Rabbit and Rodent Anesthesia
American Association of Zoo Veterinarians Conference 2012
Stephen J. Divers, BVetMed, DZooMed, DECZM(herp), DACZM, FRCVS
Department of Small Animal Medicine & Surgery, College of Veterinary Medicine, University of Georgia, Athens, GA


With over 10 million pet rabbits (Oryctolagus cuniculus), ferrets (Mustela putorius furo), and rodents (order Rodentia) in the US, these exotic mammals represent the third largest group of companion mammals (behind dogs and cats).1 This group represents an expanding component of small animal practice, with many clients expecting the same level of medicine for them as for more traditional pet species. Many diagnostic and surgical procedures warrant anesthesia; therefore acquiring skills for competent anesthesia 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 personal preferences will largely be presented here.2-5 Practitioners are also finding that sedation, especially when combined with local anesthesia, is often a viable option to general anesthesia for such procedures as sample collection and catheterization.


Maximize success of any sedation/anesthesia procedure by:

1.  Preparing the veterinary team by reviewing the species-specific anatomy/physiology of the patient, the procedure(s) to be performed, anticipating potential problems, and having back-up plans.

2.  Calculate emergency drugs, or consider a dedicated EXCEL emergency drug chart (Table 1). For critical cases prepare individual emergency drug doses before induction.

3.  Preparing all equipment and supplies ahead of time (anesthetic drugs, catheters, fluids, fluid additives, non-rebreathing anesthetic machines with appropriately sized bags and masks, surgical equipment, etc.)

4.  Ensuring the patient has been stabilized as much as possible prior to sedation or anesthesia (e.g., patient is normothermic, rehydrated, previous good plane of nutrition, fasted if needed, no metabolic derangements).

There are some basic principles that apply when using general anesthesia in 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. It should be kept in mind that patients under sedation alone must be monitored carefully as well.

Table 1. Rabbit emergency drug calculator chart


Patient ID: 319902

Weight: 1.230 kg








0.2 mg/ml

0.02 mg/kg




Epinephrine (1:1000)

1 mg/ml

0.2 mg/kg





5 mg/ml

2 mg/kg





50 mg/ml

2 mg/kg




Shock fluids


90 ml/kg/hour




Maintenance fluids


70 ml/kg/day





0.4 mg/ml

0.01 mg/kg




Preferred Anesthetic Protocols

There are many different protocols available and only those routinely used and preferred by the author are presented here (Table 2). A more complete listing can be found in the references.1-4 Many animals presented are clinically ill and/or aged, and therefore potent alpha-2 agonists like dexmedetomidine are usually avoided, unless dealing with young healthy individuals.

Table 2. Premedication, induction and maintenance drugs preferred for rabbits


(typically 10–15 min after premedication)


Additional analgesics

Butorphanola 0.2–0.4 mg/kg SQ/IM
Buprenorphineb 0.01–0.05 mg/kg SQ/IM/IV
Oxymorphonec 0.05–0.2 mg/kg SQ/IM
Plus Midazolamd 0.5 mg/kg SQ/IM

Ketamineg 5–10 mg/kg plus additional Midazolamd 0.25–0.50 mg/kg IV
or, if IV inaccessible
Ketamineg 25–30 mg/kg IM
Gas induction using sevofluranef

Isofluranee 1.5–2.5%
Sevofluranef 2–4%

Meloxicami 0.5 mg/kg SQ/IM/PO q 12 hrs
Buprenorphineb 0.01–0.05 mg/kg SQ/IM/IV, q 6–12hrs
Oxymorphonec 0.05–0.2 mg/kg SQ/IM, q 8–12 hrs

aTorbugesic, 10 mg/ml, Fort Dodge Animal Health, Fort Dodge, IA, USA
bBuprenex, 0.3 mg/ml, Reckitt Benckiser Pharmaceuticals Inc., Richmond, VA, USA
cNumorphan, 1 mg/ml, Endo Pharmaceuticals Inc, Chadds Ford, PA, USA
dMidazolam, 5 mg/ml, Baxter Healthcare Corp, Deerfield, IL, USA
eIsoflo, Abbot Animal Health, Abbott Park, IL, USA
fSevoflo, Abbot Animal Health, Abbott Park, IL, USA
gKetaset, 100 mg/ml, Fort Dodge Animal Health, Fort Dodge, IA, USA
iMetacam, 5 mg/ml, Boehringer Ingelheim VetMedica Inc, St Joseph, MO, USA

Rabbits are easily stressed patients, which often present with underlying respiratory or gastrointestinal disease. The focus should be to induce them as smoothly and quietly as possible. Preemptive analgesia and sedation are critically important. 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. While large laboratory rabbits like the 4–5 kg-New Zealand White can be readily intubated using blind or traditional techniques, the more common, smaller pet rabbits can be challenging. In addition, it is often necessary to use a smaller endotracheal tube for blind intubation, which can seriously increase resistance and reduce gas flow (Table 3). For example a 1.5-kg rabbit would likely be intubated using a 2-mm endotracheal tube blind, or a 2.5-mm tube if using direct visualization. The 2.5-mm tube offers a relative reduction in air resistance from 26 to 11, a 58% improvement in air flow! These air flow resistances become increasingly pronounced when using small tubes for small animals.

Table 3. Endotracheal tube dimensions and relative resistance

Endotracheal tube

Cuff available

Relative resistance
(R = 81n/πr4)

21-gauge catheter



1.0 mm



1.5 mm



2.0 mm



2.5 mm



3.0 mm



4.0 mm



5.0 mm



The chances of successful intubation in small rabbits and rodents can be maximized by:

1.  Ensuring adequate sedation using an opioid and midazolam as premedicants prior to induction (see Table 2). Ketamine and midazolam induction will provide 5+ minutes of good restraint and relaxation for intubation (compared to 10–20 seconds following gas induction).

