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Basic Approach to Veterinary Care
CHAPTER
13
Basic Approach to Veterinary Care
Jennifer Graham, DVM, Diplomate ABVP (Avian, Exotic Companion Mammal), Diplomate ACZM, and Douglas R. Mader, MS, DVM, Diplomate ABVP (Canine and Feline)
Housing Handling and Restraint Physical Examination Sample Collection Blood Collection Collection of Urine and Feces Dermatologic Sampling Cerebrospinal Fluid Tap Treatment Techniques Catheterization and Fluid Therapy Injection Techniques Oral Medications Enteral Support Vaccinations Pain Control Miscellaneous Procedures Anesthetic Delivery Nasolacrimal Cannulation Ear Cleaning
Rabbits are popular companion animals that present to veterinary clinics for routine and emergency care. It behooves the veterinarian, therefore, to become familiar with basic techniques used on rabbits in a clinical setting. Clinics equipped for seeing dogs and cats can be easily adapted to accommodate rabbits. This chapter reviews common procedures specific to rabbits that may be performed by the clinician.
HOUSING Housing requirements can be readily met for rabbits. Hospitalized rabbits can be kept in caging designed for avian and exotic animals, stainless steel cages designed for dogs and cats, or 174
specially designed hutch cages. It is wise to place a thick towel on the bottom of the cage to prevent the rabbit from slipping on the smooth surface and injuring its back. A rubberized mat can provide the same traction and has the added benefit of allowing urine and feces to fall through, preventing soiling of the patient. A rabbit hutch can be easily and inexpensively constructed or purchased from a pet or feed store. Hutch cage units can also be adapted for use as tabletop cages with built-in catch pans or can be suspended with wire, as is commonly done in multianimal rabbitries. Cage floors should be constructed of 14-gauge wire mesh. The mesh openings should be rectangular and no greater in size than 1 cm by 2.5 cm. This facilitates cleaning and allows feces to drop through the floor but is not so large that a rabbit might accidentally get its foot stuck. A portion of the floor should be solid, giving the animal a place to rest and helping to prevent the rabbit’s hocks from becoming sore. Keep in mind that wood is difficult to sanitize and is not permitted in facilities regulated by the U.S. Department of Agriculture. Keep a supply of good-quality feed available for hospitalized rabbits. Rabbits can be finicky eaters, so check with the owner before hospitalizing a rabbit to find out what foods the rabbit prefers. If the diet is of poor quality, it may be necessary to offer the rabbit some of the food to which it is accustomed while gradually introducing a more appropriate diet. A rapid change of diet, even a change from a poor diet to a proper one, may cause gastrointestinal upset and anorexia (see Chapters 14 and 15).3 Fresh water should always be available. Consult the owner to learn what type of watering system the rabbit uses, although rabbits easily learn to drink from sipper bottles. If water crocks are used, they should be made of a heavy ceramic so that they will not easily be tipped over. Bowls with high sides are recommended because rabbits tend to hang their dewlaps in the water when they drink. If the sides of the bowl are too low, this chronic wetting can lead to “wet dewlap” disease, which is an easily preventable moist dermatitis most often associated with colonization by Pseudomonas species. All water containers should be cleaned daily. Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
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Fig. 13-2 Restrain a stressed rabbit by covering its eyes and supporting its hindquarters.
Fig. 13-1 Restrain the rabbit with one hand under its thorax and the other hand supporting its hindquarters.
If possible, hospitalize rabbits in an area separate from potential or perceived predators. Sounds and smells from hospitalized dog, cat, or ferret patients may stress sensitive rabbits.7 Care should be taken to avoid overheating hospitalized patients. Warm areas appropriate for sick birds or reptiles may cause heat stress in euthermic or hyperthermic small mammal patients.
HANDLING AND RESTRAINT The rabbit’s skeleton represents only 7% to 8% of its body weight (as opposed to 12%-13% in cats).8 With powerful musculature on the hind limbs and the delicate nature of the skeleton, rabbits are vulnerable to fractures of the back and hind limbs. Rabbits are easily stressed and care should be taken to reduce stress to the patient during examination. Proper handling and housing of patients is necessary to avoid injury. When a rabbit is being moved, support of the hindquarters is vital to prevent injury. Rabbits used to handling can be carried with one hand under the thorax or by holding the scruff, with the second hand supporting the hindquarters (Fig. 13-1). Fractious rabbits should be carried with the head under the handler’s arm so as to minimize stress by covering the eyes; the hindquarters are supported at the same time (Fig. 13-2). Place a rabbit in a cage with its rear facing the back of the cage while supporting the hindquarters to help reduce chances of injury from the rabbit kicking. In examining a rabbit or placing it in a cage, use a nonslip mat to avoid sliding and injury. Control of the rabbit should be maintained at all times during transport and examination. A towel can be used to wrap the patient and cover the head to prevent struggling when the rabbit is transferred from a carrier to a table for examination. In similar fashion, a towel can be wrapped around the rabbit if an assistant is not available to help facilitate physical examination or the administration of medicines (Fig. 13-3). It is important to avoid overheating when a towel is being used to facilitate handling or examination. As an alternative, place your hand over the eyes to calm the
Fig. 13-3 To help facilitate physical examination or the administration of medicines when an assistant is not available, a towel can be wrapped around the rabbit to restrain it.
rabbit. Rabbits are obligate nasal breathers, so care should be taken to avoid obstructing the nostrils with a towel or hand. Because some rabbits become calm when placed on their backs, the handler can sit on the floor with the rabbit in his or her lap, hindquarters toward the handler’s body. The incisors, abdomen, genitalia, and feet can then be examined (Fig. 13-4). It is preferable for the rabbit to be examined in a room away from “predator” species such as ferrets, dogs, and cats; noises or smells from these animals may stress the rabbit.
PHYSICAL EXAMINATION In general, performance of the rabbit’s physical examination is much like that of other species.6 Observe the rabbit at a distance prior to initiating examination. Take note of movement of the rabbit and document any neurologic or musculoskeletal abnormalities. Check for any difficulty breathing and monitor general stress level. Place the rabbit in a quiet, oxygenated cage prior to examination if dyspnea is noted. Thermal support should be provided if the rabbit is hypothermic, taking care to
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Fig. 13-4 Restraint of a rabbit while the handler sits on the floor with the rabbit in his or her lap, hindquarters toward the handler’s body. Examination of the incisors, abdomen, genitalia, and feet can be accomplished with this technique.
