 38. Lichtenberger M, Orcutt C, DeBehnke D, et al: Mortality and response to fluid resuscitation after acute blood loss in mallard ducks (Anas platyrhynchos) in: 2002 Scientific Proceedings, 23rd Annual Meeting of Association of Avian Veterinarians. Monterey, CA, 2002, pp 65-67 39. Lichtenberger MK, Chavez W, Cray C, et al: Mortality and response to fluid resuscitation after acute blood loss in mallard ducks, in: 2003 Scientific Proceedings, 24th Annual Meeting of Association of Avian Veterinarians. Pittsburg, PA, 2003, pp 7-10 40. Lichtenberger M: Transfusion medicine in exotic pets. Clin Tech Small Anim Pract. 19:88- 95, 2004 41. Lichtenberger MK, Rosenthal K, Brue R, et al: Administration of Oxyglobin and 6% hetastarch after acute blood loss in psittacines birds, in: 2001 Scientific Proceedings, 22nd Annual meeting of the Association of Avian Veterinarians. Orlando, FL, 2001, pp15-18 42. Lichtenberger MK Ko J: Anesthesia and analgesia for small mammals and birds. Vet Clin North Am Exotic Anim Pract. 10(2):293-315, 2007 43. Ludder JW, Mitchel JS, Rode J: Minimal anesthetic concentration and cardiopulmonary dose response of isoflurane in ducks. Vet Surg 19:304-307, 1990 44. Ludder JW: Minimal anesthetic concentration and cardiopulmonary dose response of halothane in ducks. Vet Surg 21:319-324, 1992 45. Lukasik VM, Gentz EJ, Erb HN, et al: Cardiopulmonary effects of propofol anesthesia in chickens (Gallus gallus domesticus) J Avian Med Surg 11: 93-97, 1997 46. Machin KL, Caulkett NA: Cardiopulmonary effects of propofol and a medetomidinemidazolam- ketamine combination in mallard ducks. Am J Vet Res. 59:598-602, 1998 47. Machin KL, Caulkett NA: Cardiopulmonary Effects of propofol infusion in canvasback ducks (Aythya valisineria). J Avian Med Surg 13: 167-172, 1999 48. Machin KL, Caulkett NA: Evaluation of isoflurane and propofol anesthesia for intraabdominal transmitter placement in nesting female canvasback ducks. J Wild Dis 36: 324-334, 2000 49. Machin KL, Tellier LA, Lair S, et al: Pharmacodynamics of flunixin and ketoprofen in mallard ducks. J Zoo Wildl Med 32: 222-229, 2001 50. Machin KL, Livingston A: Assessment of the analgesic effects of ketoprofen in ducks anesthetized with isoflurane. Am J Vet Res. 63: 821-826, 2002 51. Machin KL, Avian Analgesia. Sem Avian Exot Pet Med, 14 (4): 236-242, 2005 52. Machin KL. Controlling Avian Pain. Comp Cont Ed Pract Vet. 27:299-309, 2005 53. Mama KR, Phillips LG, Pascoe PJ: Use of propofol for induction and maintenance of anesthesia in a barn owl (Tyto alba) undergoing tracheal resection. J Zoo Wildl Med 27:397-401, 1996 54. McGeown D, Danbury TC, Waterman-Pearson AE, et al.: effects of carprofen on lameness in broiler chickens. Vet Rec 144: 668-671, 1999 55. Morissey J: Avian transfusion medicine. Exotic Pet Pract 4:65-66, 1999 56. Naganobu K, Fujisawa Y, Ohde H, et al: Determination of the minimum anesthetic concentration and cardiovascular dose response for sevoflurane in chickens during controlled ventilation. Vet Surg 29:102- 105, 2000 57. Naganobu K, Ise K, Miyamoto T, et al.: Sevoflurane anaesthesia in chickens during spontaneous and controlled ventilation. Vet Rec. 11;152(2):45-8, 2003

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58. Nilson PC, Teramitsu I, White SA: Caudal thoracic air sac cannulation in zebra finches for isoflurane anesthesia. J Neurosci Methods. 30;143(2):107-115. 2005 59. Paul-Murphy JR, Brunson DB, Miletic V: Analgesic effects of butorphanol and buprenorphine in conscious African grey parrots. Am J Vet Res 60:1218-1221, 1999 60. Paul-Murphy JR, Brunson DB, Miletic V: A technique for evaluating analgesia in conscious perching birds. Am J Vet Res.:6 0(10):1213-7. 1999 61. Paul-Murphy JR, Ludders JW: Avian analgesia. Vet Clin North Am Exot Pet Pract 4:35- 45, 2001 62. Paul-Murphy J, Hess JC, Fialkowski JP: Pharmacokinetic properties of a single intramuscular dose of buprenorphine in African grey parrots. J Avian Med Surg 18:224-228, 2004 63. Pollock CG, Schumacher J, Orosz SE: Sedative effects of medetomidine in pigeons (Columba livia) J Avian Med Surg 15:95-100, 2001 64. Quandt JE, Greenacre CB: Sevoflurane anesthesia in psittacines. J Zoo Wildl Med 30:308-309, 1999 65. Reim DA, Middleton CC: Use of butorphanol as an anesthetic adjunct in turkeys. Lab Anim Sci 45: 696- 697, 1995 66. Rembert MS, Smith JA, Hosgood G: Comparison of traditional thermal support devices with the forcedair warmer system in anesthetized Hispaniolan amazon parrots (Amazona ventralis) J Avian Med Surg 15:187-193, 2001 67. Ritchie BW, Harrison GJ, Harrison LR (eds): Avian medicine, principles and application. Philadelphia, PA, Saunders, 1997 68. Rode JA, Bartholow S, Ludders JW: Ventilation through air sac canula during tracheal obstruction in ducks. J Avian Assoc Vet 4:98-102, 1990 69. Rosskopf WJ, Woerpel RW, Reed S, et al: Avian anesthesia administration, in: 1989 Proceedings of the American Animal Hospital Association, St. Louis, MO, 1989, pp 449- 457 70. Samour JH, Jones DM, Knight JA: Comparative studies of the use of some injectable anesthetic agents in birds. Vet Rec 115:6-11, 1984 71. Sandmeier P: Evaluation of medetomidine for short-term immobilization of domestic pigeons (Columba livia) and Amazon parrots (Amazona species). J Avian Med Surg 14:8- 14, 2000 72. Schumacher J, Citino SC, Kernandez K, et al: Cardiopulmonary and anesthetic effects of propofol in wild turkeys. Am J Vet Res, 58:1014-1017, 1997 73. Sladky KK, Krugner-Higby L, Meek-Walker E, et al: Serum concentrations and analgesic effects of liposome-encapsulated and standard butorphanol tartrate in parrots, Am J Vet Res 67 (5):775-781, 2006 74. Staub J, Forbes NA, Thielebein J, et al.: The effects of isoflurane anaesthesia on some Doppler-derived cardiac parameters in the common buzzard (Buteo buteo).Vet J. 166 (3): 273-6. 2003 75. Stone EG, Redig PT: Preliminary evaluation of hetastarch for the management of hypoproteinemia and hypovolemia, in: 1994 Proceedings of the Association of Avian Veterinarians. Lake Worth, FL, 1994, pp197-199 76. Thurmon JC, Tranquilli WJ, Benson GJ: Lumb & Jones’ Veterinary Anesthesia (ed 3), Baltimore. MD, Williams and Wilkins, 1996 77. Touzot-Jourde G, Hernandez-Divers SJ, Trim CM: Cardiopulmonary effects of controlled versus spontaneous ventilation in pigeons anesthetized for coelioscopy. J Am Vet Med Assoc. 1;227(9):1424-8. 2005 78. Valverde A, Bienzle D, Smith DA, et al: Intraosseous cannulation and drug administration for induction of anesthesia in chickens. Vet Surg 22:240-244, 1993 79. Varner J, Clifton KR, Poulos S, et al: Lack of efficacy of injectable ketamine with xylazine or diazepam for anesthesia in chickens. Lab Anim 33:36-39, 2004 80. Vesal N, Zare P.: Clinical evaluation of intranasal benzodiazepines, alpha-agonists and their antagonists in canaries. Vet Anaesth Analg. 33(3):143-8.2006 81. Wilson D, Pettifer GR.: Anesthesia case of the month. Mallard undergoing phacoemulsification of a cataract. J Am Vet Med Assoc. 1;225(5):685-8. 2004 82. Whittow GC (ed): Sturkie’s Avian Physiology (ed 5). San Diego, CA, Academic Press, 2000

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Anesthesia and Analgesia in Reptiles Craig Mosley DVM, MSc, DACVA Oregon State University, Corvallis, OR A. PRINCIPLES OF REPTILE ANESTHESIA Anesthetic management of reptiles is associated with many challenges as their unique physiological and anatomical adaptations often complicate anesthetic administration, maintenance and monitoring. The provision of anesthesia to this diverse class of animals requires a thorough understanding of normal physiology, pathophysiology, the action and disposition of anesthetic and related drugs, and a familiarity with the design and use of related anesthetic equipment. Thorough pre-anesthetic assessment, a carefully designed anesthetic plan with attention to premedication, induction, maintenance, monitoring, supportive care, recovery and ongoing postoperative support and analgesia all contribute to the reduction of risk associated with anesthesia. Anatomy and Physiology Reptiles have long been considered a Class of animals that reflect the evolutionary transition between the aquatic and amphibious ectothermic vertebrates and endothermic birds and mammals. Many early investigations of reptilian physiology focused on their apparent “imperfections”. More recently, investigators have begun to view reptilian physiological adaptations as ideal and advantageous, enabling exothermic animals to inhabit almost all of the available non-polar ecological niches. Although many aspects of reptilian physiology are similar to those of endothermic vertebrates, significant differences are present. Such differences may alter both the action and disposition of anesthetics and analgesics. Metabolism & Thermoregulation The reptilian resting metabolic rate is one-tenth to one-third lower than the resting oxygen consumption rate of equivalent sized mammals. However, minimum and maximum oxygen consumption rates of individual reptilian species range from almost zero to values similar to those of a resting mammal. 1 A decrease in an animal’s cellular metabolic rate may result in reductions in drug metabolism leading to increases in both the latency of onset and duration of effect and time to recovery. The ambient environmental temperature is one of the main determinants of metabolic rate in resting reptiles. As temperature decreases, oxygen demand and metabolic requirements of tissues decrease, and the metabolic capacity of various organ systems decreases. There are significant inter-species and intra-individual variations in metabolic rate. Metabolic rate is also influenced by activity level and time since last feeding. Metabolic rate can increase 3 – 40 times the resting value following a meal and may remain elevated for up to 7 days 2. However, it is unclear whether recent feeding has a clinically significant effect on anesthesia in reptiles In general, the varanid and lacertid lizards tend to have relatively high metabolic rates, and boid snakes and chelonians have lower rates. Surface dwelling squamates have higher metabolic rates than burrowing species, and species of lizards that eat insects or other vertebrates have higher metabolic rates than do herbivorous species 3. Reptiles are ectothermic and derive their body temperature from the surrounding environment. However, some reptiles, such as, large pythons and leatherback sea turtles, derive some of their body heat from muscular activity. Such endothermic-like activity is only possible in larger reptile species. Reptiles can alter their body temperature through changes in cardiovascular function. During periods of warming some reptiles increase their heart rate and the degree of right-to-left shunting to increase the fraction of blood flow that is directed to the periphery for heating and ultimate return to the body core 4. This adaptation facilitates more rapid and efficient warming of the animal. Basking and shuttling between sun and shade are very important for temperature regulation in ectotherms. In all animals, the integration of physiology and behavior is affected by the internal thermal set point or preferred body temperature (PBT). In endotherms, the PBT generally remains constant. In reptiles the PBT may vary in response to physiological challenges such as fever. In the case of fever, many reptiles will alter their behavior and physiological responses to maintain this higher body temperature. There is good evidence that reptiles down regulate their body temperature in response to hypoxia and/or inadequate tissue oxygen delivery. This is referred to as hypoxia-induced hypothermia 5 6. Hypothermia-induced by hypoxia decreases metabolic rate through the direct effect of temperature on tissue oxygen demand and through depression of the rate of aerobic metabolism 5. The PBT can also be affected by hydration status. Reductions in hydration status lead to reductions in the PBT 7. Reptiles undergoing anesthesia should be maintained at the average or the high-end of their PBT range to ensure optimal metabolic function. These values can often be found in general husbandry references or extrapolated from known species from similar natural habitats.

