Introduction

Exotic animals are routinely maintained as pets, in laboratories and zoological settings. Anesthesia of these species has improved rapidly over the last several decades with new drugs, monitoring equipment and methods of drug delivery. Their unique anatomy, physiology, and species differences make the application of anesthesia and analgesia challenging. Despite these challenges, successful anesthesia can be achieved by following a consistent protocol, including a thorough preanesthetic examination, careful anesthetic monitoring and appropriate post-anesthesia care. It is essential that veterinarians caring for exotic animals be familiar with appropriate anesthetic techniques to facilitate treatment, diagnostics, surgery and research.

Preanesthetic Assessment

Make every effort to evaluate patients prior to physical examination and anesthesia. Non-domestic species have evolved to depend on their ability to hide illness to survive making it more difficult for owners to detect medical problems. They are often significantly compromised at the time of presentation, increasing anesthetic risk. Physical examination, blood work and diagnostic imaging should be used to evaluate the patient's health status. Species differences, health, age and size influence anesthetic safety and must be taken into account. Defensive reactions in many exotic animals are common and include escape reflex, inflation, micturition, defecation, biting, vocalization, writhing, rolling and tail autonomy.

Supportive Care

Animals should be kept at their preferred optimal temperature zone (POTZ) throughout anesthetic procedures. The need for supplemental oxygen, nutrition and fluid therapy should be assessed pre and post anesthesia.

Anesthesia

Drug administration in exotic patients includes intravenous (IV), intramuscular (IM), intracoelomic (ICe), intraosseous (IO), oral (PO), topical (TO) and inhaled methods. Premedication minimizes discomfort and stress of induction, and facilitates catheter and endotracheal tube (ET) placement. Animals should be intubated when possible and trends in heart rate (HR), respiratory rate (RR), response to reflexes and discomfort closely monitored over time. Auscultation or Doppler evaluation of pulse and blood flow are extremely important as ECG leads are difficult to attach and readings can be misleading as cardiac activity can continue after death in ectotherms. Pulse oximetry, capnography, blood pressure and temperatures probes can be useful and should be used when applicable. Patients should be monitored closely during recovery to prevent self and intraspecific trauma until they are alert and responsive. Recovery in hypothermic or ectothermic patients is often prolonged. Some require ventilation, supplemental heat, stimulation, and analgesia.

Fish

Anesthesia facilitates examination, transport, obtaining diagnostic samples and is necessary for surgery. Waterborne anesthesia is the most widely used route of anesthetic administration for fishes.³ Many biologic factors such as species, age, sex, body condition as well as water quality, temperature and oxygenation affect the efficacy of fish anesthesia.⁷ Food should be withheld (for one feeding cycle) as regurgitated food can lodge in the gills and degrade the water quality. The water with the anesthetic should be of good quality, match closely the parameters from the tank or pond and be well oxygenated prior to use. Most fish depend on oxygen exchange across their gills for respiration. Pathology affecting the gills increases risk of anesthesia and warrants additional precautions. Anesthetic stages include sedation, narcosis/loss of equilibrium, and anesthesia.³

Multiple compounds have been used as fish anesthetics but the most common agents used in practice are tricaine methanesulfonate (MS-222, Finquel®) and clove oil (eugenol).^1,7 Tricaine is effective in all species and has a wide margin of safety. It comes as a powder and can be made into a stock solution or added directly into tank water. When reconstituting tricaine, it is best to use water from the fish's tank system to ensure consistency with water quality parameters. Tricaine is very acidic and must be buffered to maintain an appropriate pH. This can generally be accomplished by using sodium bicarbonate (baking soda) at equal parts on a dry matter weight to the tricaine. In general, dosages of 50–250 mg/L work for many commonly kept pet fish. Anesthesia may be improved in some cases by inducing with a higher dose (100–150 mg/L) and maintaining anesthesia with a lower dose (50–75 mg/L). First effects are noted within 5–30 minutes. For longer procedures, an anesthetic system with a recirculating pump that allows water to flow over the gills is needed.

Doppler flow detectors works well to monitor HR and rhythm. RR can be counted by observing the movement of the operculum. If respiration ceases the anesthetic concentration should be reduced or stopped and replaced with unmedicated water until the fish resumes opercular movement.

Recovery is achieved by placing the fish in anesthetic free water. As the fish recovers, RR increases, fins start to move and the fish swims with progressively better coordination. Most recover within 5–10 minutes after being placed in clean water.