Fig. 13-5 To allow for examination of the ventrum, nail trimming, and obtaining body temperature, cradle the rabbit on its back in a C-shape with the hind end supported.
avoid overheating. Although rare, a rabbit may be too stressed to undergo an examination without sedation. A sedative such as midazolam, at a dosage of 0.5 to 1.0 mg/kg IM, is often very effective in calming the animal.4 A systematic approach is needed for the examination. Take the body temperature early because the body temperature may increase from the stress of the exam.1 A rabbit can be cradled on its back in a ‘C-shape’ with the hind end supported to allow for examination of the ventrum, trimming of the nails, and taking the animal’s temperature (Fig. 13-5). Avoid using a glass thermometer because it can break if the patient struggles. In examining the rabbit, use a nonslip surface to help keep the animal from slipping and injuring itself (Fig. 13-6). The heart and respiratory rates should be calculated before the rabbit is stressed. It can be challenging though, to differentiate heart and lung sounds on auscultation; respiratory sounds superimposed over the heartbeats may create the false impression of a heart murmur. Note any ocular or nasal discharge in examining the head. The tympanic membrane should be visualized on
Fig. 13-6 Use a nonslip surface to prevent the rabbit from slipping and injuring itself while it is being examined.
otic examination. Lymphadenopathy is rare in rabbits, whose lymph nodes are located as in other species. Skin and coat quality should be documented; the plantar surface of the feet should be examined for any signs of pododermatitis. A complete oral examination is an important part of the assessment. Palpate the face and ramus of the mandible for any evidence of swelling or tooth elongation. Although the mouth can be examined without using anesthesia, rabbits are often reluctant to allow access to the oral cavity. Therefore the examination may have to be performed using light sedation or general anesthesia. Lesions may be missed when otoscopic cones are used to examine the mouth of an unanesthetized patient. Nasal or vaginal specula with an attached light source can provide a better field of view than the otoscopic cone. Rabbit mouth gags and cheek dilators are commercially available; however, lesions are occasionally missed in live patients using these instruments. To examine the oral cavity in a sedated patient, place the mouth gag over the incisors to hold the mouth open and insert the cheek dilator. Dental cameras or endoscopy, good lighting, suction, and magnification can further aid in visualization of the oral cavity. Chapter 32 offers more details and sources of instrumentation. Palpate the abdomen for masses, distention, gas, or other abnormalities. Tensing of abdominal muscles or tooth grinding may be signs of discomfort. A firm, or dough-like stomach, increased gas or fluid within the intestinal tract, and absence of normal intestinal sounds on auscultation may be noted with gastrointestinal disorders.12 Document and investigate any organomegaly.
SAMPLE COLLECTION BLOOD COLLECTION Sites used for venipuncture in rabbits can include the marginal ear veins, central ear artery, jugular vein, cephalic vein, and lateral saphenous vein. Because hematoma formation, bruising, or vessel thrombosis and skin sloughing can result, use of the ear veins and, in some breeds, the central ear artery is not ideal. Use alcohol to part the fur and allow for visualization of the vessel; clipping or plucking of the fur may be useful in some
CHAPTER 13 Basic Approach to Veterinary Care
Fig. 13-7 The rabbit’s lateral saphenous vein.
cases. Avoid damaging the delicate skin of the rabbit if clippers are used. Cephalic veins are accessible in rabbits but can be difficult to locate and hold off in smaller breeds. Venipuncture of this vein, however, should be avoided so as to preserve the integrity of the vessels if catheter placement is necessary. Additionally, venipuncture of the cephalic veins may be more stressful for the animal as the handler is working near the rabbit’s head. The lateral saphenous vein is an ideal site for venipuncture in the rabbit (Fig. 13-7). If needed, restrain the rabbit in a towel with the head covered while a rear limb is gently extended. The restrainer holds off the vein with pressure across the proximal thigh. The vessel is readily accessible across the lateral surface of the tibia just proximal to the hock. Apply gentle pressure or a pressure wrap briefly over the venipuncture site to prevent hematoma formation after the sample is obtained. Rabbits have large paired jugular veins, which are often a preferred venipuncture site for very calm or sedated rabbits or when a larger amount of blood (e.g., for blood transfusion) is being collected. However, it is difficult to visualize the jugular vein in overweight rabbits or in females with large dewlaps. For jugular venipuncture, shave the neck over the midtrachea cranial to the thoracic inlet. Position the rabbit with the front legs held over the edge of the table and the head extended up, avoiding overextension of the head. Alternatively, wrap the rabbit in a towel and place it in dorsal recumbency, extending the head and neck to allow for visualization and sampling from the jugular vein (Fig. 13-8). While cardiocentesis in rabbits is commonly performed in research for obtaining large blood samples or terminal exsanguination under anesthesia, this technique is not appropriate for clinical settings. Tables 13-1 and 13-2 contain reference ranges for hematologic and plasma biochemical values in rabbits. Hormone levels in rabbits are presented in Table 13-3.
COLLECTION OF URINE AND FECES A rabbit’s bladder can be gently pressed to obtain a urine sample during examination. To decrease chances of iatrogenic bladder rupture, however, cystocentesis may be preferable to manual bladder expression. Cystocentesis in the rabbit is similar to that in other small animal species. Although sedation or anesthesia
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Fig. 13-8 Two-person jugular venipuncture.
Table 13-1 Reference Ranges for Hematologic Values in the Rabbit12 Erythrocytes Hematocrit Hemoglobin Mean corpuscular volume Mean corpuscular hemoglobin Mean corpuscular hemoglobin concentration Platelets Leukocytes Neutrophils Lymphocytes Monocytes Eosinophils Basophils
5.1-7.9 × 106/μL 33%-50% 10.0-17.4 g/dL 57.8-66.5 μm3 17.1-23.5 pg 29-37 g/dL 250-650 × 103/μL 5.2-12.5 × 103/μL 20%-75% 30%-85% 1%-4% 1%-4% 1%-7%
Table 13-2 Reference Ranges for Serum Biochemistry Values in the Rabbit12 Serum protein Albumin Globulin Glucose Blood urea nitrogen Creatinine Total bilirubin Cholesterol Total lipids Calcium Phosphorus Sodium Potassium Chloride Bicarbonate Amylase Alkaline phosphatase Alanine aminotransferase Aspartate aminotransferase Lacticate dehydrogenase
5.4-8.3 g/dL 2.4-4.6 g/dL 1.5-2.8 g/dL 75-155 mg/dL 13-29 mg/dL 0.5-2.5 mg/dL 0.0-0.7 mg/dL 10-80 mg/dL 243-390 mg/dL 5.6-12.5 mg/dL 4.0-6.9 mg/dL 131-155 mEq/L 3.6-6.9 mEq/L 92-112 mEq/L 16-38 mEq/L 166.5-314.5 U/L 4-16 U/L 48-80 U/L 14-113 U/L 34-129 U/L
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Table 13-3 Hormone Levels in 29 Spayed/Neutered Rabbits Hormones Progesterone 17-Hydroxyprogesterone Androstenedione Testosterone Cortisol
Range (ng/mL) 0.11-0.46 0.75-22.2 0.80-4.00 0.02-0.04 4.64-11.2
From Fecteau KA, Deeb BJ, Rickel JM, et al. Diagnostic endocrinology: blood steroid concentrations in neutered male and female rabbits. J Exot Pet Med. 2007;16:256-259.