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Cardiovascular system The non-crocodilian reptile heart has three chambers, with two completely separate atria and a single anatomically continuous ventricle. The crocodilian heart is more typical of that seen in mammals and birds with two completely divided atria and ventricles. In the crocodilian heart, the foramen of Panizza allows for some intravascular shunting under circumstances of breath holding, such as diving. In non-crocodilian reptiles the ventricle is divided into two main chambers by a septum-like structure called the Muskelleiste or muscular ridge. This ridge originates from the ventral ventricular wall and runs from the ventricular apex to base, dividing the ventricle into two main chambers; the cavum pulmonale and the cavum dorsale 8,9. The cavum pulmonale and the cavum dorsale are comparable in function to the right and left ventricles of mammals, respectively. The dorsolateral border of the muscular ridge is free, permitting the flow of blood between the cavum pulmonale and cavum dorsale. However, during ventricular systole the muscular ridge presses against the dorsal wall of the ventricle and separates the cavum pulmonale from the cavum dorsale, thus although exhibiting anatomical continuity of the subchambers, in a functional sense, the heart is capable of acting as a two-circuit pump. Cardiac shunting occurs commonly in reptiles 10-12. Cardiac shunts can occur in both directions and may occur simultaneously in both directions 11,13,14. The direction of the net shunt determines whether the systemic or pulmonary circulation receives the majority of the cardiac output. Intracardiac shunting has three important functions. First, shunting serves to stabilize the oxygen content of the blood during respiratory pauses. Second, the right-to-left shunt is partly responsible for facilitating heating by increasing systemic blood flow. Third, a right-to-left shunt directs blood away from the lungs during breath holding. During anesthesia, cardiac shunting can affect systemic arterial oxygen content and the uptake and elimination of inhaled anesthetics. The size and direction of the shunts are ultimately controlled by pressure differences between the pulmonary and systemic circuits and washout of blood remaining in the cavum venosum (an anatomical subchamber of the cavum dorsale described in many reptiles) 11-13,15. The pressure differences are principally controlled by cholinergic and adrenergic factors that regulate the vascular resistance of the pulmonary and systemic circulation 11,16-21. Large right-to-left shunts limit the amount of anesthetic uptake early in the anesthetic period and slow anesthetic elimination at the end of anesthesia. Such shunts can delay the induction to and recovery from inhaled anesthesia. Changes in the level and direction of shunts may account for the unexpected awakening seen in some reptiles anesthetized with inhalant anesthetics. Intracardiac shunts also have implications for patient monitoring, in particular airway gas monitoring and pulse oximetry. Blood pressure in reptiles is controlled by mechanisms similar to those described in mammals 22. The cardiovascular system of reptiles responds to both cholinergic and adrenergic stimulation in a manner similar to mammals and the presence of a baroreceptor reflex has been described 23. The resting blood pressures of reptiles tend to be stable in the absence of external stimuli but may vary with temperature, activity or state of arousal 24,25. In contrast to mammals, systemic arterial blood pressures vary greatly among various reptilian species, making it difficult to identify a “normal” arterial blood pressure 22. “Normal” blood pressure in reptiles may be more profoundly affected by environmental stresses such as habitat and temperature, species activity level and size compared to the role of these factors on influencing blood pressure in mammals. This greater variability may originate from a reptile’s poor ability to regulate normal homeostasis independent of temperature and environment. Chelonians tend to have the lowest mean arterial pressures (15-30 mmHg) while some varanids have resting arterial pressures (60-80 mmHg) similar to mammals 26. In the green iguana, normal resting systemic arterial blood pressures are reported to be in the range of 40-50 mmHg while pulmonary arterial pressures are in the range of 15-30 26. The systemic blood pressures in snakes correspond to the gravitational stress they are likely to experience 27 28,29. Snakes from arboreal habitats tend to have higher arterial pressures than those that are primarily aquatic. An allometric relationship between arterial blood pressure and body mass has also been described in snakes. As body mass increases, so does blood pressure 30. Several anesthetics, such as sevoflurane, isoflurane, halothane, propofol, tiletamine-zolazepam, and ketamine, have been shown to induce cardiopulmonary changes in reptiles similar to those seen in mammals 31-39. Pulmonary system The most significant difference between the respiratory physiology of reptiles, mammals and birds is the lower oxygen consumption rate of reptiles. This difference reflects the lower reptilian metabolic rate. Both reptile respiratory anatomy and physiology vary markedly across species. The lungs of non-crocodilian reptiles are suspended freely in the common pleuroperitoneal cavity and are not located in a closed pleural space. In reptiles, the lungs tend to be sac-like with varying degrees of partitioning. Highly aerobic species such as the varanids tend to have highly partitioned lungs with numerous septae and invaginations that increase the surface area for gas exchange. Chelonians and lizards tend to have paired lungs where most snakes have a single functional right lung. The functional units

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of the lung are referred to as ediculi and faveoli. Ediculi or faveoli are analogous structures to mammalian alveoli. Most reptile lungs exhibit areas of both type of parenchyma. There is little detail regarding the trachea and extrapulmonary bronchial tree system in reptiles. The tracheal rings of chelonians tend to be complete necessitating care when placing an endotracheal tube. In addition the trachea may bifurcate quite proximal, so inadvertent endobronchial intubation may occur. Many snakes also possess a tracheal lung, the significance of which is unclear. The lungs of reptiles tend to have a larger tidal volume but a smaller respiratory surface area. Because reptiles lack a diaphragm, they rely on the thoracic musculature for ventilation. Because both inspiration and expiration are active processes, the respiratory depression associated with anesthesia may be more profound than that observed in species in which expiration is a passive process. Because the muscles of ventilation include many of the same muscles used for locomotion, these two functions are relatively incompatible. Chelonians are faced with additional respiratory challenges since expansion of the thoracic cavity by movement of the ribs is not possible. The dorsal surface of the lungs is attached to the carapace and the ventral surface is attached to the abdominal viscera. Inspiration is accomplished by enlarging the visceral cavity, and expiration occurs by forcing the viscera up against the lungs, driving air out. This is accomplished by contraction of various posterior abdominal muscles and several pectoral girdle muscles. Control of Respiration. The control of respiration in reptiles is poorly understood. Both peripheral receptor and centrally-mediated control have been proposed. It seems most likely that there is an interaction between a central system which generates the pattern of respiration and afferent chemoreceptor input 40,41. Both carbon dioxide and pH changes appear important for stimulating normal ventilation but there is evidence that even under normoxic conditions oxygen tension may play a role in normal ventilation 42. Although there is some species variation, reptiles are generally viewed as episodic breathers 43,44 45. Pulmonary vascular perfusion is also intermittent and changes in perfusion are in concert with respiratory rate and rhythm 18 46,47 48. Ambient temperature has variable effects on the frequency, tidal volume and minute ventilation 49 and due consideration should be given to maintaining the optimal temperature for a particular species. Effects of Inspired CO2 and O2. The response of reptiles to inspired CO2 is quite variable. Inspiration of greater than 4% CO2 in snakes and lizards produces an increase in tidal volume and a decrease in respiratory frequency, and an overall decrease in minute ventilation 50,51. In turtles, specifically Pseudemys scripta and Chrysemys picta, the response to an increase in CO2 is an increase in minute ventilation as a result of increases in both respiratory frequency and tidal volume 18,52-54. In turtles, breathing less than 21% but more than 10% oxygen produces little change in the respiratory pattern. At inspired oxygen concentrations below 10%, some species increase ventilation while others retain their resting minute ventilation.and others may decrease ventilation 45,50,55-57. In those species in which minute ventilation decreases or remains unchanged, metabolic oxygen consumption decreases. During anesthesia, most reptiles are maintained using an inhalant anesthetic delivered in 100% oxygen. The delivery of a high oxygen concentration may further compound respiratory depression by blunting the contribution of oxygen to stimulate normal ventilation. In several reptiles species exposure to 100% oxygen significantly decreases minute ventilation 50,52,58-60 suggesting that high inspired oxygen may be responsible for at least some of the respiratory depression seen during anesthesia. The magnitude of this effect is likely small compared to the effects of anesthetics on central control of respiration and the muscles of respiration. However, there is some evidence that in the green iguana, recoveries from isoflurane anesthesia may be faster when the animal is ventilated with room air rather than 100% oxygen, possibly by improving ventilation and the subsequent removal of the inhalant from the body 61. Interestingly, in studies using Dumeril’s monitors (Varanus dumerili) no significant differences in recovery times from either isoflurane or sevoflurane anesthesia were found between animals ventilated with room air or those ventilated with 100% oxygen 62. This may reflect differences in study methods or species differences. Renal system Reptiles cannot produce urine more concentrated than plasma, making the excretion of nitrogenous wastes more difficult for terrestrial reptiles. Most reptiles excrete nitrogenous waste as uric acid (uricotelic). Some turtles and crocodilians can also excrete urea. Uric acid is produced in the liver and, unlike ammonia and urea, it is very insoluble in water and is excreted as a semisolid. In the reptilian kidney tubule, urine is very dilute so that uric acid remains in solution. Urine empties into the cloaca and then into the bladder or large intestine where water is reabsorbed causing the uric acid to precipitate. This results in the excretion of nitrogenous waste with relatively little water. The bladder of

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some reptiles can be used for the storage of water. Reptilian urine is not a good indicator of renal function. Many reptiles have specialized salt excreting glands that allow for the excretion of very high concentrations of sodium, potassium, and chloride. Many reptiles living in extremely arid environments can tolerate the marked fluctuations in total body water and plasma osmolarity that can occur in these environments. When faced with limited water supplies, plasma osmolarity can rise to levels higher than those known in any other vertebrate species. Hepatic system The reptilian liver appears to be similar in structure and function to the liver of other vertebrates. Although there is little detail known about the reptilian liver, it is assumed that it probably plays important roles in tolerance to anaerobic metabolism, hypothermia and adaptation to the physical environment. The liver of reptiles has a lower metabolic capacity compared to mammalian livers 63, and the metabolic rate is very sensitive to changes in temperature 64. The lower metabolic rates for the reptilian liver probably accounts for at least some of the prolonged effects commonly seen with drugs such as antibiotics. This may partly contribute to the prolonged anesthetic recoveries seen when using drugs that require extensive hepatic metabolism for termination of their clinical effect. B. CLINICAL ANESTHESIA I. Patient Presentation/Assessment Regardless of species or procedure, a thorough preanesthetic assessment should be performed on all patients. Patient assessment should include a complete history, species identification, and a full physical examination. Any additional supporting diagnostic tests such as blood work and imaging should be performed. Because most anesthetics produce some degree of cardiopulmonary depression, all animals should be physiologically stable prior to the induction of anesthesia. Unfortunately, in some reptiles, the size, disposition or anatomy may prevent even the performance of a routine physical examination. In these animals an assessment of body weight and general appearance may assist in determining the general health status of the animal. Species identification and information on the natural habitat of an animal may be useful when presented with a novel species. All animals should be kept at their preferred body temperature (PBT) throughout the anesthetic period and recovery. Performing any anesthetic-related procedure early in the day allows animals predisposed to prolonged recoveries to recovery during regular working hours rather than late into the night when support staff and patient supervision my be reduced 65. II. Premedication Premedications are used to facilitate handling and intravenous catheterization, reduce handling stress, and reduce the negative side effects associated with the administration of higher doses of drugs used for the induction or maintenance of anesthesia. Not all drugs administered prior to the induction of anesthesia will produce sedation, while others will not necessarily reduce the dose of drugs used for the induction or maintenance of anesthesia. Thus the goal of premedication should be established prior to selecting drugs for premedication. If the primary goal of premedication is to facilitate restraint it may be most appropriate to administer a combination of ketamine and an analgesic. If little chemical restraint is required, the premedication selection will be directed toward achieving pre-emptive analgesia. See Tables 2, 3 and 5 for drug doses commonly used in reptiles. Routes of Drug Administration Intramuscular. The intramuscular route of drug administration is most common in reptiles. Historically, hind limb and tail sites have been avoided because of concerns related to the first-pass effect associated with passage of any administered drug through the kidneys via the renal portal system. However, studies in some reptiles (turtles and the green iguanas) suggest that this may be more of a theoretical than practical concern, because only a small amount of blood from the hind limbs and tail pass through the kidney 66,67. However, it is probably best to avoid hind limb and tail administration of nephrotoxic drugs or those highly metabolized or excreted by the kidneys. The epaxial muscles provide a suitable injection site in most snakes. In lizards, the muscle mass of the forelimb (triceps and biceps), hindlimb (quadriceps, semimembranosus and semitendinosus) and tail can be used. Caution should be used in species known to autotomize (drop) their tails (many geckos), beacause it is possible for an animal to “shed” their tail during handling. In chelonians, injections are most often administered in the triceps muscle. The cranial surface of the foreleg should be avoided since the proximity of the radial nerve to injection sites in this area increases the risk of damage to this nerve. The pectoral muscles can also be used although in many species there is a lack of significant muscle mass in this area.

