Anesthesia of reptiles (Proceedings)

General principles

Reptiles are very resilient and capable of surviving physiological changes e.g. severe hypoxemia, that would rapidly kill a mammal.

Hypothermia should not be used as a method of restraint. It will induce immobility but will not provide analgesia. Hypothermia will also impair drug metabolism, digestion and immune function.

For short non-painful procedures such as radiography, the vagal-vagal response can be used. Pressure over the eyes of most large lizards and crocodilians for about a minute will cause a decrease in heart rate and blood pressure and the reptile may become inactive enough for minor non-stimulating procedures.

In reptiles, administration of safe and effective anesthesia and analgesia presents unique challenges to the veterinary practitioner. Compared with mammalian and avian species, reptile anatomy and physiology differs considerably and knowledge of normal anatomy and physiology of the species to be anesthetized is essential. In particular, reptile respiratory anatomy and function is different from mammalian and avian species and affects response to anesthetic agents. In addition, the pathophysiology of many reptilian diseases is different from similar conditions described in mammals. The design of anesthetic and analgesic protocols in reptile patients is further complicated by species and individual differences in response to commonly used anesthetic and analgesic agents. Knowledge and understanding of reptilian anesthesia and analgesia has increased of the past decade, however relatively few studies have investigated the cardiopulmonary responses of reptiles to various anesthetic agents. Assessment of cardiopulmonary performance in reptiles is complicated by the fact that anesthesia monitoring devices (eg, pulse oximetry) commonly used in domestic animals are often of limited value in reptiles and have not been validated for use in reptilian species. Investigations into effective reptile analgesia have become more numerous and have shown species differences in response to analgesic agents and protocols. Effective management of preoperative, intraoperative, and postoperative pain in reptiles is important to eliminate the negative impact of pain on many body functions including the immune system and cardiopulmonary system.

Preanesthetic evaluation

A detailed history of the patient should be obtained including environmental conditions as well as husbandry practices. Many disease problems diagnosed in reptiles are directly associated with inadequate husbandry such as improper nutrition and failure to provide the animal with appropriate temperature and humidity conditions. A complete visual and physical evaluation of the patient is essential. Abnormalities should be recorded while particular attention should be paid to the cardiopulmonary system. Rate and depth of respiration should be evaluated for signs of respiratory tract disease. Baseline heart and respiratory rates should be recorded. Often, reptiles are presented with chronic disease processes and physical and laboratory findings indicating dehydration, malnutrition and presence of secondary bacterial infections are common. If possible, a venous blood sample should be collected for determination of hematological and plasma biochemical parameters. Minimally, a blood sample should be collected for determination of blood glucose, total protein and packed cell volume. Based on physical examination and laboratory findings, abnormalities and deficiencies should be treated prior to anesthesia. The patient should be placed in a temperature and humidity controlled environment. Small animal incubators are ideal for this purpose and supportive measures including fluid therapy, nutritional support and antimicrobial therapy should be instituted. The patient should also be evaluated for any signs of pain and discomfort and if indicated, analgesic therapy should be initiated. For elective surgical procedures were postoperative pain is anticipated, administration of pre-emptive analgesia is most effective for the management of intraoperative and postoperative pain.

Pre-anesthetic preparation

Whenever possible, the patient should have a thorough physical examination and their medical history should be reviewed before induction. An accurate weight is very important and the shell weight is included in body weight of chelonians.

The reptile should be physiologically stable before anesthesia and whenever possible, abnormalities such as dehydration, anemia and acid-base disturbances should be corrected. It should be maintained within its preferred optimal temperature range at all times.

Ideally reptile anesthesia should be performed in the morning so that if recovery is prolonged, the animal can be monitored and if necessary, ventilated during regular working hours rather than late at night.

Regurgitation and aspiration are rarely a problem but pre-operative fasting is recommended due to the animal's impaired digestion.

Pre-medication

The prolonged clearance of most parenteral drugs, and very variable effects of sedatives limit their use. µ2 adrenergic agonists (e.g. medetomidine and xylazine) alone give minimal sedation but when combined with ketamine can produce chemical immobilization. Atropine may be used to treat bradycardia but is not indicated as a premedicant to reduce oral secretions. Opioids may be given preoperatively for analgesia, although the efficacy is not known.