Table 13-4 Reference Ranges for Urinalysis Values in the Rabbit Urine volume Large breeds Average breeds Specific gravity Average pH Crystals present
Casts, epithelial cells, or bacteria present Leukocytes or erythrocytes present Albumin present
20-350 mL/kg per day 130 mL/kg per day 1.003-1.036 8.2 Ammonium magnesium phosphate, calcium carbonate monohydrate, anhydrous calcium carbonate Absent to rare Occasional Occasional in young rabbits
From Quesenberry KE. Rabbits. In: Birchard SJ, Sherding RG, eds. Saunders manual of small animal practice. Philadelphia: WB Saunders; 1994:1346.
reduces the chance that the stressed rabbit will struggle and damage internal structures, tranquilizers are often unnecessary. With the rabbit in dorsal recumbency, stretch the animal by holding the hind limbs in one hand and the scruff in the other. Alternatively, wrap the rabbit in a towel, hold the animal firmly, and cover its eyes. Locate the bladder on the ventral midline just cranial to the pelvic brim. A small-diameter needle (22- to 25-gauge) attached to a sterile 6-mL syringe provides an adequate sample for complete urinalysis after preparation of the antepubic region. Ultrasound can help with bladder visualization and detection of any bladder abnormalities. Urinalysis reference values are presented in Table 13-4. Urethral catheterization can be performed in sedated or anesthetized patients to collect a urine sample or for therapeutic flushing of the bladder in cases of urinary sludging. In most rabbits, a well-lubricated 9-Fr sterile catheter can be used for cathe terization. In the female, place the animal in sternal recumbency and locate the urethral os on the floor of the vagina. Placing the male in a sitting position allows for extrusion and catheterization of the penis. Fresh feces can be acquired from the enclosure of the rabbit or from the animal during physical examination. A direct smear of fresh feces in saline should be examined microscopically for the presence of protozoal organisms; several nonpathogenic
protozoa may thus be found. Fecal flotation is used to diagnose coccidia, cryptosporidia, and helminths. Coccidiosis is a common cause of diarrhea in young rabbits.
DERMATOLOGIC SAMPLING Some rabbit mites may be visualized under low magnification of skin brushings. Cheyletiella, bacteria, and yeast can be found on acetate tape strips applied to the skin and examined microscopically.5 As in other mammalian species, skin scrapings can be used to diagnose some parasitic conditions including sarcoptic or demodectic mange. Fungal and bacterial cultures as well as biopsies can be performed on skin lesions. Dark-field microscopy is used to examine smears for Treponema paraluiscuniculi.
CEREBROSPINAL FLUID TAP The collection of cerebrospinal fluid (CSF) may be indicated to help diagnose neurologic disease. Anesthesia, not sedation, is mandatory. The techniques are similar to those used in cats. The best site to collect CSF is the cerebellomedullary cistern. Position the rabbit in lateral recumbency with the head flexed toward the chest at a 90-degree angle to the vertebral column. Shave the fur on the nape of the neck from the occipital protuberance to the level of the third cervical vertebra and laterally past the margins of the atlas. The cranial margins of the wings of the atlas and the occipital protuberance are the landmarks for needle placement. A 22-gauge, 1.0- to 1.5-in.-long spinal needle should enter the skin midway between these points and be directed perpendicular to the skin approximately toward the animal’s nose. A stylet is usually not necessary because of the relatively small size of most rabbits. After the needle has penetrated the dura and arachnoid membranes, watch carefully for the appearance of CSF. The fluid should be allowed to drip into empty glass or plastic tubes. Do not attach a syringe or manometer since the movements caused by syringe attachment will likely cause bleeding, contaminating the fluid (Allen Sisson, personal communication, 2009). If the fluid cannot be submitted to the laboratory right away, one drop of serum from the rabbit should be added to 10 drops of CSF to preserve the cells for cytological evaluation. If serum must be added for cell preservation to the other tube for delayed cytologic evaluation, another 10 drops must be collected in a separate tube for cell counts and protein analysis. Only 10 drops of fluid are needed in one tube if the fluid can be analyzed immediately. Normal values for constituents of CSF in rabbits are presented in Table 13-5. A lumbar puncture, not a cerebellomedullary injection, should be used if a myelogram is to be done, since the contrast media will generally not flow past an extradural obstruction if the injection is made cisternally.
TREATMENT TECHNIQUES CATHETERIZATION AND FLUID THERAPY While many hospitalized rabbits can be managed with subcutaneous fluids, critically ill animals require intravenous fluids via catheter in the cephalic (Fig. 13-9) or lateral saphenous vein. A 24- or 26-gauge catheter can be placed in smaller rabbits, while a 22-gauge catheter can be used in rabbits weighing more than 3 kg. Alternatively, a butterfly catheter can be taped in place for short-term or bolus fluid therapy. Although jugular catheters
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Table 13-5 Reference Ranges for Cerebrospinal Fluid in the Rabbit Constituent
Concentration
Glucose Urea nitrogen Creatinine Cholesterol Total protein Alkaline phosphatase Carbon dioxide Sodium Potassium Chloride Calcium Magnesium Phosphate Lactic acid Nonprotein N
75 mg/dL 20 mg/dL 17 mg/dL 33 mg/dL 59 mg/dL 5.0 U/dL 41.2-48.5 mEq/L 149 mEq/L 3.0 mEq/L 127 mEq/L 5.4 mEq/L 2.2 mEq/L 2.3 mEq/L 1.4-4.0 mg/dL 5.6-16.8 mg/dL
Fig. 13-10 Subcutaneous fluid administration in the loose skin located on the rabbit’s dorsum.
From Weisbroth SH, Flatt RE, Kraus AL. The biology of the laboratory rabbit. New York: Academic Press, 1974:65.
Fig. 13-11 Intramuscular injection into the large lumbar muscles on either side of the rabbit’s spine, just cranial to the pelvis.
Fig. 13-9 An intravenous catheter placed in the rabbit’s cephalic vein.
can be used, anesthesia is recommended for their placement to facilitate the procedure and reduce stress to the animal. Avoid placement of intravenous catheters in the marginal ear veins, because sloughing of the ear tips can occur. A rabbit’s normal water consumption is estimated to be 100 to 150 mL/kg per day. Care must be taken in administering intravenous fluids to rabbits to avoid volume overload; 50 to 70 mL/kg per day administered intravenously can usually be tolerated. Infusion pumps help to regulate fluid delivery. A combination of crystalloids and colloids can be used to treat hypovolemic shock. If peripheral vessels are collapsed from dehydration, intraosseous catheters can be placed in the proximal humerus, greater trochanter of the femur, or tibial crest. Administer fluids intraosseously until the patient is adequately rehydrated or an intravenous catheter is placed. Prior to inserting the intraosseous catheter, clip the fur and surgically prepare the selected site in the sedated rabbit. Using sterile gloves, palpate the site of insertion of and insert a spinal needle or intraosseous catheter ranging in size from 18- to 23-gauge and from 1 to 1.5 in. in length,
depending on the rabbit’s size, parallel to the long axis of the bone and into the medullary cavity. Flush the needle gently with sterile saline and attach a male adapter. Apply antimicrobial ointment to the insertion site and a light dressing applied to hold the catheter in place. The subcutaneous route of fluid administration can be used on most hospitalized rabbits that have normal blood pressure and are accepting oral feedings. Rabbits can easily tolerate 120 mL/kg per day of subcutaneous fluids divided into 2 or 3 treatments. The loose skin located on the dorsum of the rabbit is an ideal location for subcutaneous fluid administration (Fig. 13-10).