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Intravascular. While intravenous drug administration is not always feasible in reptiles, the combination of good technique, practice, appropriate patient selection, and skilled physical restraint can facilitate predictable access to the ventral coccygeal vein in even very small snakes and lizards and the dorsal coccygeal vein in tortoise and freshwater turtles. In sea turtles, the dorsal cervical sinus has also been used for intravenous administration of drugs 68. Intravascular injection decreases the latency of onset of action of an administered drug. It also decreases the variability in uptake that is associated with intramuscular injections in reptiles. Some drugs produce tissue irritation following intramuscular irritation. Intravenous administration of these irritant drugs may obviate such tissue irritation. Techniques for catheterization of the coccygeal vein in both lizards and crocodilians have been described 69. Intravenous catheterization of the coccygeal or abdominal veins is most often performed “blind”. In some species of turtles and tortoises the jugular vein can be visualized, however; visualization of the jugular vein most often requires a skin incision and blunt dissection. Venous sinus sites are not ideal sites for intravenous catheter placement. Although over-the-needle catheters are most frequently used, a technique utilizing a small guage wire stylet through a needle (Seldinger technique) can be used to facilitate difficult catheterization. When required, cut-down procedures should be preformed in conjunction with a local or general anesthetic. Lidocaine diluted down to a 1% solution with sterile saline can be used for local infiltration. Although toxic doses have not been determined in reptiles it is probably best to use less than 8-10 mg/kg. The most common sites for vascular access and associated technical tips are presented in Table 1. Intraosseous. Intraosseous catheterization is occasionally used to secure intravascular access in dogs, cats, and birds. Intraosseous catheter placement has been described in the green iguana (Iguana iguana) and sea turtles 31,70,71. This is a technique best suited for use in lizards and can be is performed in most species. One study, examining kidney function in green iguanas (Iguana iguana), found similar renal uptake of the radioactive substance whether administered introsseously or intravenously 70. This suggests that intraosseous drug administration is a suitable alternative to intravenous administration. To this authors knowledge propofol is the only anesthetic drug that has been studied for intraosseuos administration but many other anesthetic and non-anesthetic drugs have been administered successfully via this route. Sites for intravascular access in reptiles: Squamates (snakes) 1) Coccygeal vein is located on the ventral midline of the tail. The needle should be inserted sufficiently caudal to the vent in order to avoid the hemipenes and anal sacs. The vessel is entered via a ventral midline approach, the needle is advanced with gentle suction until the vein or a vertebral body are contacted. 2) Jugular vein can be used but requires a skin incision to visualize. An incision is made 4 to 7 scutes cranial to the heart at the junction of the ventral scutes and lateral body scales. The vein is then identified using blunt dissection just medial to the tips of the ribs. 3) Palatine vein is easily visualized in larger snakes and is located medial to the palatine teeth in the roof of the mouth. The technique is greatly facilitated by short term anesthesia but it is possible to collect blood from these vessels in awake animals using a mouth speculum 4) Heart, Use of the heart for venipuncture is not recommended except for emergency situations. 5) Intraosseous, there are to this authors knowledge no intraosseous sites described for drug administration in snakes. Squamates (lizards) 1) Coccygeal vein is located on the ventral midline of the tail. The needle should be inserted sufficiently caudal to the vent in order to avoid the hemipenes. The vessel can be entered from either a ventral midline approach or laterally. The ventral approach is simple to perform the needle is advanced with gentle suction until the vein or a vertebral body are contacted. The lateral technique involves inserting the needle just ventral to the transverse process of the vertebral body and walking the needle ventral until the vein is contacted. 2) Ventral abdominal vein is located on the ventral midline of the abdomen and can be entered percutaneuosly or via a small skin incision for direct visualization of the vessel. 3) Cephalic vein is located on the dorsal surface of the distal foreleg. A skin incision is generally required for visualization. 4) Jugular vein is located on the lateral surface of the neck at about the level of the tympanum and may be palpated in some species but is generally difficult to visualize. A small skin incision is often required for direct visualization. The jugular veins tend to be located more dorsal than those in mammals. There is a large lymphatic sinus close to the vein and contamination with this lymph fluid occurs frequently. 5) Intraosseous techniques have been described for the distal femur, proximal tibia, and

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proximal humerus (heard 2001). The techniques are similar to those described for other small animal patients. Chelonian (turtles and tortoises) 1) Dorsal coccygeal vein is located midline dorsal to the coccygeal vertebrae. It is a technique requiring minimal restraint. The needle is introduced in a craniad direction at a 45-90 degree angle from the skin 2) Dorsal cervical sinus (supravertebral) is located on the dorsolateral aspect of the neck in sea turtles. It is located one third the distance from the carapace to the head, cranial to the craniad edge of the carapace. The head is directed forward and down and the needle is introduced lateral to midline on either side. 3) Occipital venous sinus has been described in freshwater turtles and is located midline below the occipitus. It requires that the head be restrained firmly and in an extended ventroflexed (45-90 degree angle from the carapace) position. The needle is then introduced midline just caudal to the occipitus and nearly perpendicular to the spine. Lymph contamination is a possibility. 4) Subcarpacal sinus or supravertebral sinus is located under the carapace just caudal to the last cervical vertebrae and craniad to the first thoracic vertebrae. This sinus can be approached by pressing the head into the shell and palpating for the first thoracic vertebrae (incorporated into the carapace). The needle should be directed through the skin just caudal to the juncture of the last cervical vertebrae up towards the carapace and first thoracic vertebrae. 5) Jugular veins are located on the lateral sides of the neck at about the level of the tympanum. In some species venipuncture of the jugular vein is relatively straightforward and can be visualized or a small skin incision can be made to facilitate direct visualization. Unfortunately this technique requires the neck to be fully extended and in uncooperative animals a short acting anesthetic or tranquilizer may be required. 6) Intraosseous techniques have been described using the plastron/plastron bridge but like other authors (Heard 2001) this author has found most catheters end up in an intracoelomic rather than intraosseous position. The technique is described as passing a needle at an angle through the bony bridge between the plastron and carapace. Drug Selection Anticholinergics. Atropine and glycopyrrolate should probably not be used to decrease salivation but may be indicated if bradycardia develops. Anticholinergics can increase salivary viscosity and may predispose the patient to obstructions from highly viscous mucous in airways or small diameter endotracheal tubes. Anticholinergic drugs can alter intracardiac shunt fractions in reptiles. This may alter a patient’s response to anesthetic drugs, particularly inhaled anesthetics. Phenothiazines. Phenothiazines such as acepromazine tend to be relatively ineffective sedatives in reptiles. Their use requires the administration of large doses that are associated with prolonged effects. Acepromazine is not a very useful drug in reptile anesthesia 65,72,73. Ketamine. Ketamine, a phencyclidine, is not routinely used as a premedication drug in most animals. Ketamine is regarded as an anesthetic but at subanesthetic doses ketamine produces analgesic effects and can produce profound restraint. At subanesthetic doses, ketamine induces a cataleptic state characterized by the presence of uncoordinated voluntary and involuntary muscle movement that may appear in response to external stimuli. It is very important to recognize that an animal in this state should not be considered to be at a surgical plane of anesthesia. Ketamine is used frequently as a component of a premdication protocol to produce restraint in chelonians and other reptiles. Ketamine has also been used alone for restraint or the induction of anesthesia in a variety of reptiles 74-76. In snakes, ketamine alone produces hypertension, tachycardia, bradypnea and hypoventilation 32,33. Similar effects on the heart rate and respiratory rate have been observed in skinks (Tiliqua rugosa and Egernia kingii) 37. Since ketamine is also associated with muscle rigidity, it is most often combined with a drug that produces muscle relaxation (benzodiazepines, alpha2 agonists). Telazol. Telazol is a proprietary combination of tiletamine and zolazepam. Tiletamine is a long acting phencyclidine similar to ketamine, while zolazepam is a long acting benzodiazepine similar to diazepam. Telazol has been used in reptiles with variable results 77-79. In the boa constrictor (species not identified) tiletamine-zolazepam (12.5 mg/kg IM) failed to produce surgical anesthesia but produced safe immobilization associated with a transient increase in heart rate, and an increase in respiratory rate that is not associated with changes in minute ventilation, systolic blood pressure or arterial oxygen saturation 36. The combination of tiletamine and zolazepam is a less desirable combination than ketamine and midazolam because of the longer duration of action of the

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tiletamine/zolazepam that can cause more prolonged recoveries. Telazol is occasionally used in very large reptiles to reduce the injected volume; however, prolonged recoveries are likely to be observed. Benzodiazepines. Midazolam is a water-soluble benzodiazepine and can be administered both intramuscular and intravenously. Diazepam is not recommended for intramuscular use as it is very poorly absorbed via this route of administration. Midazolam (2 mg/kg) is used in combination with ketamine (20-40 mg/kg IM) to facilitate handling and to induce anesthesia in chelonians 80. Midazolam (1.5 mg/kg IM) has also been used alone in freshwater turtles (Trachemys scripta elegans) with some success 81 but fails to provide significant sedation when used alone in snapping turtles (Chelydra serpentina, 2.0 mg/kg IM) 80 and painted turtles (Chrysemys picta, 2.0 mg/kg IM) 82. Alpha-2 agonists. The alpha2 agonists produce analgesia, sedation and muscle relaxation in mammals. In some reptiles, it appears to produce desirable levels of sedation and muscle relaxation while in others little to no effect is observed. The analgesic effects of alpha-2 agonists have not been evaluated in reptiles but clinical impressions suggest an analgesic effect as well. Xylazine (2 mg/kg IM), in combination with ketamine (60 mg/kg IM), produced a variable level of light anesthesia suitable for minor procedures only in red-eared sliders (Trachemy scripta elegans) 83. More recent reports describe the use of medetomdine rather than xylazine. Medetomidine has a higher alpha2:alpha1

binding ratio than xylazine. Medetomidine (150 μg/kg IM) is an effective sedative in desert tortoises (Gopherus agassizii) 84. Medetomidine in combination with ketamine produces anesthesia of a sufficient depth to allow endotracheal intubation in several species of tortoises 85,86, red-eared slider turtles (Trachemys scripta elegans) 87, and loggerhead sea turtles (Caretta caretta) 68. The administration of medetomidine to several mammalian species is known to be associated with marked cardiovascular side effects that include arrhythmias, a decrease in cardiac output, and an increase in systemic vascular resistance. It appears that some of these changes may also occur in reptiles. Medetomidine induces a significant decrease in heart rate, respiratory rate and systolic, diastolic and mean ventricular pressures, and a decrease in ventricular partial pressure of oxygen in desert tortoises (Gopherus agassizii) 84. Medetomidine, in combination with ketamine, produces a moderate increase in arterial pressure, and moderate hypercapnia and hypoxemia in desert tortoises (Gopherus agassizii) 85. One advantage of using alpha2 agonists is that they are reversible, a property that can be of benefit when faced with prolonged recoveries. Following the administration of atipamezole, animals appear normal within 30-60 minutes. Atipamezole (500 μg/kg IV) produces marked arterial hypotension 85 but intramuscular administration does not appear to produce significant alterations in ventricular pressures 84. Thus, intramuscular, rather than intravenous, is the recommended route of administration of atipamezole. Opioids. Opioids are very poor sedatives in reptiles 65. Although they are commonly used in the perianesthetic period to provide analgesia 88, there are few studies evaluating the use of opioids for pain and analgesia. Regardless, it is strongly recommended that an analgesic be administered prior to any procedure that may be associated with significant tissue damage. III. Induction Ketamine. Both ketamine and tiletamine can be used alone to induce light anesthesia or a level of restraint adequate for endotracheal intubation. It is questionable whether satisfactory surgical anesthesia can be achieved using ketamine or telazol alone in reptiles 78 37 36 74,75. Many reptiles maintain reflex movement even when using very high doses of ketamine and tiletamine. In order to achieve a level of anesthesia appropriate for surgery, ketamine should be administered in combination with a drug that produces muscle relaxation (midazolam or medetomidine). In iguanas, tiletamine (10 mg/kg IM) was used as the sole drug for the induction and maintenance of short-term anesthesia. The mean induction time was 6.5 minutes and a level of anesthesia sufficient to allow endotracheal intubation was produced 89. However, recoveries may be protracted. Telazol (33-44 mg/kg) produced surgical anesthesia in green iguanas (Iguana iguana) but anesthesia persisted for 12 hours or more 78. Propofol. Propofol is an alkylphenol, structurally different from other anesthetics such as barbiturates, eugenols or steroids. It is prepared in an intralipid solution intended for intravenous use. In mammals, propofol produces a rapid and smooth induction of anesthesia with a very predictable duration of action. The elimination of propofol involves non-hepatic sites, most likely the lung. Propofol (3-10 mg/kg IV) is the induction drug of choice when intravenous access is available. It is a reliable means of inducing anesthesia without unnecessarily prolonging recovery time. In mammals, the administration of propofol is commonly associated with apnea and hypotension. The intraosseous administration of