2.  Maintain the animal on oxygen by holding a small face mask over the nose during induction.

Laryngoscope/Endoscope-Guided Intubation

1.  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 (rabbits and many rodents are obligate nasal breathers). 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 nasal oxygen.

2.  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 through the glottis. Many experienced lab animal vets are able to perform a blind intubation technique in rabbits; however, given the smaller size of most pet rabbits compared to the 4–5-kg laboratory New Zealand white, varied breed/conformation and disease status, direct visualization is generally preferred.

Blind Intubation

Introduce an endotracheal tube into the oral cavity and over the tongue white listening for the sound of air movement as the tube advances. The tube is positioned over the glottis when the sound of air movement is the loudest. (Alternatively, watch for the presence of condensation in the tube). The tube is gently rotated while moving forward and back until the tube slides into the trachea. At no time is any pressure used to place the tube.

If two attempts at intubation are unsuccessful, the animal should be placed on a tight-fitting face mask. Pharyngeal edema and consequent dyspnea associated with repeated intubation attempts is common. If required, antiinflammatory drugs should be administered by the intravenous or intratracheal routes (e.g., NSAIDs or, less favorably, steroids).

Ventilation & Monitoring

The author prefers to connect all intubated small mammals < 8 kg to a ventilator. The ventilator is routinely set to the non-rebreather passive circuit, with active ventilation selected if required. This has the advantage of being able to select ventilation without changing circuits. Ventilation is used whenever ETCO2 rises above 45 mm Hg or SpO2 falls below 95%. Ventilation is used routinely for laparoscopy due to abdominal insufflation using CO2. Lennox prefers to use the ventilator for selected patients. An inexpensive pressure-cycle ventilator (Small Animal Ventilator, BASi Vetronics, Bioanalytical Systems Inc, West Lafayette, IN, USA) is available and has a proven track record in exotic mammal practice. This ventilator is simple to use, and has the advantage of being able to switch from non-rebreathing to ventilator circuits by simply flicking a switch. Depth and frequency of ventilation are initially set to mimic preanesthetic respiration, but are modified to maintain end tidal capnography (ETCO2) readings of between 35–45 mm Hg.

The primary goal of anesthetic monitoring should be to minimize morbidity by preventing, identifying, and correcting hypotension, bradycardia, arrhythmias, hypoxemia, hypercapnia, and metabolic disturbances. 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 preanesthetic examination to record heart and respiratory rates and character, and if possible collect baseline clinicopathologic data. Complete blood counts and full biochemistry profiles are not always possible (and indeed anesthesia is often used to facilitate blood collection); however, hematocrit, total protein, urea, and glucose can be run as a minimum using 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 preanesthetic values.

5.  Utilize equipment that will give you the most information with least problems; ultrasound Dopplers, end-tidal capnography 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 that must not be relied upon). Pulse oximetry readings (SpO2) < 90% equates to SaO2 < 60 mm Hg, or hypoxia. Indirect blood pressure readings can often be taken using a sphygmomanometer with an ultrasound Doppler.

6.  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 (and not required to answer the telephone). No anesthetic monitoring equipment can ever replace the attentive, experienced veterinary anesthetist.

7.  Maintain a contemporaneous record throughout the anesthetic period. Record quantitative readings and qualitative evaluations every 5 minutes on a dedicated anesthesia sheet that should form part of the medical record.

Recovery and Post-Op Care

Recovery and the immediate postoperative period can be just as critical in ensuring that your small mammal patient recovers completely. Indeed, the point of initial recovery and extubation can often be the most critical stage of the anesthetic procedure, because all the support mechanisms have been withdrawn.

In general:

1.  Discontinue anesthetic gas and/or give reversal drugs if applicable (e.g., flumazenil, Baxter Healthcare Corp).

2.  Once extubated, maintain on oxygen by loose fitting mask until fully conscious.

3.  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 ambulatory.

4.  Monitor the animal frequently and administer further fluids or antagonists as and when needed.

5.  Postoperative analgesia is important and should be a continuation/modification of the preemptive analgesic protocol employed for premedication.

6.  Monitor ferrets for postanesthetic hyperthermia for at least one hour after discontinuing anesthesia.


1.  AVMA. 2007.U.S. Pet Ownership & Demographics Sourcebook. American Veterinary Medical Association. (VIN editor: the original link could not be accessed on 3/28/13; an alternative link is https://ebusiness.avma.org/EBusiness50/files/productdownloads/sourcebook.pdf)

2.  Carpenter JW. Exotic Animal Formulary. 3rd ed. St. Louis, MO: WB Saunders Co.; 2005.

3.  Heard DJ. Anesthesia, analgesia, and sedation of small mammals. In: Quesenberry KE, Carpenter JW, eds. Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery. 2nd ed. Philadelphia, PA: WB Saunders Co.;2004:356–369.

4.  Heard DJ. Lagomorphs (rabbits, hares and pikas). In: West G, Heard D, Cauklett N, eds. Zoo Animal & Wildlife Immobilization and Anesthesia. ed. Ames, IA: Blackwell Publishing;2007:647–653.

5.  Heard DJ. Rodents. In: West G, Heard D, Cauklett N, eds. Zoo Animal & Wildlife Immobilization and Anesthesia. ed. Ames, IA: Blackwell Publishing;2007:655–663.


Speaker Information
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Stephen J. Divers, BVetMed, DZooMed, DECZM(Herp), DACZM, FRCVS
Department of Small Animal Medicine & Surgery
College of Veterinary Medicine
University of Georgia, Athens, GA

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