INJECTION TECHNIQUES Administer medications via the intraosseous, intravenous, intramuscular, subcutaneous, or oral route. Injection techniques in rabbits are similar to those used in cats. Intraosseous, intravenous, and subcutaneous routes are described in the discussion of catheterization and fluid therapy, above. Intramuscular injections can be given into the large lumbar muscles on either side of the spine, just cranial to the pelvis (Fig. 13-11). One person
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Fig. 13-12 Oral medication being administered to a rabbit
Fig. 13-13 Nasogastric tube in an obese adult male Flemish
wrapped in a towel in “burrito” fashion.
giant rabbit.
can restrain and administer an intramuscular injection in a rabbit by tucking the rabbit under an arm, as described under “Handling and Restraint.” To avoid damaging the sciatic nerve, the cranial aspect of the rear leg (i.e., the quadriceps) should be used if the hind limbs are used for injection. Because some medications, including enrofloxacin, may cause pain, necrosis, or abscessation when the agent is given undiluted into subcutaneous or muscular tissues, dilute such medications with fluids and administer them subcutaneously.
palatable. Because it will not easily pass through a nasogastric tube owing to its high fiber content, Oxbow Critical Care Fine Grind (Oxbow Animal Health) has recently been developed for use in feeding tubes. Syringe feeding is most often used in the anorectic rabbit but may be stressful to the animal. Several methods can be used to syringe feed a rabbit, depending on its size and demeanor.4 It is quickest to feed a rabbit directly out of a 60-mL syringe with a catheter tip if the animal will tolerate it. The tip of the syringe is introduced into the diastema and slowly depressed. In smaller patients or those rabbits that resist feeding, 1-mL syringes or small oral feeding syringes can be used. This method is more time-consuming but can be a very effective way to deliver the formula to some rabbits that would otherwise refuse feeding. Orogastric tubes can be used for single dosing but are inappropriate for chronic use. For orogastric feeding, premeasure and mark an 18- to 22-Fr round-tip rubber catheter for the distance from the mouth to the last rib. Keep the rabbit’s mouth open with a mouth gag, flex the rabbit’s neck, and pass the tube through the oropharynx into the stomach. Test the tube’s placement by auscultating as air is injected through the tube into the stomach or checking for negative pressure when the tube is backed into the esophagus. Unless the rabbit is very calm, sedation or anesthesia will be required to minimize stress. Nasogastric tubes are used in clinical settings but may be more stressful to maintain than syringe feeding. Measurement of the tube is as described for orogastric tubes, using the tip of the nose and the last rib as landmarks. A pediatric feeding tube (3.5 to 5 Fr) can be used for most rabbits. Place a topical anesthetic such as 2% lidocaine gel or several drops of pro paracaine (Ophthaine, Solvay Animal Health, Inc., Mendota Heights, MN) can be placed in the nasal opening several minutes prior to placement. Insert the tube through the ventromedial nasal meatus and pass it ventrally and medially with the head flexed. Check tube placement with a lateral radiograph or as described for orogastric tube placement. Secure the tube with a drop of cyanoacrylate glue on the furred area above the nose and with tape glued or sutured to the top of the head (Fig. 13-13).
ORAL MEDICATIONS Suspensions are preferable to tablets when medications are administered to rabbits orally. Tablets can be made into sus pensions by compounding pharmacists when needed, or, alternatively, some tablets can be crushed and placed into a favorite treat or jam. It is helpful to use a towel in which to wrap a fractious rabbit in “burrito” fashion when oral medications are being given (Fig. 13-12). Place the medication as far back as possible in the oral cavity to prevent the rabbit from spitting it out.
ENTERAL SUPPORT Anorectic rabbits are commonly presented for medical evaluation. Dental disease, gastrointestinal disease, neurologic disease, and other systemic diseases are often complicated by secondary anorexia.11 Failure to provide nutritional supplementation to an anorectic rabbit can result in hepatic lipidosis in as little as 2 to 3 days. Oxbow Critical Care (Oxbow Animal Health, Murdock, NE) is a timothy hay-based syringe-feeding formula that is mixed with water to provide an excellent high-fiber mixture for anor ectic herbivores. Reconstitution of the product is simple and results in a homogenous mixture. Soaking and blending pellets and greens with water is an alternative to this commercial diet but is more time-consuming to prepare and generally results in a less homogenous mixture. The Oxbow Critical Care diet has an excellent fiber level at 21% to 25% and is highly
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VACCINATIONS No routine vaccinations are recommended for pet rabbits.
PAIN CONTROL Pain control is important for successful recovery of the compromised rabbit (see Chapter 31). Pain assessment may be more difficult in rabbits than in other species because they typically do not vocalize when they are experiencing pain. The rabbit may sit very still in the back of the cage in a hunched position while grinding its teeth and being oblivious to its surroundings. Other clinical signs of pain in the rabbit may include decreased fecal production, head elevation, aggression, isolation, rapid shallow breathing.2 Preemptive analgesia is recommended whenever possible. Nonsteroidal anti-inflammatory drugs (NSAIDs) are estimated to be effective for 12 to 24 hours in rabbits, while the effects of opioid drugs may last for only a few hours. Buprenorphine is effective for 6 to 12 hours, while butorphanol is effective for 2 to 4 hours. Abdominal or visceral pain may respond better to opioid analgesia, while NSAIDs are generally more effective for somatic or integumentary pain. Multimodal analgesia, combining agents from different classes, should be more effective than a single agent alone and allows the agents to be used at lower dosages.
MISCELLANEOUS PROCEDURES ANESTHETIC DELIVERY There are a variety of ways to administer anesthetic agents to rabbits, including topical, injectable, inhalant, and combination protocols. Anesthesia involves many considerations, such as the stability of the patient, monitoring, anesthetic agents used, and others.9,10 Critically ill patients should be stabilized prior to anesthetic administration. Topical anesthesia may be useful prior to procedures such as catheter placement. A topical preparation containing 2.5% lidocaine and 2.5% prilocaine (lidocaine 2.5%/prilocaine 2.5% cream, Hi-Tech Pharmaceutical, Amityville, NY) enables percutaneous insertion of catheters into rabbit veins without causing any detectable pain or discomfort. After application, these topical preparations may take 45 to 60 minutes to become fully effective. A variety of injectable anesthetic/analgesic combinations have been used in rabbits (see Chapter 31). Injectable anesthetic protocols may include parasympatholytics, phenothiazines, benzodiazepines, alpha-2-adrenergic agonists, ketamine, propofol, and others. The veterinarian should be aware of the specifics of different anesthetic drugs used in rabbits; for example, the use of tiletamine/zolazepam has been associated with nephrotoxicity in rabbits. Epidural anesthesia/analgesia is becoming more commonplace with small mammals. Local anesthetics, alpha-2 adrenergic agonists, and opioid agonists have been injected into the epidural space of small mammals to control pain. The advantages of epidural anesthesia and analgesia include few or no systemic effects compared with intramuscularly or intravenously administered drugs, quicker recovery time because of decreased gas anesthetic needed, and postsurgical pain relief. Disadvantages may include inability to place a spinal needle because of
Fig. 13-14 Location of the nasolacrimal duct in a rabbit. The arrow indicates the opening of the single nasolacrimal duct medial to the lid margin in the conjunctiva of the lower eyelid.
the small size of the epidural and intervertebral space, potential for trauma to the spinal cord, and potential for death or serious complications if analgesia is administered incorrectly. Care must also be taken in calculating volumes of local anesthetic for infiltration so as to avoid toxicity. Inhalant anesthesia is the primary component of most anesthetic regimens in small mammals (see Chapter 31). Isoflurane and sevoflurane are commonly used inhalant anesthetic agents. Induction with an inhalant anesthetic is typically achieved either with an induction chamber or face mask. Rabbits, however, are generally premedicated to lower the excitatory response typically seen with induction. Restrain the animal carefully to prevent injury during this period. A variety of commercially made induction chambers and face masks are available for use with rabbits. Additionally, masks can be fashioned out of syringe cases or other materials. For maintenance of anesthesia, endotracheal intubation, using blind and direct techniques, is ideal to protect the upper airway and assist ventilation (see Chapter 31). To facilitate intubation, hyperextend the head and neck of the rabbit to allow for the alignment of the larynx and trachea with the oropharynx. Care should be taken to make sure that the rabbit is adequately premedicated and relaxed to allow for atraumatic intubation. In addition, it is important to avoid overextension of the neck, which can result in damage to the spine.