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propofol (5 and 10 mg/kg) has been evaluated in the green iguana (Iguana iguana). In this species, the administration of propofol is associated with a prolonged period of apnea 31. Inhaled Anesthetics. Inhaled anesthetics can be used for the induction of anesthesia. The least soluble of the inhalant anesthetics, sevoflurane, desflurane or isoflurane, are preferred because the solubility of an inhaled anesthetic is inversely related to the times for both induction of, and recovery from, anesthesia. In some reptiles, induction of anesthesia with an inhaled anesthetic can be very prolonged because of breath holding. Mask induction of chelonians can be very difficult because of breath holding and limited access to the head. The induction of anesthesia using inhaled anesthetics is generally easier in snakes and lizards but prolonged periods of breath holding may occur in these species as well. In some species, respirations can sometimes be stimulated by stroking the animals lateral thorax. The average induction time for green iguanas (Iguana iguana) using isoflurane in 100% oxygen administered by face mask is approximately 20 minutes. The prior administration of butorphanol does not effect the duration of induction 90. In Dumeril’s monitors (Varanus dumerili), induction times with sevoflurane (11.20 ± 3.77 min) are significantly faster than the inductions times using isoflurane (13.00 ± 4.55 min) 62. The addition of nitrous oxide (34% oxygen; 66% nitrous oxide) to the carrier gas significantly reduces the time to induction of anesthesia with sevoflurane 62. In addition to mask induction with an inhaled anesthetic, many reptiles can be tracheally intubated while awake and then manually ventilated to induce anesthesia. This technique can reduce the time for induction of anesthesia but it may be associated with higher levels of stress. The use of premedications and topical administration of local anesthetic to the glottis may help reduce this stress. Muscle Relaxants. Both depolarizing (succinylcholine) and nondepolarizing muscle relaxants (atracurium and rocuronium) are used in reptiles 79,91-94 95. Muscle relaxants act by competitive inhibition of acetylcholine at the neuromuscular junction, leading to paralysis. They are used primarily to facilitate immobilization and tracheal intubation of crocodilians 79,91-93 but are also occasionally used in chelonians 95. Muscle relaxants are not anesthetics and have no analgesic or amnesic properties. The routine use of muscle relaxants for immobilization of reptiles should be avoided. Their use may be indicated (but always in combination with analgesic and amnestic drugs) for managing very dangerous and aggressive species or in field situations when a very rapid immobilization is required to limit the potential for animal injury. Endotracheal intubation. Intubation is easily accomplished in most reptiles. In most reptiles the glottis is in a relatively rostral position in the mouth at the base of the tongue. The glottis is easily visualized and intubation is accomplished via direct visualization. A small drop of lidocaine (diluted to 1%) can be used to desensitize the glottis and may facilitate tracheal intubation. In some aquatic reptiles anatomical modifications of glottal folds may obscure direct visualization of the glottis. The animal should be intubated with the largest diameter tube that can easily be placed. The mucous of reptiles tends to be very viscous and mucoid plugs can form in endotracheal tubes during longer procedures. Attention to this possibility is important and can be recognized as an inability of the lungs to fully deflate during expiration. The trachea of chelonians bifurcates quite rostrally and endobronchial intubation is possible. The tracheal rings in chelonians and crocodiles are complete and in most reptiles cuffed endotracheal tubes are avoided to prevent accidental over inflation and damage to the tracheal structures. IV. Maintenance of Anesthesia. Inhalant anesthesia is commonly used for maintenance of anesthesia in reptiles. The physical properties of the newer inhaled anesthetics afford minimal uptake and metabolism and predictable recovery. The administration of inhalant anesthetics is normally carried out with oxygen as the carrier gas and can reduce the risk of hypoxia, despite the observation that reptiles are more tolerant of periods of hypoxemia than mammals or birds 96. Methoxyflurane and halothane are no longer readily available and are not inhalant anesthetics recommended for reptiles. Isoflurane, sevoflurane and desflurane are more appropriate choices. Isoflurane, sevoflurane, desflurane have all been evaluated in reptiles 35,39,62,90,97-102. The MAC of sevoflurane in Dumeril’s monitor (Varanus dumerili) has recently been found to be 2.51 ± 0.5%, this is similar to values in mammals (2.1-2.3%) 99. The range of MAC values for isoflurane reported for reptiles (1.54% to 3.14%) is more variable than that reported for mammals and birds. This may simply be a reflection of the techniques used for MAC determination, the body temperature of the patient, or actual species differences. Using comparable techniques, the MAC of isoflurane in the green iguana (Iguana iguana) (2.1 ± 0.6%) and Dumeril’s monitor (Varanus dumerili) (1.54 ± 0.17%) were found to be significantly different 90,98. There is also greater variability in MAC values in green iguanas than those observed in

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Dumeril’s monitors. The pronounced right to left intracardiac shunting in snakes, turtles and non-varanid lizards may account for some of these differences. In many aquatic reptiles that are capable of long periods of dive-induced breath holding, significant right-to-left shunting produces end-tidal anesthetic concentrations of inhaled anesthetics that may not be entirely reflective of those in the blood and hence the brain. Concentrations in the lung may substantially overestimate levels in the brain leading to erroneously elevated MAC when using traditional methods of MAC determination. That many reptiles either fail to become adequately anesthetized or induce to anesthesia very slowly with an inhaled anesthetic likely reflects the impact of significant right-to-left intracardiac shunting on the uptake of an inhaled anesthetic. A right-to-left intracardiac shunt results in a reduction of the volume of blood that is exposed to the inhalant at the gas exchange interface. In contrast, it is not uncommon to observe deep anesthesia in reptiles, even after very few breaths. This may be the result of the accumulation of inhaled anesthetic in the sac like structure of reptilian lungs and the breathing patterns observed in most reptiles. Many reptiles are episodic breathers that take several breaths that are followed by a prolonged inspiratory pause. Such ventilation patterns are energetically efficient and may have developed to best meet the low metabolic oxygen demand of reptiles. This ventilation pattern, in association with the sac like structure of the reptilian lung, affords continual access to oxygen without unnecessary energy expenditure. As a consequence, the lung may function as a reservoir of inhaled anesthetic that is available to the patient during breath holding. Thus the extent of right-to-left cardiac shunting may have more of an impact on the speed of induction of anesthesia using an inhaled anesthetic than does ventilation rate. Dose dependent cardiovascular depression occurs during isoflurane anesthesia of the green iguana (Iguana iguana) 39. Both blood pressure and heart rate decrease in a dose dependent manner. It is likely that similar cardiovascular depression occurs in other reptiles. However, the effects on heart rate are likely to be more variable. Ventricular blood pressures and heart rates in desert tortoises (Gopherus agassizii) do not change with increasing doses of sevoflurane anesthesia 35. Interestingly, the dose of isoflurane required to induce cardiovascular arrest in healthy green iguanas is much greater than the maximum percent delivered by most commercial isoflurane vaporizers (5%) 97. Even at levels 4 times greater than MAC (2.1%), isoflurane failed to induce cardiovascular arrest, suggesting a wide safety margin for this anesthetic when used in the healthy green iguana. Standard inhalant equipment used in small animal anesthesia is suitable for administering inhalant anesthetics to most reptiles. An anesthetic machine equipped with a flowmeter, precision vaporizer and either a non-rebreathing circuit or a circle system is often used. In very small patients weighing less than 1 kg, a non-rebreathing or a pediatric circle system is preferred. The dead space associated with a standard adult circle system may lead to substantial rebreathing of expired gases. However, in reptiles it has been shown that adding carbon dioxide to the inhaled gases may actually improve ventilation during inhalant anesthesia 34,103. Oxygen flow rates should meet or exceed the oxygen consumption of the patient. The flow rates used for standard small animal patients are suitable for most reptiles: 50 to 100 mL/kg/min when using a re-breathing system and 200 to 300 mL/kg/min when using a non-rebreathing system (Bain, Ayres TPiece). For some vaporizers, the lower limit of oxygen flow rate required to maintain vaporizer accuracy is about 200 mL/min. This should be the lower limit regardless of patient size. Ventilators are very useful when anesthetizing reptiles since most, if not all, become apneic during general anesthesia. Most commercial ventilators are not well adapted to delivering the small tidal volumes required by many reptiles. It is important to recognize that, in addition to the ventilator-delivered tidal volume, the fresh gas flow rate contributes to the delivered tidal volume during inspiration. This is most significant in very small animals when high oxygen flow rates are used. Ventilators designed for small mammals are particularly useful when ventilating small reptiles.

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Dosages of commonly used anesthetic drugs -Chelonians Glycopyrrolate IV, IM,SC: 0.01-0.04 mg/kg* :May increase viscosity of secretions increasing risk of obstruction 73 Atropine IM,IP 0.04 mg/kg*: May increase viscosity of secretions increasing risk of obstruction 104 Acepromazine IM 0.1-0.5 mg/kg*: Minimal effect 105 Medetomidine IM, IV 50-100 μg/kg (tortoises), 150-300 μg/kg (aquatic turtles): Variable sedation when used alone, best combined with ketamine 68,84-87 Xylazine IM 2 mg/kg: Did not improve anesthesia over ketamine alone in red-eared sliders (Trachemys scripta elegans) 83 Atipamezole IM, IV 500 μg/kg: May be best to administer IM 84-86 Midazolam IM 1.5-2.0 mg/kg : May be unreliable on its own, can facilitate handling, mask inductions and may improve the overall quality of anesthesia, best in combination with ketamine 80-82 Ketamine IM, IV 5-20mg/kg (in combination): Best combined with alpha-2 agonistor benzodiazepine, Doses up 60 mg/kg have been used. 68,80,83,85-87 Tiletamine/zolazepam IM 3.5-10 mg/kg :Prolonged recoveries likely 77,78 Propofol* IV, IO 3-5 mg/kg Predictable effects and recovery, first choice for induction of Anesthesia 65 Isoflurane* Inhaled 2-3% on vaporizer: MAC not determined 65 Sevoflurane Inhaled 4-5% on vaporizer MAC not determined 65 * Doses not determined experimentally, extrapolated or anecdotal Dosages of commonly used anesthetic drugs - Lizards and snakes Glycopyrrolate IV, IM,SC 0.01-0.04 mg/kg* :May increase viscosity ofsecretions increasing risk of Obstruction 73 Atropine IM,IP 0.04 mg/kg* : May increase viscosity of secretions increasing risk of obstruction 104 Acepromazine IM 0.1-0.5 mg/kg* :Minimal effect 105 Medetomidine IM,IV,IO 150 μg/kg* : Not commonly used 65 Midazolam IM 0.5-2.0 mg/kg*: Minimal sedation 72 Ketamine IM, IV,IO 22-88 mg/kg (alone)10-15 mg/kg(combined with medetomidine): Best used in combination with medetomidine 74,75,76,33,32,37 Tiletamine/zolazepam IM, IV, IO 3-6 mg/kg Prolonged recoveries likely, even at high doses animals may remain responsive 77,78,36,79,89 Propofol IV, IO 5-10 mg/kg : Predictable effects and recovery, first choice for induction of Anesthesia 31 Isoflurane Inhaled 2-3% on vaporizer MAC 1.5-2.1% 90,98,100,102 Sevoflurane Inhaled 4-5% on vaporizer MAC 2.5-3.1% 99,102 Desflurane Inhaled 9-10% on vaporizer MAC 8.9% 102 IO for lizards only * Doses not determined experimentally, extrapolated or anecdotal V. Monitoring and Perianesthetic Support The goal of anesthesia is to achieve and maintain a reasonable surgical plane of anesthesia while preventing anesthetic overdose. Safety during anesthesia is prevented by titration of the inhaled anesthetic in response to an individual animal’s requirements. The necessity for such adjustments are determined by careful patient monitoring. Comprehensive monitoring includes assessment of several reflexes and a determination of the response of the cardiopulmonary system to anesthesia. Reflexes In 1957 Kaplan and Taylor 106 published a study involving the use of ether, nembutal (sodium pentobarbital) and urethane in adult turtles (Pseudemys spp). They recorded heart rates and rectal temperatures, and observed the degree of muscle tone, voluntary movements, pupillary diameter, and presence or loss of the corneal reflex in order to assess depth of anesthesia. They defined deep or surgical anesthesia as a plane of anesthesia associated with muscular relaxation, absence of response to painful stimuli, and loss of movement. Kaplan and Taylor were pioneers in this area; anesthetic depth in reptiles is still determined using some of the same qualitative parameters they defined. Interestingly, when reptiles are induced with inhalant anesthetics muscle relaxation frequently begins at midbody and moves cranially, then caudally. Tail tone is lost last. This has been demonstrated in lizards using halothane and sevoflurane 34 62 and in turtles using ether 107. These features can be used when assessing depth during induction and recovery.