Local anesthesia

The toxic doses of local anesthetics have not been investigated in reptiles but in mammals, it varies from 5-20 mg/kg for lidocaine. In small patients this dose could easily be exceeded unless care is taken to calculate an appropriate dose. Lidocaine can be used to desensitize the glottis prior to endotracheal intubation.

Induction of anesthesia

When intravenous injection is not possible, dissociative drugs such as ketamine may be given IM to induce sedation or anesthesia. However, recovery from ketamine anesthesia can be extremely prolonged (days with high doses!). Combination of medetomidine with ketamine can reduce the requirement of ketamine and reversal of the medetomidine with atipamezole then allows faster recovery. The effective doses of this combination have not yet been determined for many species but the following are recommended by Heard (2001):

     • Tortoises: 50-100mcg/kg medetomidine + 5-10mg/kg ketamine IM

     • Giant Tortoises: 40-60mcg/kg medetomidine + 4-6mcg/kg ketamine IM

     • Turtles: 50-300mcg/kg medetomidine + 10-15mcg/kg ketamine IM

     • Lizards: 150mcg/kg medetomidine + 10-15mcg/kg ketamine IM, IV or IO.

Wait 45 minutes for full effect and reverse medetomidine with atipamezole at 3 times the dose of medetomidine.

When intravenous injection is possible in the awake animal, propofol (3-5mcg/kg IV or IO) is probably the induction agent of choice. The main advantage is its short duration of action and it is generally used to allow endotracheal intubation before maintenance on gaseous anesthesia.

Induction of anesthesia can also be achieved using gaseous anesthesia. The preferred inhalation agent is isoflurane or sevoflurane. Standard cat and dog facemasks can be used to induce larger lizards and syringe cases may be used as facemasks for smaller patients. Induction chambers may also be used for induction of e.g. venomous snakes but induction will be very prolonged. This method is not suitable for chelonia, particularly aquatic species which can convert to anaerobic metabolism for prolonged periods. The glottis of snakes is very obvious and endotracheal intubation is simple even in the conscious animal (see below). Therefore intubation (following lidocaine application to the glottis) and ventilation of the conscious reptile is frequently used to induce non-venomous snakes.

Maintenance of anesthesia

Whenever possible, maintenance of anesthesia is by gaseous anesthetics. Supplemental injections of ketamine or propofol could be used but the effects are variable and recovery could be significantly prolonged.

Cardiopulmonary performance of the patient should be closely monitored throughout the anesthetic event since all anesthetic agents have cardiopulmonary depressant effects. Fluid therapy is necessary for maintenance requirements or correction of fluid deficits. For maintenance, 5 -10 mL/kg/hr of a balanced electrolyte solution is administered. For small species syringe pumps are ideal since they allow for correct administration of fluids and continuous rate infusions which are superior to bolus injections. Throughout anesthesia, the reptile should be kept within the preferred temperature range of the species. Hypothermia is commonly seen in anesthetized patients and supplemental heat should be provided with heat lamps or heating blankets. During surgery, the patient should be regularly evaluated for signs of pain (movement, increase in heart and respiratory rate) and if necessary additional analgesic agents should be administered.

Intermittent positive pressure ventilation is generally required to ensure adequate gaseous exchange. This is because reptiles do not have a functional muscular diaphragm and in most reptiles, the muscles of the abdomen and trunk augment the function of the intercostal muscles. In chelonia, negative pressure in the lungs is achieved by movement of the limbs. At surgical planes of anesthesia, skeletal muscle movement will be lost so assisted ventilation is required. Also, when the patient is placed in dorsal recumbancy, the viscera may compress the simple, sac-like lungs. Ideally a ventillator properly programmed for the particular species should be implemented - though proper monitoring by an assistant should never be forsaken.

Endotracheal intubation and ventilation

As stated above, this will be necessary in virtually all reptile anesthetics and is generally easily accomplished. In all reptiles, the glottis is closed between breaths and the anesthetist must either be patient and wait for it to open or may gently force the glottis open with the bevel of the endotracheal tube. Intubation may be made easier after the application of local anesthetic to the glottis. The glottis lies at the base of the tongue and is obvious in snakes and most lizards. Chelonians have a thick fleshy tongue which can obscure visualization of the glottis and crocodilians have a well developed epiglottis (basihyal valve) which must be depressed in order to identify the glottis for intubation. Chelonia have a relatively short trachea which bifurcates shortly after entering the coelomic cavity.