NASOLACRIMAL CANNULATION Nasolacrimal duct flushing is indicated to determine and/or restore patency of the nasolacrimal ducts when rabbits present with an ocular discharge. Common causes of nasolacrimal duct obstruction can include infectious agents and dental disease. The single nasolacrimal duct in a rabbit is located medial to the lid margin in the conjunctiva of the lower eyelid (Fig. 13-14). Most rabbits will allow flushing of the nasolacrimal duct without sedation after topical anesthetic drops are instilled. A lacrimal cannula or 24-gauge Teflon intravenous catheter can be used to flush the duct (Fig. 13-15).
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References
Fig. 13-15 Proper placement of nasolacrimal cannula for a nasolacrimal duct flush in a rabbit.
EAR CLEANING Ear cleaning may be indicated in cases of otitis externa. If a large amount of debris is present within the ear canal and the rabbit is anesthetized, insert a red rubber catheter into the ear canal and flush with warm saline. While using an otoscope, endoscope, or otoendoscope for visualization to minimize the chance of traumatizing the ear canal, an ear curette or cottontipped applicators can be used to carefully scoop out purulent debris. Alternatively, flush saline into the ear with gentle massage to soften the purulent debris, followed by removal of debris with a suction unit while visualizing the canal to avoid iatrogenic damage. Flushing should be avoided if the tympanic membrane is ruptured. Ear cleaning should be accompanied by
1. Antinoff N. Physical examination and preventive care of rabbits. Vet Clin North Am Exot Anim Pract. 1999;2:405-427. 2. Bradley T, Lightfoot T, Mayer J. Exotic pet behavior: birds, reptiles, and small mammals. St. Louis: WB Saunders; 2006;41–43. 3. Cheeke PR. Nutrition and nutritional diseases. In: Manning PJ, Ringler DH, Newcomer CE, eds. The biology of the laboratory rabbit. 2nd ed. San Diego: Academic Press; 1994:321-331. 4. Graham J. Common procedures in rabbits. Vet Clin North Am Exot Anim Pract. 2006;9:367-388. 5. Harcourt-Brown F. Textbook of rabbit medicine. Oxford: Butterworth-Heinemann; 2002:224–248. 6. Harkness JE, Wagner JE. The biology and medicine of rabbits and rodents. 4th ed. Baltimore: Williams & Wilkins; 1995:13–30. 7. Krauss AL, Weisbroth AH, Flatt RE, et al. Biology and diseases of rabbits. In: Fox JG, Cohen BJ, Loew FM, eds. Laboratory animal medicine. Orlando, FL: Academic Press; 1984:207-240. 8. Mader DR. Basic approach to veterinary care. In: Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: WB Saunders; 2004:147-154. 9. Meredith A, Crossley DA. Rabbits. In: Meredith A, Redrobe S, eds. BSAVA manual of exotic pets. 4th ed. Gloucester, Great Britain: British Small Animal Veterinary Association; 2002:76-92. 10. Meredith A, Flecknell P. Anaesthesia and perioperative care. In: BSAVA manual of rabbit medicine and surgery. 2nd ed. Gloucester, UK: British Small Animal Veterinary Association; 2006:154-165. 11. Oglesbee BL. The 5-minute veterinary consult: ferret and rabbit. Hoboken, NJ: Wiley-Blackwell; 2006:178–385. 12. Quesenberry KE. Rabbits. In: Birchard SJ, Sherding RG, eds. Saunders manual of small animal practice. 2nd ed. Philadelphia: WB Saunders; 2000:1493-1511.
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Jennifer Graham, DVM, Diplomate ABVP (Avian, Exotic Companion Mammal), Diplomate ACZM, and Douglas R. Mader, MS, DVM, Diplomate ABVP (Canine and Feline)
Housing Handling and Restraint Physical Examination Sample Collection Blood Collection Collection of Urine and Feces Dermatologic Sampling Cerebrospinal Fluid Tap Treatment Techniques Catheterization and Fluid Therapy Injection Techniques Oral Medications Enteral Support Vaccinations Pain Control Miscellaneous Procedures Anesthetic Delivery Nasolacrimal Cannulation Ear Cleaning
Rabbits are popular companion animals that present to veterinary clinics for routine and emergency care. It behooves the veterinarian, therefore, to become familiar with basic techniques used on rabbits in a clinical setting. Clinics equipped for seeing dogs and cats can be easily adapted to accommodate rabbits. This chapter reviews common procedures specific to rabbits that may be performed by the clinician.
HOUSING Housing requirements can be readily met for rabbits. Hospitalized rabbits can be kept in caging designed for avian and exotic animals, stainless steel cages designed for dogs and cats, or 174
specially designed hutch cages. It is wise to place a thick towel on the bottom of the cage to prevent the rabbit from slipping on the smooth surface and injuring its back. A rubberized mat can provide the same traction and has the added benefit of allowing urine and feces to fall through, preventing soiling of the patient. A rabbit hutch can be easily and inexpensively constructed or purchased from a pet or feed store. Hutch cage units can also be adapted for use as tabletop cages with built-in catch pans or can be suspended with wire, as is commonly done in multianimal rabbitries. Cage floors should be constructed of 14-gauge wire mesh. The mesh openings should be rectangular and no greater in size than 1 cm by 2.5 cm. This facilitates cleaning and allows feces to drop through the floor but is not so large that a rabbit might accidentally get its foot stuck. A portion of the floor should be solid, giving the animal a place to rest and helping to prevent the rabbit’s hocks from becoming sore. Keep in mind that wood is difficult to sanitize and is not permitted in facilities regulated by the U.S. Department of Agriculture. Keep a supply of good-quality feed available for hospitalized rabbits. Rabbits can be finicky eaters, so check with the owner before hospitalizing a rabbit to find out what foods the rabbit prefers. If the diet is of poor quality, it may be necessary to offer the rabbit some of the food to which it is accustomed while gradually introducing a more appropriate diet. A rapid change of diet, even a change from a poor diet to a proper one, may cause gastrointestinal upset and anorexia (see Chapters 14 and 15).3 Fresh water should always be available. Consult the owner to learn what type of watering system the rabbit uses, although rabbits easily learn to drink from sipper bottles. If water crocks are used, they should be made of a heavy ceramic so that they will not easily be tipped over. Bowls with high sides are recommended because rabbits tend to hang their dewlaps in the water when they drink. If the sides of the bowl are too low, this chronic wetting can lead to “wet dewlap” disease, which is an easily preventable moist dermatitis most often associated with colonization by Pseudomonas species. All water containers should be cleaned daily. Copyright © 2012 by Saunders, an imprint of Elsevier Inc.
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Fig. 13-2 Restrain a stressed rabbit by covering its eyes and supporting its hindquarters.
Fig. 13-1 Restrain the rabbit with one hand under its thorax and the other hand supporting its hindquarters.
If possible, hospitalize rabbits in an area separate from potential or perceived predators. Sounds and smells from hospitalized dog, cat, or ferret patients may stress sensitive rabbits.7 Care should be taken to avoid overheating hospitalized patients. Warm areas appropriate for sick birds or reptiles may cause heat stress in euthermic or hyperthermic small mammal patients.