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Cardiovascular Direct auscultation of cardiac function is a simple method of assessing heart rate and rhythm. External auscultation is best preformed using a stethoscope with a small pediatric bell but this technique can be difficult due to interference from scales or the carapace and plastron in chelonians. A dampened gauze placed between the chest wall and the stethoscope bell can reduce interfering noise from scales. In anesthetized animals a small esophageal stethoscope works very well for direct auscultation of the heart. The stethoscope tubing should be advanced in increments until the point where maximal sound intensity is reached. It is not uncommon for some reptiles to have heart rates of 20 beats per minute or less. If the esophageal stethoscope is not advanced slowly it is easy to bypass the heart and place the stethoscope in the stomach. This may predispose the animal to regurgitation. An excellent alternative to direct auscultation is the use of an ultrasonic Doppler device that detects blood flow in major vessels and the heart itself. There are a variety of probes; adult and pediatric flat probes and pencil probes. These probes are most easily placed over the heart and held in place with tape. Alternatively, the carotid, coccygeal or femoral arteries may be used as the sites for probe placement. In chelonians, the shell generally precludes use of the heart. Pediatric probes have greater sensitivity in detecting flow in small vessels and are preferred for use in reptiles. In addition to providing an audible signal of blood flow through the vessels over which the probe is placed, the Doppler unit can also be used to assess blood pressure in a manner similar to that used during the anesthesia of non-reptilians. A small, inflatable cuff is placed around the limb or tail proximal to the probe. Blood pressure values obtained using this technique in reptiles have not been compared to direct arterial measurements; however, the technique is still useful for assessing changing trends in blood pressure. The electrocardiogram (ECG) can be used to monitor the electrical activity of the heart in reptiles. At the very least, the ECG and provides an assessment of heart rate and rhythm. Electrical activity can continue in the heart despite loss of muscular activity, a condition known as pulseless electrical activity (PEA) or electromechanical dissociation (EMD). Thus, it is best not to rely solely on an ECG for evaluation of cardiovascular function. The morphology of the reptilian ECG is similar to that of mammals with the addition of an SV wave proceeding the P wave 43. While the ECG leads on reptiles are positioned similar to the standard 3 lead configuration in mammals, some modification in lead placement will improve signal strength and ECG quality. In lizards, the right and left forelimb leads are placed in the cervical region since the heart is located in the pectoral girdle 108. In snakes, the active leads are placed two heart-lengths cranial and caudal to the heart 108. The heart in snakes is located 20-25% of the entire body length from the head and can often be identified by direct visualization of ventral scale movement caused by cardiac activity. In chelonians, the forelimb leads are placed on the skin between the neck and the forelimbs 108. Stainless steel suture loops or needles can be placed through the skin and attached to the leads can improve signal strength. Respiratory Direct visualization of respiratory movements can be extremely difficult in many reptiles, particularly chelonians and very small species. Additionally, chest and body wall excursions, bag movement and fogging of the endotracheal tube can be misleading and may not always represent adequate ventilation. Because most reptiles require intermittent positive pressure ventilation the utility of monitoring spontaneous respiration is reduced. Reptiles rarely breathe well when anesthetized 31,65,104,109, making mechanical ventilation appropriate in most cases. Current recommendations for ventilatory support include rates of 2-6 breaths per minutes using tidal volumes ranging from 15-30 ml/kg with peak airway pressures less than 10 cmH2O. Manual IPPV is commonly performed but several small animal specific ventilators are now available. Pulse oximetry is a non-invasive method used to assess functional hemoglobin saturation (SpO2). Under normal circumstances this value correlates closely with arterial hemoglobin saturation (SaO2). Although pulse oximetry is used frequently during reptile anesthesia, the results should be interpreted with caution. Pulse oximetry was specifically developed for use in humans, using the oxygen binding characteristics of mammalian hemoglobin to guide the development of the technology. A reflectance probe for pulse oximetry is most commonly placed in either the esophagus or cloaca. The heart rate reported by the pulse oximeter should correlate with the heart rate determined using direct methods (auscultation). The efficacy of this technology has only been assessed in a single reptilian species the green iguana (Iguana iguana). In this species, values obtained during pulse oximetry with an esophageal reflectance probe placed in the esophagus (SpO2) correlate closely with arterial hemoglobin saturation (SaO2) of blood taken from the abdominal aorta 61. Other investigators have not been able to establish such a relationship between arterial hemoglobin saturation and hemoglobin saturations determined using pulse oximetry 39. Capnometry measures the amount of carbon dioxide in the expired gas during the ventilatory cycle. Endtidal refers to the fact that the quantitative measurement derived during capnometry refers to the

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concentration of carbon dioxide in the last portion of the expired volume, the end-tidal volume. This gas most accurately reflects the gas contained in the gas exchange portions of the lung, rather than the gas in the conducting airways. End-tidal carbon dioxide (ETCO2) concentrations are generally reflective of the carbon dioxide concentrations in arterial blood although the level of carbon dioxide is generally lower due to the dilution of the expired carbon dioxide by non-carbon dioxide containing gases in the conducting airways. Much more information can be obtained from a capnogram, a graphic representation of the ETCO2 concentrations over the entire respiratory cycle. While capnography is a useful monitoring tool in mammals with normal lungs, the utility of capnography in monitoring respiratory function in reptiles has not been established. The presence of right-to-left intracardiac shunts and deadspace ventilation associated with the unique structure of many reptilian lungs makes information gathered using this monitoring modality difficult to interpret. Blood gas analysis in reptiles is subject to significant over-interpretation and misinterpretation. Numerous factors such as site of sampling, arterial versus venous blood, species, inspired oxygen concentration, thermoregulatory status and the ventilatory status of the patient (spontaneous versus controlled) will all affect interpretation of blood gas values. Reptiles tend to be much more tolerant to alterations in pH, PCO2 and PO2 than mammals and thus normal values for mammals may not be applicable to reptiles. This said, in general, normal pH in reptiles tends to be similar to mammals, provided comparisons are made at identical temperatures. Most reptiles have body temperatures below that of most mammals and consequently their normal pH values tends to be higher. PCO2 and PO2 tend to be lower in reptiles when compared to the same values in mammals. PO2 values are lower as a result of intracardiac shunting but also as a result of intrapulmonary shunting, ventilation-perfusion mismatching and there is some evidence that in some reptiles there may also be impairment to diffusion of oxygen from the lung into the blood 42. Given our current state of knowledge, it is difficult to critically evaluate blood gas analysis in reptiles. Fluid Therapy Fluids should be administered prior to anesthesia if clinically significant dehydration is noted. Fluids are best administered intravenously or intraosseously but they can also be given intraperitoneally or subcutaneously. Fluid movement, distribution, and homeostasis in reptiles differs significantly from mammals. Reptiles tend to have a greater proportion of total body water in the intracellular space (45-58%) 110. For this reason, some have suggested using hypotonic replacement solutions. However, it is not clear that this is of benefit to the animal unless the dehydration is associated with pure water loss. It is probably best to use a standard balanced electrolyte solution. Some reptiles are capable of tolerating extreme alterations in total body water and plasma osmolarity when water resources are scarce. The significance of such an adaptation for fluid therapy is not clear. Each patient should be carefully assessed and the fluid therapy plan should be tailored to meet the needs of the individual patient. Thermoregulation Reptiles are exothermic animals that derive nearly all their body heat from the external environment. Thermoregulation in reptiles is a complex interaction between the animal’s internal environment and the external environment. Body temperature is regulated primarily through complex behavior patterns and alterations in the cardiovascular system. Most reptiles have a preferred body temperature (PBT) range that is associated with optimal metabolic function. It is probably best to maintain animals in hospital care at the upper end of the PBT for that species. This is easily accomplished using circulating warm water blankets, warm water bottles, and warm forced air. Body temperatures below the PBT for the individual animal may be associated with prolonged drug effects and may impair the animal’s immune system and healing 34,36,37. VI. Recovery & Analgesia Reptiles should be monitored throughout the recovery period. Since recovery from anesthesia in reptiles can be prolonged, inhaled anesthetics are often discontinued 15-20 minutes prior to completion of the procedure. Delayed recoveries seem to be more common in less aerobic reptiles which may be the result of significant right-to-left shunting and low cardiac output that lead to a protracted elimination of the inhalant from the body. Body temperature is also very important for facilitating recovery and optimal body temperature should be maintained throughout the recovery period. Consideration for the post-operative analgesic needs of the animal should be made based on clinical signs and the anticipated degree of tissue damage associated with the procedure. Reducing the oxygen concentration by allowing the animal to breath room air may help hasten recovery 61. The analgesic management of reptiles can be challenging due to their unique physiological, anatomical and behavioural adaptations. Although reptile pain and nociception have not been

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extensively studied there is very strong evidence that reptiles are capable of nociception. The neuroanatomic components necessary for nociception have been described in 111,112. In addition, endogenous antinociceptive mechanisms 112,113 and a demonstrable modulation of nociception with pharmacological agents known to be analgesics in other species have been demonstrated 114-118. In lizards (Gekko gecko), spinal projections originating in the brainstem region (nucleus raphes inferior) that project to the superficial layers of the dorsal horn have been identified and these structures suggest the presence of tracts, similar to those found in mammals, that mediate descending inhibition of nociception 117. Neurotransmitters that are important in nociceptive modulation in mammals have been identified in reptiles 119. While endogenous opioids and opioid receptors involved in reproduction and thermoregulation have been identified in reptiles, there is little known about the role of opioids in nociception 119-122. This information suggests that, at least at the physiological level reptiles are capable of responding to noxious stimuli in a manner similar to mammals, capable of nociception. The question of whether reptiles can “feel” pain or what significance pain or nociception have on physiological homeostasis is a far more complex question to answer. Until further evidence is available it seems that it would be most ethical to assume reptiles are capable of feeling pain and to treat or manage pain when there is reasonable evidence that pain is present. Interestingly, in a recent survey of the membership of the Association of Reptile and Amphibian Veterinarians, 98% of the respondents indicated their belief that reptiles do feel pain. However, only 39% of respondents in this survey reported using analgesics in > 50% or their patients 88. The reasons for failure to use analgesics were not specifically addressed in this study. However, some possibilities include a failure to recognize painful patients, lack of efficacy data, concern of adverse effects and little of no experimentally determined dose and pharmacokinetic information. The benefits of providing adequate analgesia are well recognized in mammals. The consequences of untreated pain are consistent with impaired patient homeostasis. These alterations can result in negative energy balance, lead to immune system compromise, inhibit healing and interfere with normal behavioural process required for health 123,124. The benefits of preemptive analgesia have also been demonstrated and can not only reduce postoperative pain by decreasing central sensitization but also may also facilitate healing and prevent and/or limit the actions of detrimental neurohumoral responses to pain 125,126. In addition, the use of analgesics as part of a balanced anesthetic protocol can reduce the doses of other anesthetics. This may help reduce the negative cardiopulmonary effects of general anesthesia. Overall it has been demonstrated in humans and other mammals that appropriate analgesia is an important part of complete medical care in health and disease. C. ANALGESIC THERAPY IN REPTILES As an extremely diverse group of animals, reptiles demonstrate a wide variation in inter- and intra-species behaviors. This makes the recognition of alterations in normal behavior that may be indicative of clinically significant pain and stress particularly difficult. Thus successful treatment of pain in reptiles demands an intimate knowledge of normal species-specific behaviors. In the absence of such knowledge, the delivery of appropriate analgesic therapy is based on an assessment of the likelihood of tissue trauma associated with a particular procedure. This recommendation is not new and was suggested by Flecknell in 1984 and Morton in 1986 127,128. There are three primary classes of analgesic drugs used in reptiles: local anesthetics, non-steroidal antiinflammatory drugs (NSAID’s) and opioids. Local anesthetics provide complete anesthesia by interrupting nociception from the level of the nociceptor to the spinal cord. NSAID’s act by modulating nociception in both the periphery the spinal cord. Opioids act by modulating nociception in the periphery, the spinal cord and supraspinal areas of the central nervous system. Since reptiles have a more primitive central nervous system, the central actions of analgesics medications, particularly opioids, may not be as predictable as the more peripherally-acting drugs. However, it is well documented that reptiles have opioid receptors in the central nervous system 119,120 and that the proopiomelanocortin system (one of the 3 molecular systems from which all naturally occurring opioids are derived) is well preserved among vertebrates 129,130. The unknown actions of opioids and NSAID’s in the central nervous system of reptiles may result in unpredictable variations in the duration, potency and side-effects of these drugs when the doses are determined by extrapolation from mammalian doses. Despite the unpredictable central effects of NSAID’s and opioids, their administration may offer the advantage of an increased duration of effect compared to that associated with the administration of local anesthetics. There are very few investigations that describe the assessment of analgesics in reptiles. The cardiopulmonary effects of several opioids have been studied in indigo snakes (Drymarchon corais couperi), bullsnakes (Pituophis catenifer sayi) and immature caiman (Caiman crocodilus) 131. In general, the administration of a variety of opioids to these species is not associated with significant changes in physiological parameters (heart rate, respiratory rate) or behavior (sedation or excitement). Morphine (0.05- 1.0 mg/kg IP) and meperidine (2-4 mg/kg IP) both induce statistically significant increases in response to a hot-plate test in crocodiles (Crocodylus niloticus africana) 115,116. A dose-

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dependent response is observed with both of these opioids. A ceiling for effect is observed following the administration of 0.3 mg/kg of morphine or 2 mg/kg of meperidine.In this species. The latency of onset of action is approximately 30 minutes and the duration of effect is 2 – 2.5 hours. The hot-plate test assesses thermal nociception which may not accurately reflect nociception associated with other stimulus modalities 132. See Table 5 for recommended doses of drugs that may be useful as analgesics in reptiles. Clinical Approaches to Analgesia in reptiles Assessing Pain in Reptiles Assessing pain in non-verbal species, including human infants, is an extremely challenging endeavor. Pain is defined as a sensory or emotional experience. Adult humans can express their individual level and significance of pain. In non-verbal humans and animals behavioral assessment tends to be the best indicator of pain 133-135. However, with over 8000 different reptile species identified that exhibit a wide range of unique physiological and behavior adaptations it is exceedingly difficult to assess behavior changes in these animals. This makes the identification of behavioural alterations that may be associated with pain particularly difficult. Recognition of abnormal behavior in reptiles requires careful, often time consuming observation and changes may be very subtle. An approach similar to pain assessment in other veterinary species can be adapted for use in reptiles (please see Table 4). If possible, it is probably best to observe reptiles remotely (using a remote camera) as there is evidence that reptiles may suppress some pain behaviours when the an observer is present 136. This may be a protective response similar to that seen in some reptiles subjected to brief physical restraint 114 and is likely a normal protective behavioural response to a perceived threat, similar to that found in many other vertebrate species 137. In a survey of reptile veterinarians it was found that the anticipated level of pain extrapolated from other species (76%), behavioural changes (66%), anticipated level of pain based on prior experience in reptiles (57%) and physiological changes (32%) were commonly used to evaluate pain in reptiles 88.

