Exotic Small Mammal SEDATION AND ANESTHESIA

Lorelei D’Avolio, LVT, VTS (Clinical Practice-Exotics), CVPM

Cape Cod Veterinary Specialists

Introduction

Technicians who are confident in anesthesia protocols for dogs and cats can use much of their knowledge for exotic small mammal anesthesia; however, there are some very important differences that they must learn first. Exotic small mammals have some physiological, anatomical, and behavioral differences that make anesthesia much more challenging than in other mammals. But once these unique traits are understood, anesthesia can be a safe event for even the smallest exotic pet mammal. This manuscript and lecture will describe anesthetic techniques for rabbits, guinea pigs, small rodents, and ferrets.

PRE-ANESTHETIC PREPARATION

Exotic small mammals that are prey species — such as rabbits, guinea pigs, chinchillas, and small rodents, — are much more susceptible to stress-related complications than dogs and cats. While ferrets may tolerate a lot of noise, commotion, and activity, prey species will do better in a quiet, dimly lit environment away from loud noises. Technicians should check vital signs before any medications are administered. This includes checking temperature, pulse, respiration rate (RR), and body weight. Because handling can artificially elevate some of these parameters, obtaining the RR through observing in the enclosure is recommended. Depending on the patient, diagnostic lab tests should be acquired prior to anesthesia to correct any abnormalities or adjust anesthesia. Rabbits and rodents are prone to underlying respiratory problems, and thoracic radiographs are helpful at understanding anesthetic risk. Ferrets are prone to hypoglycemia and may require a glucose CRI. Guinea pigs often have underlying reproductive or urinary tract issues and may be anemic.

Exotic small mammals, other than ferrets, should not be fasted. Rabbits and small rodents have an anatomical valve restricting food from passing from the stomach into the esophagus, so they cannot vomit. For herbivores, like rabbits, guinea pigs, and chinchillas, it is important to have food in their stomach, cecum, and intestines to prevent hypoglycemia, dehydration, and ileus. For omnivorous small rodents, like rats, mice, gerbils, and hamsters, fasting is exceptionally dangerous and can contribute to hypoglycemia under anesthesia. Removing food for an hour prior to anesthesia is acceptable and will help ensure that the oral cavity is clear, but fasting for longer periods should be avoided. Ferrets are also susceptible to hypoglycemia, and they have a very short gastrointestinal transit time of three to four hours. But because they can vomit, they should be fasted, but for no longer than three to four hours.

Intravenous or intraosseous catheters should be used whenever possible. Rabbits, ferrets, guinea pigs, and chinchillas can have 22, 24, or 26g IV catheters easily placed in the cephalic veins. Guinea pigs and ferrets can have very tough skin and may require a small nick in the skin to facilitate threading the catheter. Guinea pig cephalic veins are very lateral in comparison to other species. If IV access is not possible, IO should be considered. These are relatively easy to place using a spinal needle, IV catheter, or regular hypodermic needle. Most guinea pigs or chinchillas require a 20 to 22-g needle while a gerbil or hamster may require 25 to 27g. The length of the needle should be long enough to extend 1/3 to 1/2 the length of the bone. A sterile piece of cerclage wire or stainless-steel suture may be needed to clear out boney material if a stylette is not used. IO catheter placement is considered painful and appropriate local analgesia or anesthesia/sedation is mandatory. Placement also requires proper aseptic technique and should be sutured in place for stability. When placed properly, risks of complications are rare but could include infection or emboli of fat or bone marrow. The sites for placement in rodents include the proximal humerus, proximal tibia, and proximal femur. IO catheters should not remain in place for more than 72 hours.

If IV or IO fluids are not possible, warm fluids should be administered prior to any surgical procedure and, prior to or immediately after induction subcutaneously. Fluids should be warmed to prevent hypothermia and can easily be given in the subscapular area. Veterinary technicians will need to draw predetermined doses into syringes rather than using a hanging bag because doses are sometimes only a few milliliters.

