ANESTHESIA IN EXOTIC PETS

ANESTHESIA IN EXOTIC PETS

Geraldine Diethelm Mader, Vet. Med.

There are books written on the subject of anesthesia in all the different species. However, in private clinical practice, it is not necessary to know fifty ways to anesthetize a hamster. What is needed is a standard protocol for the average patient. A standardized protocol should be established for all the different patients that are treated by your hospital. Whether the anesthesia is performed by a veterinary technician or by the veterinarian, it is important that these protocols are followed. Once you get used to a particular drug or anesthetic protocol, you can learn what to expect, and will be better able to deal with problems should they arise.

In general you should not vary from these routines unless a specific situation dictates the need. The following protocols are those that the authors have used with great success in exotic practice.

GENERAL CONSIDERATIONS

Fasting

Because of their small size and rapid metabolisms, it is generally not recommended to submit small mammals to a prolonged fast prior to surgery. To minimize the risk of regurgitation and aspiration, it may be wise to withhold food for no more than three hours prior to the procedure. Guinea pigs will store food in their mouths, and it is not uncommon for them to spit this food up after they are induced. Careful swabbing of their oral cavities with cotton swabs will usually suffice.

Birds have relatively little stores of glycogen in their livers, and as a result, will experience rapid hypoglycemia if fasted for prolonged periods. The three hour limit is appropriate for avian patients as well.

Reptiles do not have a problem with regurgitation during anesthesia. Therefore, It is usually not necessary to fast these animals prior to surgery unless you are planning on a gastric procedure, in which case a fast and an enema may be essential.

Pre-operative

All of the general principles of anesthesia and pre-operative evaluation that apply to mammal patients also apply to exotic pets. A thorough pre-operative physical examination by the surgeon, appropriate laboratory screening, radiographs etc., should all be performed, or at least recommended to the client.

Patients should always be classified as to their risk of anesthesia (healthy pet, elective procedure vs. patient with major health problems), and the potential surgical risk (simple restraint for laboratory sample collection vs. prolonged procedure with potential for major blood loss).

Caution should be taken with patients having a low PCV, as these animal’s do not have a lot of blood to start with, so even minor hemorrhage can be lethal in a small pet. Consider pre-operative transfusions in those animals at risk, have blood ready for intra-operative transfusions, or at least pre-medicate patients with fluids prior to the procedures.

Since hypothermia is of paramount concern in these little patients, the fluids should always be prewarmed prior to administration. In addition, due to the generally higher metabolic rates in these little animals, it is wise to use a 5% dextrose solution IV, or a 2.5% dextrose in a balanced electrolyte solution either pre-operatively, intra-operatively, or immediately post-operatively.

Most of the exotic pets that we deal with have a relatively large surface area to volume ratio (in general, the smaller the pet, the larger the ratio). As mentioned, this larger surface area means that these patients are prone to hypothermia. Some form of supplemental heating is mandatory for these patients.

Circulating warm water blankets, water filled gloves (USE EXTREMEM CAUTION WITH THIS TECHNIQUE!), warmed anesthetic gasses (tube warmers), heated fluids, heated lavage fluids, and for surgical prep, the use of chlorhexidine rather than alcohol. The avian patients, with their unique respiratory system, are also prone to hypothermia. Since, in order for a bird to respire, gas needs to constantly flow across the parabronchi, there is considerable potential for heat loss. Again, many of the above principles for preventing hypothermia apply.

Reptiles, being ectothermic, can be totally unpredictable in their response to anesthesia. To better control this, it is best to eliminate temperature as a variable. Every reptile group has its own preferred optimal temperature. For instance, a garter snake thrives at temperatures around the high 70’s to the low 80’s, whereas the green iguana needs temperatures in the low 90’s to flourish. So, when performing anesthesia in these patients, where a garter snake may do just fine at 80 degrees, the iguana may be slower to induce, may be more difficult to maintain, and may take much longer to recover.