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An approach to pain assessment in reptiles A. BEHAVIOUR Species considerations Requires proper species identification and familiarity with species-specific behaviours. Basic species differences will impact behavioral patterns and these will be important when attempting to differentiate normal from abnormal behaviours. • Predominant activity pattern (diurnal, nocturnal) • Predated or predator species • Habitat (arboreal, aquatic, terrestrial, fossorial) Individual patient considerations • Stage of ecdysis o Some may become more aggressive during this time • Hibernation status o Hibernating animals or those inclined to hibernate may be more docile and less responsive than normal • Socialization o Altered response to human interaction (normally docile animal to biting), poor response to caregiver • Concurrent illness o Patient may be incapable of exhibiting behaviours associated with pain or behaviours associated with disease may be mistaken for pain behaviours • Owner assessment o Owners are often more familiar with their animals normal behaviour however owners may also be biased based on their own understanding and belief regarding their animals conditions Environmental considerations • Enclosure o Home enclosures often more “interesting”, compared to hospital enclosures, providing animal with plenty of opportunity to exhibit normal behaviours • Preferred optimal body temperature observer o Ambient environmental temperature is one of the main determinants of metabolic rate in resting reptiles and consequently normal behaviour may influenced by alterations in metabolic rate Locomotor activity • Posture o Hunched, guarding of affected body area, not resting in normal posture • Gait o Must differentiate neurological and mechanical dysfunction from pain induced lameness • Other o Excessive scratching or flicking foot tail or affected area o Unwillingness to perform normal movements (look up, step up, thrash with tail) o Exaggerated flight response Miscellaneous • Appetite o Reduced appetite may be related to underlying disease but may also be related to pain. • Eyes o May be held closed when painful or ill • Colour change o Species capable of colour change may do so in response to stress and/or pain • Abnormal respiratory movements o May be associated with primary respiratory disease but also pain affecting the muscles and tissues involved in respiration. B. ANTICIPATED LEVEL OF PAIN The anticipated level of pain is commonly used to evaluate pain in reptiles and is based on the likelihood and severity of tissue trauma associated with a particular procedure or condition. This is a well-accepted approach in veterinary medicine, particularly when dealing with less familiar species 127,128. However in addition to significant species differences, significant individual differences in response to therapy and response to tissue trauma can be seen. C. PHYSIOLOGICAL DATA Most physiological parameters have been shown to be relatively poor indicators of pain in most species. Physiological parameters can be influenced by disease and excitement. In addition, the physiological parameters of reptiles may be influenced by a number of metabolic processes such as activity level, temperature alterations and feeding. D. RESPONSE TO PALPATION In some species a negative response to palpation can be a useful indicator of pain. However, in reptiles this may be less sensitive as most reptiles will withdraw from touch regardless of whether the animal is experiencing pain. Analgesic Therapy in Reptiles

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A carefully designed analgesic plan should not only include the specific analgesic drugs, route of drug administration but it should also include steps to monitor patient response to therapy and address the provision of ongoing supportive patient care. Route of Drug Administration It is probably important to consider the route of drug administration in reptiles for many reasons including ease of drug administration, unique anatomical or physiological structures and variability in drug bioavailability and uptake among various routes of drug administration. Intramuscular drug administration is commonly used in reptiles. Historically, hind limb and tail injection sites have been avoided because of concerns related to the first-pass effect associated with passage of any administered drug through the kidneys via the renal portal system. However, at least in some species this may be more of a theoretical than practical concern as only a small amount of blood from the hind limbs and tail pass through the kidney 66,67 whereas in other species a small amount from the hind limbs pass through the renal portal system while substantial amounts of blood from the tail pass through the renal portal system 138. Regardless it may be best to avoid hind limb and tail administration of potentially nephrotoxic drugs or those highly metabolized or excreted by the kidneys. There is some recent evidence from the green iguana that the bioavailability of intramuscular ketoprofen may be reduced 139. In addition, time to effect may be variable depending upon the absorption rate of the drug from the site of deposition. Absorption may be affected by the physiochemical properties of the drug, tissue blood flow and cardiac output. Oral drug administration may be more desirable for patients who require chronic analgesic therapy, as most will become intolerant of repeated intramuscular injections. Oral drug administration can be difficult in some patients, although the use of feeding tubes can markedly simplify and facilitate this process. A more important consideration when contemplating oral drug administration may be the marked differences in gastrointestinal function among reptile species. Many are strict carnivores (snakes) and may fast for days to months between meals, while others are primarily herbivores (turtles, tortoises and some lizards) and tend to feed more or less continuously. These differences will presumably impact the bioavailability and pharmacokinetics among the different reptile species of drugs given orally. However, recently a study in green iguanas found that the bioavailability and pharmacokinetics of meloxicam given orally to be essentially the same as intravenous drug administration (Hernandez-Divers, personal communication). While intravenous drug administration is not always feasible in reptiles, the combination of good technique, practice, appropriate patient selection, and skilled physical restraint can facilitate predictable venous access. Intravascular injection ensures complete bioavailability of a drug and may avoid the tissue irritation associated with some drugs when given intramuscularly. Drugs There are three primary classes of analgesic drugs used in reptiles: local anesthetics, non-steroidal anti-inflammatory drugs (NSAID’s) and opioids. Opioids It is well documented that reptiles have opioid receptors in the central nervous system 119,120 and that the proopiomelanocortin system (one of the 3 molecular systems from which all naturally occurring opioids are derived) is well preserved among vertebrates 129,130. The importance of the different opioid receptors in modifying pain perception is far less clear. Opioids are the only class of analgesic drugs that have been assessed for clinical analgesic properties in reptiles. However, each study should be carefully evaluated for the quality of experimental design and strength of its conclusions. Based on recent studies, butorphanol, despite its frequent use 88, does not appear to produce significant analgesic effects in some reptiles 90,118 136. However, other studies suggest that butorphanol, at least at some doses, may reduce the intensity of motor reactions in response to an electrical stimuli applied to the tails of green iguanas 140. In the same study buprenorphine (0.02 and 0.04 mg/kg IM) did not significantly alter the motor reaction in response to an lectrical stimuli when compared to saline. There is evidence that morphine may be effective as an analgesic in at least some reptile species 115,116,118,140. It should be noted that time for onset of action appears to be prolonged (2-8 hours) following morphine administration and the duration of effect may vary considerably among species 115,116,118. This may be related to slow receptor binding kinetics or slow absorption from the intramuscular or subcutaneous injection site. Opioids appear to be safe for use in reptiles producing no discernable alterations in heart rate and behaviour (sedation or excitement) but do cause significant respiratory depression in some species 39,118,131. Local anesthetics

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Local anesthetics act by interrupting transmission of sensory and motor neurons. In reptiles local anesthetics are commonly used to facilitate minor surgical interventions but they can also be used as analgesics. Recently a technique has been described to block the mandibular nerve in crocodilians 141. The limited duration of analgesic effect and accompanying motor paralysis associated with local anesthetics limit their use primarily to the immediate peri-operative period or when hospitalized. Local anesthetic toxicity can be avoided by careful attention to total dose of local anesthetic administered to a patient. It must be kept in mind that many reptile patients are very small and large doses can accidentally be administered. In general, the toxic doses of local anesthetics in mammals (dogs) should not be exceeded; lidocaine (toxic dose 10-22 mg/kg) and bupivacaine (toxic dose 5 mg/kg) 142. Additionally, excessive dilution of local anesthetics may decrease their efficacy. Dilution of commercially available concentrations of local anesthetics should probably not exceed 50% on a per volume basis. A 1% lidocaine solution is available commercially and may be preferred for use in patients at greater risk for local anesthetic toxicity (i.e. small patients). NSAIDs The role of cyclo-oxygenase in the pathophysiology of pain and inflammation of reptiles has not been studied. However, reported clinical experience strongly supports the efficacy of NSAID’s in reptiles and they continue to be popular. Recent studies examining the pharmacokinetics of meloxicam in the green iguana showed excellent bioavailability of this drug given orally (Hernandez-Divers, personal communication). There is also some evidence of enterohepatic or possibly urinary resorption of meloxicam. The results suggest that plasma levels associated with analgesia are maintained for 24 hours after a single dose. However it should be noted that plasma levels of NSAID’s do not always correspond to their clinical effect and thus it is difficult to recommend effective and safe dosing intervals. A pilot examination of the pharmacokinetics of ketoprofen administered intravenously and intramuscularly in green iguanas has also been completed and revealed that bioavailability (78%) was decreased when administer intramuscularly and terminal half life was greater than comparable studies in dogs 139. Again this may suggest dosing intervals in reptiles should be greater compared to mammals, this recommendation is routinely followed for other drugs in reptiles, especially those associated with significant toxicity. Limited toxicity studies involving meloxicam were not associated with any clinically apparent abnormalities nor were there any histopathological lesions that could be associated with toxicity present at necropsy. However, mild biochemical and haematological abnormalities were noted that could not be clearly explained (Hernandez-Divers, personal communication). Thus until further studies in reptiles become available it is probably still best to consider the possibility that side effects similar to those seen in mammals (GI irritation, renal compromise, platelet inhibition) may occur in reptiles. Therefore hydration status, concurrent medications (steroids), presence of coagulopathy, GI disease and renal disease should all be addressed prior to administering these drugs. Other drugs Ketamine administered at sub anesthetic doses is being used as an analgesic in many mammalian species but its analgesic potential in reptiles has not been studied. Ketamine alone and used at anesthetic doses is associated with hypertension, tachycardia, bradypnea and hypoventilation 32,33,37. Ketamine may be a useful analgesic adjunct in select cases. The alpha2 agonists produce analgesia, sedation and muscle relaxation in mammals. In reptiles it appears to produce desirable levels of sedation and muscle relaxation. The analgesic effects of alpha-2 agonists have not been evaluated in reptiles but clinical impressions suggest it may be capable of producing an analgesic effect. Medetomidine induces cardiopulmonary effects in reptiles similar to those seen in mammals, bradycardia, hypertension and a reduction in arterial oxygen partial pressures. 84 85. Other analgesic drugs and adjuncts such as tramadol, gabapentin, amantadine, the tricyclic antidepressant and various forms of nutraceutical and physical therapy have not been explored in reptiles but may have a role to play as our understanding of nociception, pain and analgesic treatments increase in reptiles. Dosages of drugs with potential analgesic effects in reptiles

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Butorphanol* IM 1-8 mg/kg :May not be effective as an analgesic in reptiles 90,104,118,140,1 43 Buprenorphine* IM, IV, SC 0.4-1.0 mg/kg : May not be effective as an analgesic in reptiles 140,144 Morphine IC, IM 0.05-4.0 mg/kg (crocodiles) 1.5-6.5 mg/kg (turtles) 1.0 mg/kg (green iguanas) :Ceiling effect seen at 0.3 mg/kg in Nile crocodiles (Crocodylus niloticus africana) : Duration of action may persist for up to 24 hours in turtles 114,116,118,140 Meperidine IC 1-4 mg/kg: Ceiling effect seen at 2 mg/kg in Nile crocodiles (Crocodylus niloticus africana) 116 Ketamine IM, IV, SC 10-100 mg/kg: High doses are associated with anesthesia, Low doses <10 mg/kg likely associated with analgesia without sedation 32,33,37,73- 75,143 Xylazine* IM 1-1.25 mg/kg 144

Medetomidine IM, IV, IO 50-100 μg/kg (tortoises) 150-300 μg/kg (aquatic), 150 μg/kg (snakes and lizards): Lower doses may be effective for analgesia 68,84-87 Meloxicam IM, IV, PO 0.1-0.2 mg/kg q24-48 h#: Bioavailability and pharmacokinetics for oral and intravenous administration in green iguanas (Iguana iguana) 144, (Hernandez- Divers, personal communication) Carprofen* IM, IV, SC 2-4 mg/kg followed by 1-2 mg/kg q24-72 h# 73,143 Ketoprofen IM, SC 2 mg/kg q 24-48 h# 139,143 Flunixin meglumine* IM 0.1-0.5 mg/kg q 24-48 h# 73 Lidocaine (2%)* Local infiltration: Toxic dose unknown, recommend < 5 mg/kg: Dilute to 0.5% to increase volume 73,143 Bupivacaine (0.5%)* Local infiltration Toxic dose unknown, recommend < 2 mg/kg: Dilute to 0.25% to increase volume 72 * Doses not determined experimentally, extrapolated or anecdotal # Repeat dosing pharmacokinetics has not been studied and is based on extrapolation

