Stephen J. Hernandez-Divers, BVetMed, DZooMed, MRCVS, DACZM
Zoological Medicine, Department of Small Animal Medicine & Surgery, College of Veterinary Medicine, University of Georgia, Athens, GA
Principles of Small Mammal Anesthesia
Being a good anesthesiologist is an indispensable skill for the exotic animal practitioner. Short anesthetic procedures are commonplace for completing tasks such as thorough examination, phlebotomy, radiography, or other short diagnostic/therapeutic procedures. In addition, many small mammal clinical problems necessitate surgery. Readers are directed to detailed reviews that are available on the subject, as there is only space for personal preferences to be presented here.1-4
Preparation
Maximize your success by: 1) preparing all the equipment ahead of time (anesthetic drugs, emergency drugs, catheters, fluids, fluid additives, non-rebreathing anesthetic machines with appropriately sized bags and masks, surgical equipment, etc. 2) making sure the patient has been stabilized prior to the anesthetic episode as much as possible, (warmed, rehydrated, in a good plane of nutrition, fasted if needed, no metabolic derangements 3) prepare yourself (understand anatomy/physiology of the patient, understand procedure, anticipate and prepare for problems).
There are some basic principles that apply when anesthetizing small mammalian patients in addition to the sound, basic principles of domestic animal anesthesiology. These additional comments are based on the fact that exotic mammals are often small in size, have high metabolic rates, have high body surface area: volume ratios and are prone to hypothermia, are often catecholamine driven prey animals that 'stress' easily, are typically presented in advanced stages of disease (often respiratory) with little respiratory or cardiovascular reserve, and have anatomy that challenges endotracheal intubation and intravenous access.
However, there are solutions that can help address these problems;
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Problem
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Potential Complication
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Solution
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Small size
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Mechanical obstruction of airway due to positioning; compression of thoracic cavity during handling/surgery;
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Being careful to keep head/neck extended; not placing heavy drapes, equipment or resting hands on the animal during procedures
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High metabolic rate
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Hypoglycemia, esp. if fasted prior to sx procedure; metabolize certain drugs faster; have higher fluid maintenance requirements; maybe affected by anesthetic drugs more severely
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Plan for higher fluid requirements and ALWAYS administer fluids pre-, peri- and post-operatively; consider dextrose in fluid solution; either do not fast, or fast for the minimal amt of time necessary; utilize drug dosages based on pharmacokinetic studies for that species; be ready to reverse drugs if effect is undesirable or be ready to support the animal fully through anesthetic period
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High surface area: volume ration
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Hypothermia; small animals lose heat more readily; during abdominal/thoracic sx the amount of heat loss is greatly increased; hypothermia can lead to decreased anesthetic requirements, prolonged recovery, bradycardia and terminal fibrillation
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Provide warmth from the point of induction; monitor body temperature carefully and frequently; plan for different types of supplemental heat during anesthetic episode and use them simultaneously (radiant source, heating pad, warm fluids, warm air); administer heat prior to animal becoming hypothermic and continue until the animal can thermoregulate
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Catecholamine driven prey animals
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Stressed prey species release endogenous catecholamines that sensitize the myocardium (among other effects); this effect is much more pronounced in certain species (esp. rabbits)
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Minimize handling pre-op; plan for hospital environment to be stress-free (shelter boxes, quiet wards away from predators); pre-medicate patient with anxiolytics prior to induction; utilize strategies such as rapid induction agents and induction chambers (instead of face-mask) that minimize handling
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Present with underlying disease
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Little respiratory or cardiovascular reserve
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Perform a pre-anesthetic evaluation; minimize anesthetic time as much as possible; carefully monitor patient; plan to ventilate animal; plan to assist blood pressure with fluids and other drugs (respiratory stimulants, positive inotropes, chronotropes, vasoconstrictors etc).
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Difficult to intubate
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No control over ventilation; potential for hypoxemia
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Study anatomy of the animal prior to scheduling procedure and research intubation methods; practice on cadavers; many small mammals can be safely intubated with practice and proper equipment; if unable to intubate, realize limitations of lack of control of airway, keep procedure short; utilize well-fitting mask and appropriate lab animal anesthetic circuit
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Difficult to gain vascular access
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Unable to administer maintenance fluids during anesthetic procedure to maintain proper blood pressure and blood flow to vital organs; unable to administer emergency agents if needed
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Study anatomy of the animal prior to scheduling procedure and research catheterization methods; many small mammals can be catheterized, either IV or IO with practice and proper equipment; if unable, administer maintenance fluid requirements SQ before the procedure (or at induction), and repeat just prior to recovery
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Referred Sedative/Anesthetic/Analgesic Protocols
There are many different protocols available and only those routinely used and preferred by the author are presented here. Most of the animals seen by the author are clinically ill and/or aged, and therefore potent alpha-2 agonists like medetomidine are usually avoided.