Pre Medication And Induction

Pre-anesthetic sedatives and anxiolytic medication is recommended for rabbits, guinea pigs, and chinchillas as these species release systemic catecholamines and cortisol under stress, which can result in anesthetic complications. For these species, sedation or tranquilization is often required to allow IV catheterization and endotracheal intubation. There are some controversial opinions about whether to use premedication with small rodents or not because of their small size. In the event one of the potential adverse side effects of a premedication occurs, one must be able to detect and manage the problem. For example, if a gerbil is given acepromazine and ketamine and begins to seize, there is little a veterinary technician can do without having IV/IO access. Conversely, the benefits of pre medicating animals that are so prone to stress-related side effects can be profound. Decisions should be made with the doctors on an individual basis based on the ability of the staff to be able to monitor and treat potential premedication side effects. Each patient should be evaluated to determine what the appropriate premedication may include rather than a standard “cocktail” for all animals. One attribute unique to rabbits is that many of them produce atropinase, an enzyme that renders atropine useless, and other anticholinergics less effective. If a rabbit requires an anticholinergic, it is recommended to try glycopyrrolate.

The following drug options for premedication, sedation, and often secondary induction can be species and dose-dependent:

• Teaming a full mu opioid (such as hydromorphone or fentanyl) with a benzodiazepine (such as midazolam) can provide effective and safe analgesia and sedation in most healthy or ill patients. These combinations can be given IM or SQ; however, diazepam can have erratic absorption and is painful when given via the SQ or IM route. Therefore, choosing a more water-soluble benzodiazepine such as midazolam is preferred.

• Alpha-2 adrenergic agonists such as dexmedetomidine are used in exotic animal species; however, just as in small animals, they have profound cardiovascular side effects and are not appropriate for all patients. They are best used in young and/or healthy rabbits.

• Mixtures such as dexmedetomidine and ketamine are commonly used in combination with midazolam to pre medicate rabbits. Ketamine should be used with caution in animals with cardiac or renal issues. Ketamine should not be used alone as convulsions can occur.

• Acepromazine can be combined with an opioid; however, it is used less frequently than in dogs and cats due to concern over its side effects and reliability.

• The neuroactive steroid alflaxalone has a growing body of literature on its use in small mammals, and dosing for exotic small mammals is currently being extrapolated from cats and dogs. Reports on alflaxalone use in rabbits indicate minimal negative cardiovascular effects and rapid recoveries; however, there is little scientific research on these species.

• Propofol can be used as an induction agent for patients with an IV catheter placed. It is not recommended to use this if the patient is not going to be intubated due to the possibility of apnea with this drug.

These drug options can be used in combinations and at varying doses. Often, they will create a deep enough plane of sedation to allow for IV catheter placement and/or intubation. Once sedated, they can be put on Isoflurane or Sevoflurane for the transition from induction to maintenance anesthesia.

If patients are not sedated enough from premedications to be intubated or maintained on a mask without induction, they will require an induction agent. Using a mask with oxygen and Isoflurane or Sevoflurane is historically considered a very safe induction method as opposed to adding more injectable drugs. However, technicians must be diligent in preventing additional stress to these patients. If it is determined that a patient is going to have a mask or box induction, it is important that technicians advocate for the patient and as part of the anesthetic team and help determine if other anxiolytics, sedatives, or analgesics can be used. Directly placing a mask over a patient’s head or placing the entire patient in an induction chamber without any premedication should be avoided.

Intubation

Some exotic small mammals can be easily intubated, such as the ferret. The procedure is like that of intubating a cat, but with a smaller diameter ET tube. Rabbits and rodents have very small chest cavities in comparison to their large abdominal cavities. When in dorsal recumbency, the viscera can put added pressure on the thoracic cavity. They also may have undiagnosed respiratory illnesses. Combining these factors with the use of drugs that may suppress the respiratory system creates a potential respiratory disaster if these patients are not intubated. Unfortunately, intubating them has more challenges than intubating ferrets.

Because rabbits primarily breathe through their nose, many practitioners do not routinely intubate them and instead use a mask covering the nose to deliver inhalant anesthetic agents. If there are no anesthetic complications, this technique will suffice. However, in the event of respiratory arrest, there is very little one can do to ventilate the patient. It is possible that by pushing oxygen through a tight-fitting mask over the nose, the lungs may become inflated; however, it is also possible that the stomach will become dilated with oxygen rather than the lungs. The oral cavity of rabbits is very long and narrow with a large fleshy tongue and sharp angular teeth. These features make it almost impossible to visualize the glottis. Because of their anatomy, endotracheal intubation in rabbits is commonly done blindly and requires the animal to be very sedated or anesthetized. This procedure generally requires at least two people.

• While administering passive supplemental oxygen, hold the head up to straighten the trachea as much as possible.

• Apply dilute local anesthesia to the supraglottic region to reduce laryngeal spasm.