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Conclusion Reptiles are a very unique and diverse class of animals that have developed distinctive mechanisms, not found in most other animals, for managing alterations in body temperature and metabolic rate. An approach to pain management based on our current understanding of reptile physiology, nociception, pain and analgesia represents a generalized approach to pain management in this class of animals. As our knowledge and understanding increases it is likely that our approach to pain management in this class of animals will also be modified and refined to more specifically address reptile pain. New information should be evaluated objectively and without the influence of personal bias or beliefs. It is likely that reptiles have evolved unique mechanisms for managing pain and avoiding the negative consequences associated with pain that we do not yet completely understand. REFERENCES 1. Ultsch GR, Jackson DC. Long-term submergence at 3 degrees C of the turtle Chrysemys picta bellii in normoxic and severely hypoxic water. III. Effects of changes in ambient PO2 and subsequent air breathing. J Exp Biol 1982;97:87-99. 2. Hicks JW, Bennett AF. Eat and run: prioritization of oxygen delivery during elevated metabolic states. Respir Physiol Neurobiol 2004;144:215-224. 3. Andrews RM, Pough FH. Metabolism of squamate reptiles: allometric and ecological relationships. Physiol Zool 1985;58:214-231. 4. Baker LA, White FN. Redistribution of cardiac output in response to heating in Iguana iguana. Comp Biochem Physiol 1970;35:253-262. 5. Hicks JW, Wang T. Hypometabolism in reptiles: behavioural and physiological mechanisms that reduce aerobic demands. Respir Physiol Neurobiol 2004;141:261-271. 6. Hicks JW, Wang T. Hypoxic hypometabolism in the anesthetized turtle, Trachemys scripta. Am J Physiol 1999;277:R18-23. 7. Ladyman M, Bradshaw D. The influence of dehydration on the thermal preferences of the Western tiger snake, Notechis scutatus. J Comp Physiol [B] 2003;173:239-246. 8. Van Mierop LHS, Kutsche M. Comparitive anatomy of the ventricular septum. In: wenick ACG, ed. The Ventricular Septum in the Heart. Boston: Martinus Nijhoff, 1981;35-46. 9. Van Mierop LHS, Kutsche M. Some aspects of comparative anatomy of the heart. In: Johansen K,Burggren WW, eds. Cardiovascular Shunts: Phylogenetic, Ontogenic and Clinical Aspects. Copenhagen: Munksgaard, 1985;38-56. 10. Herman J, Wang T, Smits AW, et al. The effects of artificial lung inflation on pulmonary blood flow and heart rate in the turtle Trachemys scripta. J Exp Biol 1997;200:2539-2545. 11. Comeau SG, Hicks JW. Regulation of central vascular blood flow in the turtle. Am J Physiol 1994;267:R569-578. 12. Hicks JW, Ishimatsu A, Molloi S, et al. The mechanism of cardiac shunting in reptiles: a new synthesis. J Exp Biol 1996;199:1435- 1446. 13. Heisler N, Neumann P, Maloiy GM. The mechanism of intracardiac shunting in the lizard Varanus exanthematicus. J Exp Biol 1983;105:15-31. 14. Ishimatsu A, Hicks JW, Heisler N. Analysis of intracardiac shunting in the lizard, Varanus niloticus: a new model based on blood oxygen levels and microsphere distribution. Respir Physiol 1988;71:83-100. 15. Hicks JW, Malvin GM. Mechanism of intracardiac shunting in the turtle Pseudemys scripta. Am J Physiol 1992;262:R986-992. 16. Luckhardt AB, Carlson AJ. Studies on the visceral sensory nervous system. Am J Physiol 1921;56:72-112. 17. White FN. Circulation In: Gans C,Dawson WR, eds. Biology of the Reptilia, Physiology A. New York: Academic Press, 1976;275- 334. 18. Burggren WW, Glass ML, Johansen K. Pulmonary ventilation: perfusion relationships in terrestrial and aquatic chelonian reptiles. Can J Zool 1977;55:2024-2034. 19. Milsom WK, Langille BL, Jones DR. Vagal control of pulmonary vascular resistance in the turtle Chrysemys scripta. Can J Zool 1977;55:359-367. 20. Berger PJ, Burnstock G. Autonomic nervous system In: Gans C, ed. Biology of the Reptilia. New York: Academic Press, 1979;1- 57. 21. Lillywhite HB, Donald JA. Pulmonary blood flow regulation in an aquatic snake. Science 1989;245:293-295. 22. Burggren W, Farrell A, Lillywhite HB. Vertebrate cardiovascular systems In: Dantzler WH, ed. Hanbook of Physiology Section 13: Comparative Physiology. New York: Oxford University Press, 1997;254-267. 23. Berger PJ. The reptilian baroreceptor and its role in cardiovascular control. Amer Zool 1987;27:111-120. 24. Stinner JN. Cardiovascular and metabolic responses to temperature in Coluber constrictor. Am J Physiol 1987;253:R222-227. 25. Stinner JN, Ely DL. Blood pressure during routine activity, stress, and feeding in black racer snakes (Coluber constrictor). Am J Physiol 1993;264:R79-84. 26. Farrell AP. Introduction to cardica scope in lower vertebrates. Can J Zool 1991;69:1981-1984. 27. Seymour RS, Lillywhite HB. Blood pressure in snakes from different habitats. Nature 1976;264:664-666. 28. Lillywhite HB, Gallagher KP. Hemodynamic adjustments to head-up posture in the partly arboreal snake, Elaphe obsoleta. J Exp Zool 1985;235:325-334. 29. Lillywhite HB, Pough FH. Control of arterial pressure in aquatic sea snakes. Am J Physiol 1983;244:R66-73. 30. Seymour RE. Scaling of cardiovascular physiology in snakes. Amer Zool 1987;27:97-109. 31. Bennett RA, Schumacher J, Hedjazi-Haring K, et al. Cardiopulmonary and anesthetic effects of propofol administered intraosseously to green iguanas. J Am Vet Med Assoc 1998;212:93-98. 32. Schumacher J, Lillywhite HB, Norman WM, et al. Effects of ketamine HCl on cardiopulmonary function in snakes. Copeia 1997:395-400. 33. Custer RS, Bush M. Physiologic and acid-base measures of gopher snakes during ketamine or halothane-nitrous oxide anesthesia. J Am Vet Med Assoc 1980;177:870-874. 34. Bonath K. Halothane inhalation anaesthesia in reptiles and its clinical control. International Zoo Yearbook 1979;19:112-125. 35. Rooney MB, Levine G, Gaynor J, et al. Sevoflurane anesthesia in desert tortoises (Gopherus agassizii). J Zoo Wildl Med 1999;30:64-69. 36. Stirl R, Bonath KH. Cardiovascular, pulmonary and acid-base measurements in boa constrictors during tiletamine-zolazepam sedation. 5th International Congress of Veterinary Anesthesia 1994;55.

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37. Arena PC, Richardson KC, Cullen LK. Anaesthesia in two species of large Australian skink. Vet Rec 1988;123:155-158. 38. Anderson NL, Wack RF, Calloway L, et al. Cardiopulmonary effects and efficacy of propofol as an anesthetic in brown tree snakes (Boiga irregularis). Bulletin of the Association of Reptilian and Amphibian Veterinarians 1999;9:9-15. 39. Mosley CA, Dyson D, Smith DA. The cardiovascular dose-response effects of isoflurane alone and combined with butorphanol in the green iguana (Iguana iguana). Vet Anaesth Analg 2004;31:64-72. 40. Milsom WK. Mechanoreceptor modulation of endogenous respiratory rhythms in vertebrates. Am J Physiol 1990;259:R898-910. 41. Smatresk NJ. Chemoreceptor modulation of endogenous respiratory rhythms in vertebrates. Am J Physiol 1990;259:R887-897. 42. Wang T, Smits AW, Burggren W. Pulmonary function in reptiles In: Gans C,Gaunt AS, eds. Biology of the Reptilia. Ithaca, New York: Society for the Study of Amphibians and Reptiles, 1998;319. 43. Wood SC, Lenfant CJM. Respiration: Mechanics, control and gas exchange. In: Gans C,Dawson WR, eds. Biology of the Reptilia, Physiology A. New York: Academic Press, 1976;225-274. 44. Shelton G, Jones DR, Milsom WK. Control of breathing in ectothermic vertebrates In: Geiger SR,Widdicombe JG, eds. Handbook of Physiology Section 3: The Respiratory System. Bethesda, MD: American Physiological Society, 1986. 45. Glass ML, Wood SC. Gas exchange and control of breathing in reptiles. Physiol Rev 1983;63:232-260. 46. Johansen K, Hanson D, Lenfant C. Respiration in a primitive air breather, Amia calva. Respir Physiol 1970;9:162-174. 47. White FN. Redistribution of cardic output in the diving alligator. Copeia 1969:567-570. 48. Shelton G, Burggren W. Cardiovascular dynamics of the chelonia during apnoea and lung ventilation. J Exp Biol 1976;64:323-343. 49. Perry SF. Structure and function of the reptilian respiratory system In: Lenfant C,Wood CM, eds. Comparative pulmonary physiology Current Concepts. New York: Marcel Dekker, 1989;193-236. 50. Glass ML, Johansen K. Control of breathing in Acrochordus javanicus, an aquatic snake. Physiol Zool 1976;49:328-340. 51. Templeton JR, Dawson WR. Respiration in the lizard Crotaphytus collaris. Physiol Zool 1963;36:104-121. 52. Glass ML, Burggren W, Johansen K. Ventilation in an aquatic and a terrestrial chelonian reptile. J Exp Biol 1978;72:165-179. 53. Jackson DC, Palmer SE, Meadow WL. The effects of temperature and carbon dioxide breathing on ventilation and acid-base status of turtles. Respir Physiol 1974;20:131-146. 54. Jackson DC, Kraus DR, Prange HD. Ventilatory response to inspired CO2 in the sea turtle: effects of body size and temperature. Respir Physiol 1979;38:71-81. 55. Boyer DR. Comparative effects of hypoxia on respiratory and cardiac function in reptiles. Physiol Zool 1966;39:307-316. 56. Jackson DC. Ventilatory response to hypoxia in turtles at various temperatures. Respir Physiol 1973;18:178-187. 57. Hitzig BM, Allen JC, Jackson DC. Central chemical control of ventilation and response of turtles to inspired CO2. Am J Physiol 1985;249:R323-328. 58. Benchetrit G, Armand J, Dejours P. Ventilatory chemoreflex drive in the tortoise, Testudo horsfieldi. Respir Physiol 1977;31:183- 191. 59. Benchetrit G, Dejours P. Ventilatory CO2 drive in the tortoise Testudo horsfieldi. J Exp Biol 1980;87:229-236. 60. Frankel HM, Spitzer A, Blaine J, et al. Respiratory response of turtles (Pseudemys scripta) to changes in arterial blood gas composition. Comp Biochem Physiol 1969;31:535-546. 61. Diethelm G. The effect of oxygen content of inspiratory air (FIO2) on recovery times in the green iguana (Iguana iguana). Zurich: University of Zurich, 2001. 62. Bertelsen MF, Mosley CA, Crawshaw GJ, et al. Inhalation anesthesia in Dumeril's monitor (Varanus dumerili) with isoflurane, sevoflurane, and nitrous oxide: effects of inspired gases on inducion and recovery. J Zoo Wildl Med 2005;36:62-68. 63. Berner NJ. Oxygen consumption by mitochondria from an endotherm and an ectotherm. Comp Biochem Physiol B Biochem Mol Biol 1999;124:25-31. 64. Penick DN, Paladino FV, Steyermark AC, et al. Thermal dependence of tissue metabolism in the green turtle (Chelonia mydas). Comp Biochem Physiol 1996;113A:293-296. 65. Heard DJ. Reptile anesthesia. Vet Clin North Am Exot Anim Pract 2001;4:83-117, vii. 66. Holz P, Barker IK, Burger JP, et al. The effect of the renal portal system on pharmacokinetic parameters in the red-eared slider (Trachemys scripta elegans). J Zoo Wildl Med 1997;28:386-393. 67. 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 1997;28:378-385. 68. Chittick EJ, Stamper MA, Beasley JF, et al. Medetomidine, ketamine, and sevoflurane for anesthesia of injured loggerhead sea turtles: 13 cases (1996-2000). J Am Vet Med Assoc 2002;221:1019-1025. 69. Wellehan JFX, Lafortune M, Gunkel C, et al. Coccygeal vascular catheterization in lizards and crocodilians. Journal of Herpetological Medicine and Surgery 2004;14:26-28. 70. Maxwell LK, Jacobson ER. Allometric scaling of kidney function in green iguanas. Comp Biochem Physiol A Mol Integr Physiol 2004;138:383-390. 71. Whitaker BR, Krum H. Medical management of sea turtles In: Fowler ME,Miller RE, eds. Zoo and Wild Animal Medicine. 4 ed. Philadelphia: W. B. Saunders, 1999;217. 72. Redrobe S. Anaesthesia and analgesia In: Girling S,Raiti P, eds. BSAVA Manual of Reptiles. Shurdington: British Small Animal Veterinary Association, 2004;131-146. 73. Malley D. Reptiles In: Seymour C,Gleed R, eds. Manual of Small Animal Anaesthesia and Analgesia. Shurdington: British Small Animal Veterinary Medical Association, 1999;271-281. 74. Glenn JL, Straight R, Snyder CC. Clinical use of ketamine hydrochloride as an anesthetic agent for snakes. Am J Vet Res 1972;33:1901-1903. 75. Cooper JE. Ketamine hydrochloride as an anaesthetic for East African reptiles. Vet Rec 1974;95:37-41. 76. Wood FE, Critchley KH, Wood JR. Anesthesia in the green sea turtle, Chelonia mydas. Am J Vet Res 1982;43:1882-1883. 77. Gray CW, Bush M, Beck CC. Clinical experience using CI-744 in chemical restraint and anesthesia of exotic specimens. J Zoo An Med 1974;5:12-21. 78. Boever WJ, Caputo F. Tilazol (CI 744) as an anesthetic agent in reptiles. J Zoo An Med 1982;13:59-61. 79. Clyde VL, Cardeilhac PT, Jacobson ER. Chemical restraint of american alligators (Alligator mississippiensis) with atracurium or tiletamine-zolazepam. J Zoo Wildl Med 1994;25:525-530. 80. Bienzle D, Boyd CJ. Sedative effects of ketamine and midazolam in snapping turtles (Chelydra serpentina). J Zoo Wildl Med 1992;23:201-204.