Amphibians

Amphibians, in their aquatic form, have gills and respiration similar to fish. As adults they develop lungs and can respire out of water. Larval tadpoles can be immersed in an anesthetic, but it is very important to protect the airway of terrestrial adults to prevent drowning. Amphibian skin is semipermeable and can be used to administer drugs as well as oxygen.⁴ Handlers should wear moistened disposable gloves to protect the amphibian's sensitive integument and as personal protective gear to avoid exposure to secretions.¹⁰

Amphibians are poikilothermic and should be kept at their POTZ (~ 15–23°C, 59–73°F) to maintain metabolic needs and ensure proper drug absorption, metabolism and excretion.⁸ Aspiration of stomach contents is generally not a concern but it is best to fast amphibians to prevent complications secondary to post-anesthetic ileus. The amphibian heart is composed of a single ventricle and two atria, allowing varying degrees of mixing of oxygenated and deoxygenated blood as well as cardiovascular shunting which can result in slower than expected induction or recovery periods.^4,10 Respiration can occur by pulmonic, buccopharyneal and cutaneous mechanisms making amphibians more resistant to hypoxia associated with apnea. ^4,10 Intubation is more difficult and when performed should be done with non-cuffed tubes. The glottis is located at the base of the tongue, followed by a very short trachea and paired fragile saclike lungs.

Buffered tricaine is the anesthetic recommended for most procedures and prepared as for fish. Varying concentrations are used depending on the level of anesthesia desired, life stage, stability of the patient, as well as species specific drug sensitivity. Younger life stages (0.1–0.5 g/L) and aquatic frogs (1–2 g/L) require lower dosages than terrestrial toads (2–3 g/L) for induction. Maintenance doses of 0.1–0.2 g/L often suffice to maintain appropriate anesthetic depth for many procedures. Anesthetic should cover approximately half the patient, ensuring nostrils, are kept above water as sedation ensues. First effects are generally noted within 15–30 min. Induction rates are faster in smaller animals and those with gills.^8,10 When the desired anesthetic state has been attained, the amphibian should be removed from the induction solution. In most cases the patient will remain anesthetized long enough to complete the necessary procedures. If the animal becomes light, a tricaine solution, 50% of the induction strength, can be applied to the patient to adjust the anesthetic depth as needed.

HR is easiest to monitor with Doppler devices placed over the heart. ECGs can be used but the myocardium can continue beating after death and readings do not reflect cardiac output. Marked decreases in HR (> 20%) warrant attention and removal or reduced concentrations of anesthetic solutions.

Amphibians often exhibit an excitement phase during induction. As they become more deeply anesthetized, gular respiratory movements decrease, withdrawal and palpebral reflexes diminish and finally the righting reflex is lost. Marked respiratory depression with complete loss of gular and pulmonary ventilation occurs when animals reach a surgical plane of anesthesia. Respiratory drive may also cease under hyperoxic conditions and room air can speed the return of voluntary ventilation.⁸ Amphibians ability to respire transdermally allows them to tolerate apnea for extended periods.¹⁰ Pulse oximetry can be used in larger patients and decreases of > 5% SpO₂ warrant reevaluation. Recoveries are prolonged compared to other animals with withdrawal reflex and gular respirations returning first followed by the righting reflex.

Reptiles

Reptiles like amphibians, are poikilothermic, and changes in temperature affect many physiologic processes. Patients should be maintained at their POTZ (usually 20–25°C) to ensure proper drug absorption, metabolism and excretion.⁵ Chelonians, lizards, and snakes have a cardiopulmonary system similar to amphibians. Oxygenated and deoxygenated blood can mix and be shunted to bypass the lungs during apneic periods. This may delay uptake or exhalation of inhalation agents resulting in slower than expected induction or recovery periods. Reptiles have a renal portal system that is able to direct blood flow through or bypass the kidneys to the caudal vena cava and liver. For this reason parenterally administered drugs should be given in the cranial half of the body. They lack a diaphragm and rely on movement of abdominal muscles to move air during inspiration and expiration. The glottis of snakes is rostrally located and easily visualized for intubation. In lizards and chelonians the glottis is at the base of the tongue which can be very fleshy in some species which makes them more challenging to intubate. Non-cuffed tubes are recommended for all reptiles but especially for chelonians, as they have complete tracheal rings. Lizards and Chelonia have paired, saclike lungs whereas most snakes have a vestigial left lung and functional right lung that ends in a terminal air sac.