Induction and Anestesia

Injectable anesthetics are used more frequently in reptiles than in birds. However, the agents are used more for restraint than for anesthesia. Ketamine and Telazol (tiletamine/zolazepam) are the drugs of choice. Telazol needs to be reconstituted, and if not used within 14 days, goes bad. It has the advantage over ketamine in that the dose is much smaller, being especially useful in larger patients (large constrictors, tortoises).

Ketamine, due to its acidic pH, has a tendency to cause discoloration to the scales in reptiles when injected superficially. These marks usually result in permanent discoloration, so, it is prudent to advise your client prior to using it.

Reports from England suggest that Propofol makes an excellent pre-medicant in iguanids. At a dose of 1 ml/kg, IV in the tail vein provides 10 - 15 minutes of anesthesia, allowing for intubation or minor procedures. Keep in mind that Propofol does not produce analgesia. Recently, Bennet et. al recommends a dose of 5-10 mg/kg IO as an induction agent. The animals were then maintained at 0.5 mg/kg/min.

Supplemental oxygen via an endotracheal tube was recommended. As with the other exotics, for prolonged procedures, it is wise to maintain the reptile patient with a volatile anesthetic. Again, isoflurane is the agent of choice as it permits rapid induction and compared to the other agents, a more rapid recovery.

In most lizards mask induction is possible. Snakes and turtles can hold their breath for minutes to hours. If injectable anesthetic induction is not feasible, then placing these animals in an induction chamber, under supervision, is possible.

Reptiles are generally easy to intubate. In snakes and varanid lizards, the glottis is immediately inside of the mouth, making intubation an easy task, even in awake animals. In most lizards and the turtles, the glottis is caudal to the fleshy tongue. In these animals it is usually necessary to pre-midicate with Telazol or ketamine prior to intubation.

In reptiles, unlike in mammals, the glottis is always closed except during inspiration. This means that the anesthetist must be patient when performing the intubation. Do not force the tube into a closed glottis, rather wait until the patient takes a breath and then gently pass the tube into the trachea. Snakes and lizards have a «C» shaped tracheal cartilage, the dorsal surface being comprised of a membrane. Turtles and the crocodylia have complete tracheal rings. When intubating snakes and liazards, extreme care must be taken no to damage this delicate dorsal tracheal membrane. It is preferable to use shouldered «Cole» tubes rather than endotracheal tubes with a cuff. Reptiles lack a diaphragm. Their breathing is voluntary, not involuntary as it is in mammals. This means that when they are fully anesthetized, they become apneaic - or sill stop breathing. They must be ventilated in order to maintain steady state anesthesia.

Induction is accomplished by administering one breath every ten seconds. The pressure should not exceed 15 cm H2O (11 mm Hg). Once induced, maintenance can be accomplished with one breath every 30 seconds. Recovery can be achieved with no more than one breath every minute until spontaneous respiration returns.

The stimulus for reptiles to breath is a decreasing oxygen tension, as opposed to mammals, whose stimulus to breath is an increase in carbon dioxide. This is important for post-anesthetic recovery since it is imperative to discontinue the oxygen as soon as the procedure is finished. For recovery, continue to ventilate using only room air.

Post-operative

The most important thing to remember post-operatively is to maintain the patient’s warmth. The pet should be moved to a warmed recovery cage or incubator. In addition, fluid balance must be addressed, as dehydrated patients will have a more difficult time metabolizing or eliminating the anesthetic. Post-operative analgesia should always be a consideration. Few analgesia studies have been done in birds and reptiles, but substantial work has been done in small mammals due to their extensive use in laboratory research. Several analgesics are listed at the end of the proceedings.

PATIENT MONITORING

Over the years several monitoring devices have been employed to evaluate anesthetized patients. The electrocardiogram has been the gold standard. Although the significance of the tracings may not be fully understood for all the species, it usually provides reliable indication of the patient’s pulse rate. In addition, to the observant anesthetist, it may also provide clues toward impending changes in the patient’s condition. Many of the smaller patients, and most of the reptilian patients, are impossible to monitor with an E.C.G. due to either their rapid heart rates, or the small electrical potential produced.