HANDLING AND RESTRAINT The rabbit’s skeleton represents only 7% to 8% of its body weight (as opposed to 12%-13% in cats).8 With powerful musculature on the hind limbs and the delicate nature of the skeleton, rabbits are vulnerable to fractures of the back and hind limbs. Rabbits are easily stressed and care should be taken to reduce stress to the patient during examination. Proper handling and housing of patients is necessary to avoid injury. When a rabbit is being moved, support of the hindquarters is vital to prevent injury. Rabbits used to handling can be carried with one hand under the thorax or by holding the scruff, with the second hand supporting the hindquarters (Fig. 13-1). Fractious rabbits should be carried with the head under the handler’s arm so as to minimize stress by covering the eyes; the hindquarters are supported at the same time (Fig. 13-2). Place a rabbit in a cage with its rear facing the back of the cage while supporting the hindquarters to help reduce chances of injury from the rabbit kicking. In examining a rabbit or placing it in a cage, use a nonslip mat to avoid sliding and injury. Control of the rabbit should be maintained at all times during transport and examination. A towel can be used to wrap the patient and cover the head to prevent struggling when the rabbit is transferred from a carrier to a table for examination. In similar fashion, a towel can be wrapped around the rabbit if an assistant is not available to help facilitate physical examination or the administration of medicines (Fig. 13-3). It is important to avoid overheating when a towel is being used to facilitate handling or examination. As an alternative, place your hand over the eyes to calm the
Fig. 13-3 To help facilitate physical examination or the administration of medicines when an assistant is not available, a towel can be wrapped around the rabbit to restrain it.
rabbit. Rabbits are obligate nasal breathers, so care should be taken to avoid obstructing the nostrils with a towel or hand. Because some rabbits become calm when placed on their backs, the handler can sit on the floor with the rabbit in his or her lap, hindquarters toward the handler’s body. The incisors, abdomen, genitalia, and feet can then be examined (Fig. 13-4). It is preferable for the rabbit to be examined in a room away from “predator” species such as ferrets, dogs, and cats; noises or smells from these animals may stress the rabbit.
PHYSICAL EXAMINATION In general, performance of the rabbit’s physical examination is much like that of other species.6 Observe the rabbit at a distance prior to initiating examination. Take note of movement of the rabbit and document any neurologic or musculoskeletal abnormalities. Check for any difficulty breathing and monitor general stress level. Place the rabbit in a quiet, oxygenated cage prior to examination if dyspnea is noted. Thermal support should be provided if the rabbit is hypothermic, taking care to
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Fig. 13-4 Restraint of a rabbit while the handler sits on the floor with the rabbit in his or her lap, hindquarters toward the handler’s body. Examination of the incisors, abdomen, genitalia, and feet can be accomplished with this technique.
Fig. 13-5 To allow for examination of the ventrum, nail trimming, and obtaining body temperature, cradle the rabbit on its back in a C-shape with the hind end supported.
avoid overheating. Although rare, a rabbit may be too stressed to undergo an examination without sedation. A sedative such as midazolam, at a dosage of 0.5 to 1.0 mg/kg IM, is often very effective in calming the animal.4 A systematic approach is needed for the examination. Take the body temperature early because the body temperature may increase from the stress of the exam.1 A rabbit can be cradled on its back in a ‘C-shape’ with the hind end supported to allow for examination of the ventrum, trimming of the nails, and taking the animal’s temperature (Fig. 13-5). Avoid using a glass thermometer because it can break if the patient struggles. In examining the rabbit, use a nonslip surface to help keep the animal from slipping and injuring itself (Fig. 13-6). The heart and respiratory rates should be calculated before the rabbit is stressed. It can be challenging though, to differentiate heart and lung sounds on auscultation; respiratory sounds superimposed over the heartbeats may create the false impression of a heart murmur. Note any ocular or nasal discharge in examining the head. The tympanic membrane should be visualized on
Fig. 13-6 Use a nonslip surface to prevent the rabbit from slipping and injuring itself while it is being examined.
otic examination. Lymphadenopathy is rare in rabbits, whose lymph nodes are located as in other species. Skin and coat quality should be documented; the plantar surface of the feet should be examined for any signs of pododermatitis. A complete oral examination is an important part of the assessment. Palpate the face and ramus of the mandible for any evidence of swelling or tooth elongation. Although the mouth can be examined without using anesthesia, rabbits are often reluctant to allow access to the oral cavity. Therefore the examination may have to be performed using light sedation or general anesthesia. Lesions may be missed when otoscopic cones are used to examine the mouth of an unanesthetized patient. Nasal or vaginal specula with an attached light source can provide a better field of view than the otoscopic cone. Rabbit mouth gags and cheek dilators are commercially available; however, lesions are occasionally missed in live patients using these instruments. To examine the oral cavity in a sedated patient, place the mouth gag over the incisors to hold the mouth open and insert the cheek dilator. Dental cameras or endoscopy, good lighting, suction, and magnification can further aid in visualization of the oral cavity. Chapter 32 offers more details and sources of instrumentation. Palpate the abdomen for masses, distention, gas, or other abnormalities. Tensing of abdominal muscles or tooth grinding may be signs of discomfort. A firm, or dough-like stomach, increased gas or fluid within the intestinal tract, and absence of normal intestinal sounds on auscultation may be noted with gastrointestinal disorders.12 Document and investigate any organomegaly.
SAMPLE COLLECTION BLOOD COLLECTION Sites used for venipuncture in rabbits can include the marginal ear veins, central ear artery, jugular vein, cephalic vein, and lateral saphenous vein. Because hematoma formation, bruising, or vessel thrombosis and skin sloughing can result, use of the ear veins and, in some breeds, the central ear artery is not ideal. Use alcohol to part the fur and allow for visualization of the vessel; clipping or plucking of the fur may be useful in some
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Fig. 13-7 The rabbit’s lateral saphenous vein.
cases. Avoid damaging the delicate skin of the rabbit if clippers are used. Cephalic veins are accessible in rabbits but can be difficult to locate and hold off in smaller breeds. Venipuncture of this vein, however, should be avoided so as to preserve the integrity of the vessels if catheter placement is necessary. Additionally, venipuncture of the cephalic veins may be more stressful for the animal as the handler is working near the rabbit’s head. The lateral saphenous vein is an ideal site for venipuncture in the rabbit (Fig. 13-7). If needed, restrain the rabbit in a towel with the head covered while a rear limb is gently extended. The restrainer holds off the vein with pressure across the proximal thigh. The vessel is readily accessible across the lateral surface of the tibia just proximal to the hock. Apply gentle pressure or a pressure wrap briefly over the venipuncture site to prevent hematoma formation after the sample is obtained. Rabbits have large paired jugular veins, which are often a preferred venipuncture site for very calm or sedated rabbits or when a larger amount of blood (e.g., for blood transfusion) is being collected. However, it is difficult to visualize the jugular vein in overweight rabbits or in females with large dewlaps. For jugular venipuncture, shave the neck over the midtrachea cranial to the thoracic inlet. Position the rabbit with the front legs held over the edge of the table and the head extended up, avoiding overextension of the head. Alternatively, wrap the rabbit in a towel and place it in dorsal recumbency, extending the head and neck to allow for visualization and sampling from the jugular vein (Fig. 13-8). While cardiocentesis in rabbits is commonly performed in research for obtaining large blood samples or terminal exsanguination under anesthesia, this technique is not appropriate for clinical settings. Tables 13-1 and 13-2 contain reference ranges for hematologic and plasma biochemical values in rabbits. Hormone levels in rabbits are presented in Table 13-3.