For very small species, 18-20G intravenous catheters or urinary catheters may be used as ET tubes. A ventilation rate of 2-4 breaths / minute is usually adequate and enough time should be allowed for the patient to exhale through very small diameter tubes. Excessive ventilatory pressure should be avoided to prevent rupture of the lungs i.e. just enough to move the body wall in snakes, lizards and crocodilians or just enough to move the limbs in chelonia.

Anesthetic monitoring

In order to determine anesthetic depth, the presence or absence of reflexes is commonly determined. In Reptiles & Amphibians under the surgical plane of anesthesia the righting reflex will be usually be absent. Absence of the corneal and tongue withdrawal reflex often indicates too deep a level anesthetic plane.

Anesthetic depth

Assessing the depth of anesthesia can be very difficult in reptiles. The degree of muscle tone (jaw, cloacal) and response to toe, tail or cloacal pinch or to surgery are used to assess depth. Palpebral and corneal reflexes (in species with eye lids) may be monitored, although corneal reflex and cloacal tone are not usually lost at surgical anesthetic levels. Respiratory rate and depth are not usually useful because most reptiles become either apneic or markedly bradypneic at surgical levels of anesthesia. The heart rate increases in response to painful stimuli and a sudden increase is often indicative of inappropriate anesthetic depth.

Cardiovascular monitoring

In order to monitor the heart, it is necessary to know its position. In lizards, the position of the heart varies from between the forelimbs (e.g. iguanas, chameleons, skinks) to almost mid-body (monitors and tegus). Snake hearts are usually located 20-25% of the distance from head to tail. External auscultation is not usually very rewarding but esophageal stethoscopes are useful and placement of a Doppler flow detector directly over the heart of lizards and snakes will allow monitoring of blood flow. Esophageal stethoscopes are also useful in chelonia and the Doppler probe is usually placed in the thoracic inlet in these species.

Electrocardiography can be used for monitoring the heart rate although lead position may need to be adjusted to obtain and adequate trace. The electrodes are placed in the cervical region for lizards with hearts located in the pectoral girdle and in snakes they are placed two heart lengths cranial and caudal to the heart. In tortoises and turtles the cranial leads are placed on the skin between the neck and the forelimbs.

Pulse oximetry is generally not reliable in reptiles. Cloacal / esophageal probes may be tried but it is generally difficult to obtain a consistent reading. Although not validated for the use in reptiles, pulse oximetry is a useful tool to monitor trends in relative arterial oxygen saturation. Both transmission and reflectance probes can be used. In most species an esophageal reflectance probe can be placed at the level of the carotid artery. Arterial blood gas analysis is challenging in reptiles since a peripheral artery is not readily available. Interpretation of arterial blood gas parameters should be done with caution since arterial blood gas analyzers are calibrated based on the human oxygen hemoglobin dissociation curve. Values should be interpreted at 37°C since correction of arterial blood gas parameters to the patient's temperature will further compromise the accuracy of obtained values. Both, pulse oximetry and arterial blood gas analysis may provide valuable data to assess trends in cardiopulmonary performance. Capnography is of limited value in reptiles because they can develop cardiac shunts. However, changes in end-tidal CO2 concentrations may indicate complications such as airway leaks and airway obstruction. As mentioned above the reptile should regularly be evaluated for any evidence of pain during surgery. Additional administration of systemic analgesic agents as well as local anesthetic agents (eg, bupivacaine) may be indicated.

All reptiles will exhibit profound respiratory depression following induction of anesthesia as well as during the maintenance phase. Consequently, as previously stated, it is essential to assist ventilation and provide intermittent positive pressure ventilation (IPPV). Small animal ventilators (pressure or volume driven) can effectively be used in reptiles. Although no studies have determined safe and effective ventilation in reptiles it is recommended to administer IPPV between 4 to 8 breaths/minute. Peak airway pressure should not exceed 10 to 15 cm H2O.

Fluid administration

Dehydration should preferably be corrected before the induction of anesthesia (see critical care lecture). Intra-operative fluids can be given at the rate of 5-10 ml/kg/hr to maintain vascular access and replace fluid loss.