• Insert an appropriate-sized endotracheal tube.

• Listen for the loudest and clearest breath sounds through the endotracheal tube. While listening, attempt to feed the tube into the presumed trachea.

• Hair plucked from the patient can be used to detect whether the endotracheal tube is in the trachea.

• Visualization of condensation created by breath on the walls of a clear endotracheal tube is also a confirmation of proper placement.

• Using rigid or flexible endoscopes is also an option for practices with this equipment. These techniques often require a third person to drive the camera and can take longer than an experienced technician with excellent blind intubation skills.

A commercially available modified laryngeal mask that covers the supraglottic region (V-gel™ [Docsinnovent, Inc]) has been developed for rabbits and felines. Although endotracheal intubation is preferred, V-gel offers an easier approach to the airway when endotracheal intubation proves too difficult or time-consuming. V-gel is not ideal for animals requiring oral surgery as it takes up a significant amount of space in the mouth, and there is potential for fluid leakage around the endotracheal tube if the patient is undergoing a dental procedure. Providing positive pressure ventilation is also not considered a guarantee with the V-gel, as the potential for forced air slipping around the seal is present.

Another use of this device is to run a guide tube through the V-gel into the trachea, remove the V-gel, and then slide an endotracheal tube over the guide tube for intubation.

Guinea pigs are one of the most difficult to intubate. Their anatomy is unique with the presence of a palatal ostium. This is a tiny opening in the caudal portion of the oral pharynx which is made of tissue that is easily damaged by the prodding of an endotracheal tube and can bleed profusely, even causing asphyxiation. Additionally, the oral cavity is long and narrow with prominent cheek teeth. One way to potentially intubate a guinea pig is by using a tiny laryngoscope blade such as a size 0 Miller to provide visualization of the palatal ostium. If an endotracheal tube is passed through the opening, blind intubation of the trachea may be possible. Alternatively, using an appropriately sized ear cone can be used in a similar fashion. In hospitals where small-sized endoscopes are utilized, they can also be used to assist in visualization. If guinea pigs are not intubated, technicians need to continually swab out their oral cavity to prevent aspiration of mucosal secretions or food/feces they may be holding in their oral cavity.

The smallest patients most veterinary hospitals have the ability and equipment to intubate are chinchillas and rats. These rodents can be intubated in a similar fashion to the rabbit, in that the technique is mostly blind. Proper positioning with the ventral neck flexed is essential. One technique that may help visualize the glottis is the use of a syringe tube with the plunger taken out and cut at a 25-degree angle. Once visualization is made, a tiny tube or even an IV catheter can be used to establish a patent airway.

Maintenance And Monitoring

For shorter procedures, injectable anesthetic premedication and induction agents may produce anesthesia of sufficient duration, especially when utilizing proper multimodal analgesia. For longer procedures, delivery of inhalant anesthetic agents such as isoflurane and sevoflurane via a non-rebreathing system is best. Exotic small mammals generally require higher levels of gas anesthesia than dogs and cats. Pre medications such as ketamine and dexmedetomidine will decrease inhalant anesthetic agent requirements while more painful procedures may increase the need for higher dosages. The key is for the veterinary technician to constantly be monitoring the patient and adjusting as needed. These small patients should have their chest and head slightly elevated above their abdominal cavity to allow the chest to expand easier.

Veterinary technicians should be comfortable assessing anesthetic depth by observing and recording reflexes. In rabbits, loss of pedal withdrawal, auricular (ear pinch), and palpebral reflexes indicate appropriate depth, while loss of corneal reflex and dilated pupils can be a sign of deep anesthesia and cerebral hypoxia. Mucous membrane color, capillary refill time, and temperature of extremities can be used to gauge cardiovascular function and tissue perfusion. Another valuable and easy monitoring technique is to use an ultrasonic Doppler to monitor the heart rate and rhythm. Placing the probe directly over the heart works well. Some small rodents, such as hamsters and mice, are at risk of eye proptosis and technicians should monitor this using lubrication and by applying pressure against the eyes.

Conventional monitoring equipment can be used for exotic small mammals; however, pediatric settings that can read a heart rate up to at least 350 bpm should be selected. An SPO2 monitor that reads higher heart rates will work well on ears or thin extremities. They do not work well on rabbit feet as rabbits have very thick fur and no hairless pads. Alligator clamps on ECG leads can cause pressure necrosis or tear thin skin. Using needles, specially manufactured flattened clips, or modified alligator clips is important. Mainstream capnography can be useful in monitoring trends in intubated animals. The approximate normal range for end-tidal carbon dioxide (EtCO2) is between 35 to 45mmHg.