COLLECTION OF URINE AND FECES A rabbit’s bladder can be gently pressed to obtain a urine sample during examination. To decrease chances of iatrogenic bladder rupture, however, cystocentesis may be preferable to manual bladder expression. Cystocentesis in the rabbit is similar to that in other small animal species. Although sedation or anesthesia
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Fig. 13-8 Two-person jugular venipuncture.
Table 13-1 Reference Ranges for Hematologic Values in the Rabbit12 Erythrocytes Hematocrit Hemoglobin Mean corpuscular volume Mean corpuscular hemoglobin Mean corpuscular hemoglobin concentration Platelets Leukocytes Neutrophils Lymphocytes Monocytes Eosinophils Basophils
5.1-7.9 × 106/μL 33%-50% 10.0-17.4 g/dL 57.8-66.5 μm3 17.1-23.5 pg 29-37 g/dL 250-650 × 103/μL 5.2-12.5 × 103/μL 20%-75% 30%-85% 1%-4% 1%-4% 1%-7%
Table 13-2 Reference Ranges for Serum Biochemistry Values in the Rabbit12 Serum protein Albumin Globulin Glucose Blood urea nitrogen Creatinine Total bilirubin Cholesterol Total lipids Calcium Phosphorus Sodium Potassium Chloride Bicarbonate Amylase Alkaline phosphatase Alanine aminotransferase Aspartate aminotransferase Lacticate dehydrogenase
5.4-8.3 g/dL 2.4-4.6 g/dL 1.5-2.8 g/dL 75-155 mg/dL 13-29 mg/dL 0.5-2.5 mg/dL 0.0-0.7 mg/dL 10-80 mg/dL 243-390 mg/dL 5.6-12.5 mg/dL 4.0-6.9 mg/dL 131-155 mEq/L 3.6-6.9 mEq/L 92-112 mEq/L 16-38 mEq/L 166.5-314.5 U/L 4-16 U/L 48-80 U/L 14-113 U/L 34-129 U/L
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Table 13-3 Hormone Levels in 29 Spayed/Neutered Rabbits Hormones Progesterone 17-Hydroxyprogesterone Androstenedione Testosterone Cortisol
Range (ng/mL) 0.11-0.46 0.75-22.2 0.80-4.00 0.02-0.04 4.64-11.2
From Fecteau KA, Deeb BJ, Rickel JM, et al. Diagnostic endocrinology: blood steroid concentrations in neutered male and female rabbits. J Exot Pet Med. 2007;16:256-259.
Table 13-4 Reference Ranges for Urinalysis Values in the Rabbit Urine volume Large breeds Average breeds Specific gravity Average pH Crystals present
Casts, epithelial cells, or bacteria present Leukocytes or erythrocytes present Albumin present
20-350 mL/kg per day 130 mL/kg per day 1.003-1.036 8.2 Ammonium magnesium phosphate, calcium carbonate monohydrate, anhydrous calcium carbonate Absent to rare Occasional Occasional in young rabbits
From Quesenberry KE. Rabbits. In: Birchard SJ, Sherding RG, eds. Saunders manual of small animal practice. Philadelphia: WB Saunders; 1994:1346.
reduces the chance that the stressed rabbit will struggle and damage internal structures, tranquilizers are often unnecessary. With the rabbit in dorsal recumbency, stretch the animal by holding the hind limbs in one hand and the scruff in the other. Alternatively, wrap the rabbit in a towel, hold the animal firmly, and cover its eyes. Locate the bladder on the ventral midline just cranial to the pelvic brim. A small-diameter needle (22- to 25-gauge) attached to a sterile 6-mL syringe provides an adequate sample for complete urinalysis after preparation of the antepubic region. Ultrasound can help with bladder visualization and detection of any bladder abnormalities. Urinalysis reference values are presented in Table 13-4. Urethral catheterization can be performed in sedated or anesthetized patients to collect a urine sample or for therapeutic flushing of the bladder in cases of urinary sludging. In most rabbits, a well-lubricated 9-Fr sterile catheter can be used for cathe terization. In the female, place the animal in sternal recumbency and locate the urethral os on the floor of the vagina. Placing the male in a sitting position allows for extrusion and catheterization of the penis. Fresh feces can be acquired from the enclosure of the rabbit or from the animal during physical examination. A direct smear of fresh feces in saline should be examined microscopically for the presence of protozoal organisms; several nonpathogenic
protozoa may thus be found. Fecal flotation is used to diagnose coccidia, cryptosporidia, and helminths. Coccidiosis is a common cause of diarrhea in young rabbits.
DERMATOLOGIC SAMPLING Some rabbit mites may be visualized under low magnification of skin brushings. Cheyletiella, bacteria, and yeast can be found on acetate tape strips applied to the skin and examined microscopically.5 As in other mammalian species, skin scrapings can be used to diagnose some parasitic conditions including sarcoptic or demodectic mange. Fungal and bacterial cultures as well as biopsies can be performed on skin lesions. Dark-field microscopy is used to examine smears for Treponema paraluiscuniculi.
CEREBROSPINAL FLUID TAP The collection of cerebrospinal fluid (CSF) may be indicated to help diagnose neurologic disease. Anesthesia, not sedation, is mandatory. The techniques are similar to those used in cats. The best site to collect CSF is the cerebellomedullary cistern. Position the rabbit in lateral recumbency with the head flexed toward the chest at a 90-degree angle to the vertebral column. Shave the fur on the nape of the neck from the occipital protuberance to the level of the third cervical vertebra and laterally past the margins of the atlas. The cranial margins of the wings of the atlas and the occipital protuberance are the landmarks for needle placement. A 22-gauge, 1.0- to 1.5-in.-long spinal needle should enter the skin midway between these points and be directed perpendicular to the skin approximately toward the animal’s nose. A stylet is usually not necessary because of the relatively small size of most rabbits. After the needle has penetrated the dura and arachnoid membranes, watch carefully for the appearance of CSF. The fluid should be allowed to drip into empty glass or plastic tubes. Do not attach a syringe or manometer since the movements caused by syringe attachment will likely cause bleeding, contaminating the fluid (Allen Sisson, personal communication, 2009). If the fluid cannot be submitted to the laboratory right away, one drop of serum from the rabbit should be added to 10 drops of CSF to preserve the cells for cytological evaluation. If serum must be added for cell preservation to the other tube for delayed cytologic evaluation, another 10 drops must be collected in a separate tube for cell counts and protein analysis. Only 10 drops of fluid are needed in one tube if the fluid can be analyzed immediately. Normal values for constituents of CSF in rabbits are presented in Table 13-5. A lumbar puncture, not a cerebellomedullary injection, should be used if a myelogram is to be done, since the contrast media will generally not flow past an extradural obstruction if the injection is made cisternally.