Recovery

This will normally occur more slowly than in mammals and the endotracheal tube is left in position until spontaneous respiration or attempts by the patient to remove the tube are made. The reptile is ventilated with room air to speed the return of spontaneous breathing. The patient is maintained in its preferred temperature range. Recovery will be significantly prolonged if anesthesia was achieved using ketamine or similar drugs.

During the recovery period the patient should be placed in a quiet, temperature controlled environment. Small animal incubators are ideal for this purpose since they allow visualization of the patient without disturbing or handling it. Temperature in the incubator should be set within the preferred body temperature range of the particular species. Increasing the temperature above this range is not recommended since it will increase metabolism and demand of the tissues for oxygen which may not be met by a patient with some degree of respiratory compromise following anesthesia. Reptiles should be closely monitored for evidence of respiratory depression and ventilation may have to be assisted well into the recovery period. Only fully recovered animals should be returned to their enclosure. Following a surgical procedure the reptile should also be regularly evaluated for any evidence of postoperative pain such as increased aggressiveness or abnormal posture. If needed, additional analgesics should be administered and the analgesic protocol should be re-evaluated. It may be necessary to combine different analgesic agents to provide the most effective pain relief for the patient.

Analgesia

While all vertebrates experience pain, there may be different pathways and receptors between mammals and reptiles.⁴ Prevention and treatment of pain is essential in order to provide state of the art care for the reptilian patient. An effective analgesic protocol should be in place for any surgical or painful procedure. Although information on effective analgesic drugs and dosages is increasing, it is well established that there are pronounced species and individual differences in response to analgesic agents. Also, effective dosages and treatment intervals vary considerably between species. Recognition of pain in reptiles may be challenging and it is essential to be familiar with the species to be treated and its normal behavior. Clinical signs of pain include abnormal body position, reluctance to move or to lie down, restlessness, anorexia, increased aggression, increased respiratory rate and trembling.

The most effective method of pain management is prevention of pain. Local anesthetic techniques are commonly used in reptiles for minor procedures (eg, abscess debridement, mass removal). Techniques for local anesthesia described for mammalian species can also be applied to reptiles, although consideration must given to anatomic differences. Topical, regional and local infiltration techniques can be used in many reptile species. Similar to mammalian species, pre-emptive analgesic techniques are recommended in reptiles and will include pre-operative administration of analgesics such as opioid agents. Opioid agents (eg, butorphanol, buprenorphine) are most effective in the treatment of acute pain. Administration of these agents prior to a noxious stimulus will often provide effective preemptive analgesia and will also provide various degrees of sedation prior to induction of anesthesia. Balanced analgesic regimen have been described in domestic animals and should also be administered to reptiles.5 A combination of an opiod agent (eg, butorphanol (0.4-2 mcg/kg) or buprenorphine (0.02-0.2 mcg/kg)) with a long-acting local anesthetic (eg, bupivacaine 1-2 mcg/kg) agent is often more effective in relieving pain then the use of either one of these drugs alone. While chronic pain is often neglected in reptiles, many patients are presented with chronic disease processes associated with pain and discomfort including renal disease and metabolic bone disease. Nonsteroidal anti-inflammatory agents (NSAIDs) can be used in reptiles; however, information on effective drugs and dosages is scant. Meloxicam (0.1-0.2 mcg/kg PO) has been used successfully for the management of chronic pain as well as postoperative pain in reptiles.

References

Bennett RA. Reptile Anesthesia. In Mader DR (ed) Reptile Medicine and Surgery. WB Saunders Co, Philadelphia, Pennsylvania. pp 241-247, 1996.

Heard DJ. Reptile Anesthesia. Veterinary Clinics of North America: Exotic Animal Practice. Vol 4, No 1, January 2001, pp 83-117.

Lawton MPC. Anesthesia. In PH Benyon, NA Forbes and MPC Lawton (eds) BSAVA Manual of Psittacine Birds. BSAVA, Cheltenham, UK. 1996 pp 49-59

Greenacre CB, Takle G, Schumacher J, Klaphake EK, Harvey RC. 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.

Robertson SA. Analgesia and analgesic techniques. Vet Clin North Am Exotic Anim Pract. 2001;1-18.