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Species
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Premedication
(or sedation)
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Induction
(typically 10-15 min after premedication)
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Maintenance
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Additional Analgesics
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Ferret
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Butorphanol 0.1-0.4 mg/kg SQ/IM q 4 hrs
Midazolam 0.25-0.5 mg/kg SQ/IM q 4-8 hrs
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Isoflurane or sevoflurane by mask
Ketamine 5-10 mg/kg IV or 10-15 mg/kg IM
Propofol 3-6 mg/kg IV
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Isoflurane 1-2.5%
Sevoflurane 2-4%
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Meloxicam 0.2 mg/kg SQ/IM/PO q 12-24 hrs
Oxymorphone 0.05-0.2 mg/kg SQ/IM, q 8-12 hrs
Buprenorphine, 0.01-0.03 mg/kg SQ/IM/IV, q 6-12 hrs
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Rabbit
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Butorphanol 0.1-0.5 mg/kg SQ/IM, q 2-3 hrs
Midazolam 0.5-1 mg/kg SQ/IM q4-8 hrs
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Ketamine 15 mg/kg IV or 25-30 mg/kg IM
Gas induction provides little time for intubation. In addition apnea often occurs with isoflurane, less so with sevoflurane.
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Isoflurane 1.5-2.5%
Sevoflurane 2-4%
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Meloxicam 0.5-1.0 mg/kg SQ/IM/PO q 12 hrs
Oxymorphone 0.05-0.2 mg/kg SQ/IM, q 8-12 hrs
Buprenorphine, 0.01-0.05 mg/kg SQ/IM/IV, q 6-12 hrs
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Rat, Mouse,
Gerbil,Hamster,
Guinea Pig,
Chinchilla
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Butorphanol 2-4 mg/kg SQ/IM/IP, q 2-3 hrs
Midazolam 1-2 mg/kg SQ/IM/IP q4-6 hrs
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Ketamine 25-50 mg/kg IM/IP if intubating
Isoflurane (5%) or sevoflurane (8%) with induction chamber if to be maintained by mask
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Isoflurane 1.5-3%
Sevoflurane 2-5%
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Meloxicam 1-2 mg/kg SQ/PO q 12 hrs
Oxymorphone 0.2-0.5 mg/kg SQ q 8-12 hrs
Buprenorphine, 0.05-0.1 mg/kg SQ q 6-12 hrs
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Ferrets
The anatomy and physiology of the ferrets is much like of other small carnivores including the domestic dog and cat, and therefore can be approached in a very similar manner. Intubation is seldom a problem; however intravenous catheter placement may have to follow induction, rather than precede it unless the animal is heavily sedated.
Rabbits
These are easily-stressed patients which often present with underlying respiratory or gastro-intestinal disease. The focus should be to induce them as smoothly and quietly as possible (pre-medication is a must). High levels of circulating catecholamines, combined with the stress of handling/restraint, hypoxemia, hypercarbia and unpredictable responses to anesthetic agents can lead to respiratory and cardiac arrest. They are challenging to intubate (several techniques facilitate this procedure: blind, endoscope-assisted) but at the same time, are one of the patients that often need assisted ventilation the most. The chances of successful intubation can be maximized by:
1. Sedation/premedication prior to induction using ketamine. This will provide 5-10 minutes of restraint for intubation (compared to 10-20 seconds following gas induction).
2. Maintain the rabbit on oxygen by holding a small face mask over the nose.
3. Use gauze to open the mouth fully and hyperextend the head and neck. Insert the laryngoscope (Wisconsin 00, long blade) or endoscope to visualize the epiglottis and ensure that it is not engaged above the soft palate (obligate nasal breather). Applying mild dorsal pressure on the soft palate will cause the epiglottis to fall ventrally and expose the glottis. Apply lidocaine to the glottis, and wait for 1 minute, while maintaining on nasal oxygen.
4. Slide the endotracheal tube over a 1-2.7 mm endoscope (or endoscopic laryngoscope) and insert the endoscope into the glottis before sliding the tube into the trachea. Alternatively use a Wisconsin 00 long bladed laryngoscope to visualize the insertion of the glottis. Many experienced lab animal vets are able to perform a blind intubation technique; however given the smaller size of most pet rabbits compared to new Zealand whites, varied breed/conformation and disease status, direct visualization is recommended.