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81. Oppenheim YV, Moon PF. Sedative effects of midazolam in red-eared slider turtles (Trachemys scripta elegans). J Zoo Wildl Med 1995;26:409-. 82. Harvey-Clark C. Midazolam fails to sedate painted turtles (Chysemys picta). Bulletin of the Association of Reptilian and Amphibian Veterinarians 1993;3:7-8. 83. Holz P, Holz RM. Evaluation of ketamine, ketamine/xylazine, and ketamine/midazolam anesthesia in red-eared sliders (Trachemys scripta elegans). J Zoo Wildl Med 1994;25:531-537. 84. Sleeman JM, Gaynor J. Sedative and cardiopulmonary effects of medetomidine and reversal with atipamezole in desert tortoises (Gopherus agassizii). J Zoo Wildl Med 2000;31:28-35. 85. Dennis PM, Heard DJ. Cardiopulmonary effects of a medetomidine-ketamine combination administered intravenously in gopher tortoises. J Am Vet Med Assoc 2002;220:1516-1519. 86. Lock BA, Heard DJ, Dennis P. Preliminary evaluation of medetomidine/ketamine combinations for immobilization and reversal with atipamezole in three tortoise species. Bulletin of the Association of Reptilian and Amphibian Veterinarians 1998;8:6-9. 87. Greer LL, Jenne KJ, Diggs HE. Medetomidine-ketamine anesthesia in red-eared slider turtles (Trachemys scripta elegans). Contemp Top Lab Anim Sci 2001;40:9-11. 88. Read MR. Evaluation of the use of anesthesia and analgesia in reptiles. J Am Vet Med Assoc 2004;224:547-552. 89. Mauthe von Degerfeld M. Personal experiences in the use of association tiletamine/zolazepam for anaesthesia of the green iguana (Iguana iguana). Vet Res Commun 2004;28 Suppl 1:351-353. 90. Mosley CA, Dyson D, Smith DA. Minimum alveolar concentration of isoflurane in green iguanas and the effect of butorphanol on minimum alveolar concentration. J Am Vet Med Assoc 2003;222:1559-1564. 91. Brisbin LL. Reactions of the American alligator to several immobilizing drugs. Copeia 1966:129-130. 92. Messel H, Stephens DR. Drug immobilization of crocodiles. J Wildl Manage 1980;44:295-296. 93. Spiegel RA, Lane TJ, Larsen RE, et al. Diazepam and succinylcholine chloride for restraint of the American alligator. J Am Vet Med Assoc 1984;185:1335-1336. 94. Klide AM, Klein LV. Chemical restraint if three reptilean species. J Zoo An Med 1973;4:8-11. 95. Kaufman GE, Seymour RE, Bonner BB, et al. Use of rocuronium for endotracheal intubation of North American Gulf Coast box turtles. J Am Vet Med Assoc 2003;222:1111-1115. 96. Belkin DA. Anoxia: tolerance in reptiles. Science 1963;139:492-493. 97. Mosley CA, Dyson D, Smith DA. The cardiac anesthetic index of isoflurane in green iguanas. J Am Vet Med Assoc 2003;222:1565- 1568. 98. Bertelsen MF, Mosley CA, Crawshaw GJ, et al. Minimum alveolar concentration of isoflurane in mechanically ventilated Dumeril monitors. J Am Vet Med Assoc 2005;226:1098-1101. 99. Bertelsen MF, Mosley CA, Crawshaw GJ, et al. Anesthetic potency of sevoflurane and nitrous oxide in mechanically ventilated Dumeril's monitors (Varanus dumerili). J Am Vet Med Assoc 2005;In Press. 100. Maas A, Brunson D. Comparison of anesthetic potency and cardiopulmonary effects of isoflurane and sevoflurane in colubrid snakes. American Association of Zoo Veterinarians 2002;306-308. 101. Brosnan RJ, Pypendop BH, Barter LS, et al. Pharmacokinetics of inhaled anesthetics in green iguanas (Iguana iguana). Am J Vet Res 2006;67:1670-1674. 102. Barter LS, Hawkins MG, Brosnan RJ, et al. Median effective dose of isoflurane, sevoflurane, and desflurane in green iguanas. Am J Vet Res 2006;67:392-397. 103. Calderwood HW. Anesthesia for reptiles. J Am Vet Med Assoc 1971;159:1618-1625. 104. Schumacher J. Reptiles and amphibians In: Thurmon JC, Tranquilli WJ,Benson GJ, eds. Lumb and Jones' Veterinary Anesthesia. 3 ed. Philadelphia: Williams & Wilkins, 1996;670-685. 105. Millichamp NJ. Surgical techniques in reptiles In: Jacobson ER,Kollias GV, eds. Exotic Animals. New York: Churchill Livingstone, 1988;49-59. 106. Kaplan HM. Anesthesia in amphibians and reptiles. Fed Proc 1969;28:1541-1546. 107. Bello AA, Bello-Klein A. A technique to anesthetize turtles with ether. Physiol Behav 1991;50:847-848. 108. Murray MJ. Cardiology and circulation In: Mader DR, ed. Reptile Medicine and Surgery. Toronto: W. B. Saunders, 1996;95 103. 109. Bennett RA, Divers SJ, Schumacher J, et al. Anesthesia. Bulletin of the Association of Reptilian and Amphibian Veterinarians 1999;9:20-27. 110. Jarchow JL. Hospital care of the reptile patient In: Jacobson ER,Kollias GV, eds. Exotic Animals. New York: Churchill Livingstone, 1988;28-30. 111. Liang YF, Terashima S. Physiological properties and morphological characteristics of cutaneous and mucosal mechanical nociceptive neurons with A-delta peripheral axons in the trigeminal ganglia of crotaline snakes. J Comp Neurol 1993;328:88-102. 112. Stoskopf MK. Pain and analgesia in birds, reptiles, amphibians, and fish. Invest Ophthalmol Vis Sci 1994;35:775-780. 113. Gans C, Gaunt AS. Muscle architecture and control demands. Brain Behav Evol 1992;40:70-81. 114. Mauk MD, Olson RD, LaHoste GJ, et al. Tonic immobility produces hyperalgesia and antagonizes morphine analgesia. Science 1981;213:353-354. 115. Kanui TI, Hole K, Miaron JO. Nociception in crocodiles: Capsaicin instillation, formalin and hot plate tests. Zool Sci 1990;7:537-540. 116. Kanui TI, Hole K. Morphine and pethidine antinociception in the crocodile. J Vet Pharmacol Ther 1992;15:101-103. 117. ten Donkelaar HJ, de Boer-van Huizen R. A possible pain control system in a non-mammalian vertebrate (a lizard, Gekko gecko). Neurosci Lett 1987;83:65-70. 118. Sladky KK, Miletic V, Paul-Murphy J, et al. Analgesic efficacy and respiratory effects of butorphanol and morphine in turtles. J Am Vet Med Assoc 2007;230:1356-1362. 119. de la Iglesia JA, Martinez-Guijarro FI, Lopez-Garcia C. Neurons of the medial cortex outer plexiform layer of the lizard Podarcis hispanica: Golgi and immunocytochemical studies. J Comp Neurol 1994;341:184-203. 120. Reiner A. The distribution of proenkephalin-derived peptides in the central nervous system of turtles. J Comp Neurol 1987;259:65- 91. 121. Lindberg I, White L. Reptilian enkephalins: implications for the evolution of proenkephalin. Arch Biochem Biophys 1986;245:1-7.

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122. Ng TB, Hon WK, Cheng CH, et al. Evidence for the presence of adrenocorticotropic and opiate-like hormones in the brains of two sea snakes, Hydrophis cyanocinctus and Lapemis hardwickii. Gen Comp Endocrinol 1986;63:31-37. 123. Kona-Boun JJ, Silim A, Troncy E. Immunologic aspects of veterinary anesthesia and analgesia. J Am Vet Med Assoc 2005;226:355-363. 124. Muir WW. Pain and stress In: Gaynor J,Muir WW, eds. Handbook of Veterinary Pain Management. Toronto: Mosby, 2002;46-59. 125. Lascelles BD, Waterman AE, Cripps PJ, et al. Central sensitization as a result of surgical pain: investigation of the pre-emptive value of pethidine for ovariohysterectomy in the rat. Pain 1995;62:201-212. 126. Woolf CJ, Chong MS. Preemptive analgesia--treating postoperative pain by preventing the establishment of central sensitization. Anesth Analg 1993;77:362-379. 127. Flecknell PA. The relief of pain in laboratory animals. Lab Anim 1984;18:147-160. 128. Morton DB. Assessment of pain. Vet Rec 1986;119:435. 129. Zagon IS, Sassani JW, Allison G, et al. Conserved expression of the opioid growth factor, [Met5]enkephalin, and the zeta (zeta) opioid receptor in vertebrate cornea. Brain Res 1995;671:105-111. 130. Polzonetti-Magni A, Facchinetti F, Carnevali O, et al. Presence and steroidogenetic activity of beta-endorphin in the ovary of the lizard, Podarcis s. sicula raf. Biol Reprod 1994;50:1059-1065. 131. Hinsch H, Gandal CP. The effects of etorphine (M-99), oxymorphone hydrochloride and meperidine hydrochloride in reptiles. Copeia 1969:404-405. 132. Waterman AE, Livingston A, Amin A. Analgesic activity and respiratory effects of butorphanol in sheep. Res Vet Sci 1991;51:19-23. 133. Holton L, Reid J, Scott EM, et al. Development of a behaviour-based scale to measure acute pain in dogs. Vet Rec 2001;148:525- 531. 134. Pritchett LC, Ulibarri C, Roberts MC, et al. Identification of potential physiological and behavioral indicators of postoperative pain in horses after exploratory celiotomy for colic. Applied Animal Behaviour Science 2003;80:31-43. 135. van Dijk M, de Boer JB, Koot HM, et al. The reliability and validity of the COMFORT scale as a postoperative pain instrument in 0 to 3-year-old infants. Pain 2000;84:367-377. 136. Fleming GJ, Robertson S. Use of thermal threshold test response to evaluate the antinociceptive effects of butorphanol in juvenile Green iguanas (Iguana iguana). American Association of Zoo Veterinarians 2006;279. 137. Porro CA, Carli G. Immobilization and restraint effects on pain reactions in animals. Pain 1988;32:289-307. 138. Benson KG, Forrest L. Characterization of the renal portal system of the common green iguana (Iguana iguana) by digital subtraction imaging. J Zoo Wildl Med 1999;30:235-241. 139. Tuttle AD, Papich M, Lewbart GA, et al. Pharmacokinetics of ketoprofen in the green iguana (Iguana iguana) following single intravenous and intramuscular injections. J Zoo Wildl Med 2006;37:567-570. 140. Greenacre CB, Takle G, Schumacher J, et al. Comparative antinociception of morphine, butorphanol, and buprenorphine versus saline in the Green iguana, Iguana iguana, using electrostimulation. J Herp Med Surg 2006;16:88-92. 141. Wellehan JF, Gunkel CI, Kledzik D, et al. Use of a nerve locator to facilitate administration of mandibular nerve blocks in crocodilians. J Zoo Wildl Med 2006;37:405-408. 142. Liu PL, Feldman HS, Giasi R, et al. Comparative CNS toxicity of lidocaine, etidocaine, bupivacaine, and tetracaine in awake dogs following rapid intravenous administration. Anesth Analg 1983;62:375-379. 143. Bennett RA. Reptile anesthesia. Seminars in Avian and Exotic Pet Medicine 1998;7:30-40. 144. Malley D. Reptile anaesthesia and the practising veterinarian. In Practice 1997:351-368.

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