Lizards and snakes tend to require lower dosages of anesthetics than chelonians. Sedation and premedication with α2-adrenergic agonists and benzodiazepines with or without low doses of ketamine (dexmedetomidine 70–100 mcg/kg, midazolam 1–2 mg/kg, ketamine 1–5 mg/kg SC/IM) provide good results. Opiods can be used alone or in combination with above mentioned agents and are indicated prior to painful procedures (medetomidine 100 mcg/kg, midazolam 2 mg/kg, morphine 1 mg/kg SC/IM).⁶ Both isoflurane and sevoflurane are common induction and maintenance anesthetic agents. Mask, tube or chamber induction is possible but often results in apnea, vascular shunting and can be prolonged. Direct intubation allows for quicker induction but can be stressful to the patient. Propofol (1–10 mg/kg IV/IO) is useful for induction, as an aid to intubation and for non-painful procedures. Although it requires IV administration, it provides rapid effects and short recoveries compared to many other agents. Rare complications have been reported and it is recommended that injections be given into the jugular or brachial plexus.⁶ Historically used high doses of parenteral agents result in prolonged recovery, are not reversible and do not provide clinical benefits over agents discussed above.

Most reptiles lose the ability for spontaneous respiration when they reach a surgical plane of anesthesia and should be ventilated at 1–2 breaths per minute. Muscle tone, righting reflex, spontaneous respiration, jaw tone, palpebral and corneal reflexes and response to painful stimuli assist with gauging the depth of anesthesia in reptiles. Spontaneous arousal, particularly when using solely inhalant anesthesia, is common and likely related to the ability for these species to shunt blood away from the lungs. Manual ventilation ± propofol boluses are especially helpful in managing these occurrences.

Doppler devices, ECGs, audible respiratory monitors and indwelling temperatures probes are useful to monitor and assess reptile patients under anesthesia. As with amphibians, ventilation with room air is recommended during recovery to stimulate voluntary respiration.

Birds

Birds have air sacs that act as bellows to the lungs but are not involved with gas exchange. In emergencies involving tracheal obstruction, air sac cannulation can be used to provide ventilation. They have a highly efficient crosscurrent flow of air and blood, with continuous gas exchange along parabronchi. Respiratory muscles are used for both inspiration and expiration and birds, like reptiles do not have a diaphragm. Renal portal circulation may reduce the efficacy of anesthetic drugs administered in the caudal half of the animal's body including the legs.

Avian patients mask illness particularly well and are more likely to develop handling stress related complications. Preoxygenating patients when necessary and ensuring efficiency with procedures that require anesthesia is essential. Supplemental heat should be considered for procedures that last longer than 5–10 minutes. Length of preanesthetic fasting needs to be determined based on the size and metabolism of individual patients.

Inhalants, isoflurane and sevoflurane, are preferred for anesthetic induction and maintenance.² Most birds are best masked down using a face mask. Birds have complete tracheal rings and should be intubated with non-cuffed endotracheal tubes.

Physiologic parameters including HR, RR and body temperature should be closely monitored and recorded. If apnea develops, discontinue anesthetic and ventilate with oxygen while reevaluating cardiac function. ECG, Doppler and audible respiratory monitors are useful in birds to help detect changes in heart rate, blood flow and ventilation respectively. Noninvasive blood pressure and pulse oximetry are more difficult to employ reliably due to the small size and anatomy in these patients. To prevent self-trauma during recovery it is best to hold birds in a towel until they are able to stand/perch. Appropriate perianesthetic analgesia, fluid and nutritional support need to be considered for each avian patient.

Small Mammals

Rodents, rabbits, and ferrets are some of the most common exotic pets. Their small body size makes venous access, muscular injection and tracheal intubation more difficult. They have a high metabolic rate and this in particular must be taken into account for preanesthetic preparation and drug dosing. Fasting is typically not necessary with rodents and rabbits since vomiting does not occur in these species. Ferrets should only be fasted 1–3 hours due to their fast intestinal transit time. Rodents and ferrets are prone to hypoglycemia and may require parenteral dextrose during long procedures. Preexisting respiratory disease is common and supplemental oxygen during induction and recovery may be desirable.⁹

Injectable and inhalant anesthetics are both commonly used in these species depending on species, temperament, health status, and availability. Injectable agents can be given via IM, IV, SQ or IP routes. Intravascular and IM access is less practical in rodents and IP injections may be more reliable.