Maintaining normothermia is important and warming devices such as forced air warming blankets, water circulating pads, or other convective heat sources that provide heat while preventing burns are ideal. Other techniques include using aluminum foil or bubble wrap around the extremities, delivering heated fluids, and radiant heat sources. Pediatric rectal thermometers work well to monitor body temperature.

Blood pressure monitoring is a great tool to assess cardiac function under anesthesia. Due to their small size, placing direct arterial lines to invasively monitor blood pressure is very challenging. However, using a cuff and sphygmomanometer of a Doppler to non-invasively monitor peripheral blood pressure in some larger species is easy and should be part of any long anesthetic procedure. An appropriately sized cuff placed on an appendage or tail works well. Shaving small patches of hair will help obtain better results. As with many other devices, veterinary technicians should pay attention to trends rather than exact numbers, recognizing that mean arterial pressure should remain over 60 mmHg and systolic blood pressure (SaP) over 90 mmHg.

Recovery

Post-operative oxygen is recommended until the pet is conscious. Once jaw tone returns, the patient can be extubated. Technicians should examine the glottis to make sure it is clear of debris or secretions that could be aspirated or obstruct the airway. Patients should be placed in sternal recumbency and housed in a warm, quiet, dimly lit room away from predators to minimize stress. Rodents will chew at anything that is irritating to them, including uncomfortable IV/IO catheters and incisions. But unfortunately, exotic small mammals do not tolerate E-collars well and become highly stressed, sometimes refusing food. Others will adeptly wriggle out of even the most secure E-collar. Rather than taking these risks, careful monitoring of patients without an E-collar to observe inclination to chew at incision sites or catheters is critical. Additional analgesics should be administered or added, such as NSAIDs or repeated doses of short-acting opioids, that may have been eliminated from the body. Depending on the surgical procedure, these animals can and should be offered food immediately after awakening to prevent/correct potential hypoglycemia as well as distract the animal from chewing or barbering its incision. Sometimes bandages need to be applied to prevent chewing sutures or staples. Temperatures should be monitored, and patients should be kept in a heated enclosure until normothermia has returned.

References And Suggested Reading

• Vella D, Donnelly T: Rabbits: Basic Anatomy, Physiology, and Husbandry. Quesenberry K, Carpenter J (eds): Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 3, St. Louis, 2012, Elsevier/Saunders, p 162

• Lennox A, Bauck L: Small Rodents: Basic Anatomy, Physiology, Husbandry, and Clinical Techniques. Quesenberry K, Carpenter J (eds): Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 3, St. Louis, 2012, Elsevier/Saunders, p 361

• Lennox A: Rabbits: Respiratory Disease and Pasteurellosis. Quesenberry K, Carpenter J (eds): Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 3, St. Louis, 2012, Elsevier/Saunders, p 207

• Hawkins M, Pascoe P: General Topics: Anesthesia, Analgesia, and Sedation of Small Mammals. Quesenberry K, Carpenter J (eds): Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 3, St. Louis, 2012, Elsevier/Saunders, p 429

• Morrisey J, Carpenter J: Formulary. Quesenberry K, Carpenter J (eds): Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 3, St. Louis, 2012, Elsevier/ Saunders, p 571-573

• Longley L: Anaesthesia of Exotic Pets, ed1, St. Louis, 2008, Elesvier, p 27-28, 36-38, 60-62, 96-99

LORELEI D’AVOLIO LVT, VTS (Clinical PracticeExotics), CVPM

Lorelei D'Avolio is the practice administrator of New York City's only stand-alone exotic pet practice, the Center for Avian and Exotic Medicine. She received her BA in journalism from Boston University, and after earning a second bachelor's degree in Veterinary Technology from Mercy College in 2001, she has been working exclusively with exotic pets. Lorelei has lectured at many national and international veterinary symposiums and is an educator at a veterinary assistant program in Cape Cod. She also has authored and edited textbook chapters, journal articles, and professional publications. Lorelei was a founding member and is the past president of the Academy of Veterinary Technicians in Clinical Practice (AVTCP). She is also a Certified Veterinary Practice Manager (CVPM)and in 2016, she was awarded the Veterinary Technician of the Year award from the New York State Association of Veterinary Technicians.