Ultrasonic Dopplers detect pulsatile blood flow in the patient. It produces an audible signal for each pulse wave. It is considered very accurate, but has the disadvantage of not being able to provide any clues on the patient’s physiological changes. In addition, the noise that the Doppler produces is grating to listen to, and is usually best when attached to headphones.

The stethoscope, regular bell type or esophageal, is an excellent monitoring device providing that it is used. However, they are usually ineffective in reptiles.

Recent advances in anesthetic monitoring in human medicine have seen the introduction of the pulse oximeter to veterinary anesthesia. Specifically, the pulse oximeter has been used to monitor pulse and oxygenation during anesthesia and critical care settings. The pulse oximeter employs non-invasive technology and is user-friendly, requiring minimal training for proper and accurate usage by even lay staff.

These attributes make its use in anesthetic monitoring for exotic patients an attractive adjunct to the more conventional electrocardiogram or Doppler ultrasonic flow detectors.

The pulse oximeter is a spectrophotoelectric device that is applied typically via means of a clamp, preferable over a pulsating vascular bed to glabrous areas of the skin. The probe contains a diode that emits light in both the red and infra-red wavelengths. This dual-wavelength emitted light then passes through the pulsating vascular bed adjacent to the diode, and then is registered in a photodetector. The percent transmittance of the light is calculated within the unit and further equated to an oxygen saturation estimate.

Reptiles, unlike mammals which maintain respiratory drive based on many factors, including blood CO2 and pH, regulate their respiratory rate by careful balance of oxygen partial pressure (PO2) and temperature. At higher temperatures, the tissue demand for oxygen likewise increases. The increase demand is not met by an increase in respiratory rate, but rather an increase in tidal volume. Of particular clinical significance here is the ability to monitor the delivered oxygen to the tissues of anesthetized reptiles. It is not uncommon for reptiles post-procedure to remain anesthetized for prolonged periods. By monitoring SPO2 it may be possible to more accurately assess the state of the reptilian patient. In mammalian anesthesia the tendency is there to frequently bag the patient during recovery, whereas, in the reptilian patient, this increased oxygen may actually be inhibiting the return to spontaneous respiration.

The sensor traditionally has been incorporated into a finger or «clothespin-like» clamp positioned either over the patient’s finger (human) or tongue (veterinary patient). There are several new probes available for use with exotics (Surgivet). Aside from the standard tongue clips, there are now C-clamps, tail wraps and esophageal/rectal probes. Of all the species, the birds are probably the most difficult to monitor, however, it is usually possible, using one of the probes, to monitor most any patient. The esophageal probes seem to be the most effective in the avian patient. Reptilian patients have been difficult to monitor due to their thick skin and oftentimes heavy pigment, making placement of any kind of clamp-on type of sensor ineffective. The rectal or esophageal probe make the measurement of pulse rate and arterial oxygen saturation more feasible even in these patients.

Placement of the esophageal probe is best when the spectrophotometric diodes are positioned adjacent to the internal carotids/jugular vein complex, which is readily accessible through the oral cavity in an anesthetized patient. In patients where this is not logistically feasible (oral surgery), a membranous sheath can be applied over the probe so that it can be inserted rectally, with the diodes directed dorsally so that it can sense the caudal aorta and renal arteries. This is not as efficient in the avian patients. In these cases, using the C-clamp over the thigh has been effective.

CONCLUSION

Regardless of the instrumentation employed, there is no substitute for an alert, attentive anesthetist. Heart rate, respiratory rate, mucous membrane color, capillary refill time and reflexes are all reliable, useful parameters for monitoring the anesthetized patient. Don’t forget your medicine that you learned in dogs and cats. Don’t be afraid to apply the principles to your exotic patients.