Neuroleptanalgesic combinations (fentanyl, morphine or buprenorphine + midazolam) are useful for premedication/induction. Medetomidine combined with other agents has also been useful for maintenance (medetomidine + ketamine ± midazolam, medetomidine + butorphanol) of anesthesia. Midazolam used alone (0.5–2.0 mg/kg IM) can provide adequate sedation to facilitate sample collection, diagnostic imaging and treatment.⁹ Propofol may be used in animals with IV access. The author prefers to use a combination of injectable premedication followed by inhalant agents in ferrets and rabbits and more commonly uses inhalants (isoflurane, sevoflurane) via mask or chamber for both induction and maintenance in rodents. Anticholinergic drugs, such as atropine and glycopyrrolate are not routinely used.

Intubating rabbits and ferrets can be challenging. To facilitate intubation in these species a long narrow laryngeal blade should be used to visualize the larynx. Topical anesthetic may be helpful to decrease laryngospasm but should be used sparingly to prevent toxicity. Once the larynx is visualized a small diameter polypropylene catheter can be gently introduced into the trachea. Using it as a stylet, the ET is fed over the catheter, and slowly advanced into the trachea with slight rotation at which point the stylet can be removed. IV catheters can be used to intubate guinea pigs, hamsters, and other small rodents but is typically not carried out due to their small size. Additionally, viscous oropharyngeal secretions are much more likely to obstruct ET tubes in these species due to their narrow diameter.

Anesthetic depth monitoring using physiologic reflexes as in other mammals should be used. HR, RR and body temperature should be recorded. ECG is very helpful to monitor HR in these species as it is difficult to count accurately via auscultation. Instrumentation developed for research settings is available and allows noninvasive blood pressure and pulse oximetry monitoring via the tail in rodents. Capnography and pulse oximetry provide useful information to access ventilatory efficiency. Similar precautions, as with domestic animals, apply to small mammals during recovery. Particular emphasis should be placed on maintaining optimal body temperature, a clear airway and providing appropriate analgesia.

Conclusion

The same principles of anesthesia apply to exotic animals as they do in other animals. Clinicians need to assess risks and the potential for pain along with the needs of the needs of each patient. Dosages in this review may not be appropriate for all species or under all conditions. When working with unfamiliar species use the lowest dose and number of individuals to test anesthetic efficacy and safety. Familiarity of unique anatomy and physiology is vital to planning and carrying out anesthesia safely in nondomestic species. The field of exotic animal anesthesia is continually advancing and it is important to consult the newest references for the most current information.

References

1.  Brown, L. A. 1993. Anesthesia and restraint. In: Stoskopf, M. K. (ed.). Fish Medicine. W. B. Saunders Co., Philadelphia, Pennsylvania. Pp. 79–90.

2.  Gunkel, C., and M. Lafortune. 2005. Current techniques in avian medicine. Semin. Avian Exot. Pet. 14: 263–276.

3.  Harms, C. A. 2003. Fish. In: Fowler, M. E., and R. E. Miller (eds.). Zoo and Wild Animal Medicine. 5th ed. W. B. Saunders Co., Philadelphia, Pennsylvania. Pp. 7–9.

4.  Helmer, P., and D. Whiteside. 2005. Amphibian anatomy and physiology. In: O'Malley, B. (ed.). Clinical Anatomy and Physiology of Exotic Species: Structure and Function of Mammals, Birds, Reptiles and Amphibians. Elsevier Limited, London, UK. Pp. 3–14.

5.  Schumacher, J., and T. Yelen. 2006. Anesthesia and analgesia. In: Mader, D. R. (ed.). Reptile Medicine and Surgery. 2nd ed. Elsevier/Saunders, St. Louis, Missouri. Pp. 442–452.

6.  Sladky, K. and C. Mans. 2012. Clinical anesthesia in reptiles. J. Exotic Pet Med. 21: 17–31.

7.  Sneddon, L. U. 2012. Clinical anesthesia and analgesia in fish. J. Exotic Pet Med. 21: 32–43.

8.  Stetter, M. 2007. Amphibians. In: West, G., D. Heard, and N. Caulkett (eds.). Zoo Animal & Wildlife Immobilization and Anesthesia. Blackwell Publishing, Ames, Iowa. Pp. 205–209.

9.  Wenger, S. 2012. Anesthesia and analgesia in rabbits and rodents. J. Exotic Pet Med. 21: 7–16.

10. Wright, K. 2001. Restraint techniques and euthanasia. In: Wright, K. M., and B. R. Whitaker (eds.). Amphibian Medicine and Captive Husbandry. Krieger Publishing Co., Malabar, Florida. Pp. 111–121.