5. If two attempts at intubation are unsuccessful, the animal should be placed on a tight-fitting face-mask. Pharyngeal edema and consequent airway obstruction from more than 2-3 intubation attempts is common. If suspected, fast-acting, powerful anti-inflammatories should be administered IV (steroids, potent NSAIDs).
Guinea Pigs and Chinchillas
These patients are also easy to stress, and difficult to intubate. Due to the presence of the palatal ostium (a ring of tissue between the oropharynx and pharynx), intubation is more difficult and virtually impossible without the aid of an endoscope.
Mice and Rats
Large rats can be intubated as outlined above, but most small rodents are maintained using a dedicated rodent anesthesia circuit mask (http://www.harvardapparatus.com). Subclinical respiratory disease is very common (e.g. Mycoplasma, cilia-associated respiratory [CAR] Bacillus). Preventing hypoglycemia can be difficult.
Ventilatory Support
The author prefers to connect all intubated small mammals < 8kg to a pressure-cycle ventilator (Small Animal Ventilator, Ventronics, http://www.vetronics.net/products/smvent.php). This ventilator is relatively inexpensive, simple to use, and has the advantage that the animal can be maintained on a non-rebreathing circuit, and ventilated when necessary simply by flicking a switch. Depth and frequency of ventilation are initially set to mimic pre-anesthetic respiration, but are modified to maintain end tidal capnography between 35-45 mmHg.
Monitoring
The primary goal of anesthetic monitoring should be to prevent, identify, and treat hypotension, bradycardia, arrhythmias, hypoxemia and hypercapnia. Although the size and anatomy of some small mammal patients often preclude some anesthetic monitoring techniques, this should not discourage the practitioner from:
1. Performing a basic pre-anesthetic examination to record heart and respiratory rates and character, and if possible collect baseline clinicopathologic data. Complete blood counts and full biochemistry profiles are seldom possible (and indeed anesthesia is often used to facilitate sample collection), however PCV, TP, BUN, glucose can be run patient-side with small samples. Alternatively, blood samples may be collected and run immediately following induction.
2. Recording the precise time of premedication/induction; this step is often overlooked; it is important to know how long an animal has been anesthetized, and to maximize efforts in expediting procedures.
3. Gauge the depth of anesthesia by observation of mentation and reflexes.
4. Utilize simple monitoring techniques first rather than spending 15 minutes trying to get the pulse oximeter probe to work! First take a heart rate and note cardiac rhythm with a simple stethoscope, note the depth and frequency of ventilation and judge based upon pre-anesthetic values.
5. Utilize equipment that will give you the most information for the 'least hassle'; Dopplers, end-tidal capnographs and ECGs are often easier to use. Pulse oximeters can be temperamental but are still useful if a reliable pulse wave is obtained (irregular or poor pulse wave leads to untrustworthy readings). SpO2 < 90% equates to saO2 < 60 mmHg and hypoxia. Indirect blood pressure readings can often be taken using a sphygmomanometer with the Doppler. Do not rely totally on the monitoring equipment - always rely on human evaluation of the patient which means that there must be an anesthesiologist/anesthetist dedicated to the case at all times.
Recovery and Post-Operative Care
Recovery and the immediate post-operative period can be just as critical in ensuring that your small mammal patient recovers completely:
1. Once extubated maintain on oxygen until fully conscious.
2. Place the animal in an incubator and recover in a normal position (preferably sternal). Recovery areas should be quiet, warm and in a place where the animal can be readily monitored. Do not add water/food bowls or cage furniture/substrate until the animal is fully ambulatory.
3. Monitor the animal frequently and administer further fluids or antagonists as needed.
4. Postoperative analgesia is vital and should be a continuation/modification of the pre-emptive analgesic protocol employed for premedication.
References
1. Carpenter, J.W. 2005. Exotic Animal Formulary. WB Saunders Co., St Louis.
2. Heard, D.J. 2004. Anesthesia, analgesia, and sedation of small mammals. In. K.E. Quesenberry and J.W. Carpenter, (eds.). Ferrets, Rabbits and Rodents Clinical Medicine and Surgery, Second edition ed. WB Saunders Co, Philadelphia, PA. Pp. 356-369.
3. Heard, D.J. 2007. Lagomorphs (rabbits, hares and pikas). In. G. West, D. Heard, and N. Caulkett, (eds.). Zoo animal & wildlife immobilization and anesthesia, ed. Blackwell Publishing, Ames, Iowa. Pp. 647-653.
4. Heard, D.J. 2007. Rodents. In. G. West, D. Heard, and N. Caulkett, (eds.). Zoo animal & wildlife immobilization and anesthesia, ed. Blackwell Publishing, Ames, Iowa. Pp. 655-663.