TREATMENT TECHNIQUES CATHETERIZATION AND FLUID THERAPY While many hospitalized rabbits can be managed with subcutaneous fluids, critically ill animals require intravenous fluids via catheter in the cephalic (Fig. 13-9) or lateral saphenous vein. A 24- or 26-gauge catheter can be placed in smaller rabbits, while a 22-gauge catheter can be used in rabbits weighing more than 3 kg. Alternatively, a butterfly catheter can be taped in place for short-term or bolus fluid therapy. Although jugular catheters
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Table 13-5 Reference Ranges for Cerebrospinal Fluid in the Rabbit Constituent
Concentration
Glucose Urea nitrogen Creatinine Cholesterol Total protein Alkaline phosphatase Carbon dioxide Sodium Potassium Chloride Calcium Magnesium Phosphate Lactic acid Nonprotein N
75 mg/dL 20 mg/dL 17 mg/dL 33 mg/dL 59 mg/dL 5.0 U/dL 41.2-48.5 mEq/L 149 mEq/L 3.0 mEq/L 127 mEq/L 5.4 mEq/L 2.2 mEq/L 2.3 mEq/L 1.4-4.0 mg/dL 5.6-16.8 mg/dL
Fig. 13-10 Subcutaneous fluid administration in the loose skin located on the rabbit’s dorsum.
From Weisbroth SH, Flatt RE, Kraus AL. The biology of the laboratory rabbit. New York: Academic Press, 1974:65.
Fig. 13-11 Intramuscular injection into the large lumbar muscles on either side of the rabbit’s spine, just cranial to the pelvis.
Fig. 13-9 An intravenous catheter placed in the rabbit’s cephalic vein.
can be used, anesthesia is recommended for their placement to facilitate the procedure and reduce stress to the animal. Avoid placement of intravenous catheters in the marginal ear veins, because sloughing of the ear tips can occur. A rabbit’s normal water consumption is estimated to be 100 to 150 mL/kg per day. Care must be taken in administering intravenous fluids to rabbits to avoid volume overload; 50 to 70 mL/kg per day administered intravenously can usually be tolerated. Infusion pumps help to regulate fluid delivery. A combination of crystalloids and colloids can be used to treat hypovolemic shock. If peripheral vessels are collapsed from dehydration, intraosseous catheters can be placed in the proximal humerus, greater trochanter of the femur, or tibial crest. Administer fluids intraosseously until the patient is adequately rehydrated or an intravenous catheter is placed. Prior to inserting the intraosseous catheter, clip the fur and surgically prepare the selected site in the sedated rabbit. Using sterile gloves, palpate the site of insertion of and insert a spinal needle or intraosseous catheter ranging in size from 18- to 23-gauge and from 1 to 1.5 in. in length,
depending on the rabbit’s size, parallel to the long axis of the bone and into the medullary cavity. Flush the needle gently with sterile saline and attach a male adapter. Apply antimicrobial ointment to the insertion site and a light dressing applied to hold the catheter in place. The subcutaneous route of fluid administration can be used on most hospitalized rabbits that have normal blood pressure and are accepting oral feedings. Rabbits can easily tolerate 120 mL/kg per day of subcutaneous fluids divided into 2 or 3 treatments. The loose skin located on the dorsum of the rabbit is an ideal location for subcutaneous fluid administration (Fig. 13-10).
INJECTION TECHNIQUES Administer medications via the intraosseous, intravenous, intramuscular, subcutaneous, or oral route. Injection techniques in rabbits are similar to those used in cats. Intraosseous, intravenous, and subcutaneous routes are described in the discussion of catheterization and fluid therapy, above. Intramuscular injections can be given into the large lumbar muscles on either side of the spine, just cranial to the pelvis (Fig. 13-11). One person
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Fig. 13-12 Oral medication being administered to a rabbit
Fig. 13-13 Nasogastric tube in an obese adult male Flemish
wrapped in a towel in “burrito” fashion.
giant rabbit.
can restrain and administer an intramuscular injection in a rabbit by tucking the rabbit under an arm, as described under “Handling and Restraint.” To avoid damaging the sciatic nerve, the cranial aspect of the rear leg (i.e., the quadriceps) should be used if the hind limbs are used for injection. Because some medications, including enrofloxacin, may cause pain, necrosis, or abscessation when the agent is given undiluted into subcutaneous or muscular tissues, dilute such medications with fluids and administer them subcutaneously.
palatable. Because it will not easily pass through a nasogastric tube owing to its high fiber content, Oxbow Critical Care Fine Grind (Oxbow Animal Health) has recently been developed for use in feeding tubes. Syringe feeding is most often used in the anorectic rabbit but may be stressful to the animal. Several methods can be used to syringe feed a rabbit, depending on its size and demeanor.4 It is quickest to feed a rabbit directly out of a 60-mL syringe with a catheter tip if the animal will tolerate it. The tip of the syringe is introduced into the diastema and slowly depressed. In smaller patients or those rabbits that resist feeding, 1-mL syringes or small oral feeding syringes can be used. This method is more time-consuming but can be a very effective way to deliver the formula to some rabbits that would otherwise refuse feeding. Orogastric tubes can be used for single dosing but are inappropriate for chronic use. For orogastric feeding, premeasure and mark an 18- to 22-Fr round-tip rubber catheter for the distance from the mouth to the last rib. Keep the rabbit’s mouth open with a mouth gag, flex the rabbit’s neck, and pass the tube through the oropharynx into the stomach. Test the tube’s placement by auscultating as air is injected through the tube into the stomach or checking for negative pressure when the tube is backed into the esophagus. Unless the rabbit is very calm, sedation or anesthesia will be required to minimize stress. Nasogastric tubes are used in clinical settings but may be more stressful to maintain than syringe feeding. Measurement of the tube is as described for orogastric tubes, using the tip of the nose and the last rib as landmarks. A pediatric feeding tube (3.5 to 5 Fr) can be used for most rabbits. Place a topical anesthetic such as 2% lidocaine gel or several drops of pro paracaine (Ophthaine, Solvay Animal Health, Inc., Mendota Heights, MN) can be placed in the nasal opening several minutes prior to placement. Insert the tube through the ventromedial nasal meatus and pass it ventrally and medially with the head flexed. Check tube placement with a lateral radiograph or as described for orogastric tube placement. Secure the tube with a drop of cyanoacrylate glue on the furred area above the nose and with tape glued or sutured to the top of the head (Fig. 13-13).
ORAL MEDICATIONS Suspensions are preferable to tablets when medications are administered to rabbits orally. Tablets can be made into sus pensions by compounding pharmacists when needed, or, alternatively, some tablets can be crushed and placed into a favorite treat or jam. It is helpful to use a towel in which to wrap a fractious rabbit in “burrito” fashion when oral medications are being given (Fig. 13-12). Place the medication as far back as possible in the oral cavity to prevent the rabbit from spitting it out.
ENTERAL SUPPORT Anorectic rabbits are commonly presented for medical evaluation. Dental disease, gastrointestinal disease, neurologic disease, and other systemic diseases are often complicated by secondary anorexia.11 Failure to provide nutritional supplementation to an anorectic rabbit can result in hepatic lipidosis in as little as 2 to 3 days. Oxbow Critical Care (Oxbow Animal Health, Murdock, NE) is a timothy hay-based syringe-feeding formula that is mixed with water to provide an excellent high-fiber mixture for anor ectic herbivores. Reconstitution of the product is simple and results in a homogenous mixture. Soaking and blending pellets and greens with water is an alternative to this commercial diet but is more time-consuming to prepare and generally results in a less homogenous mixture. The Oxbow Critical Care diet has an excellent fiber level at 21% to 25% and is highly
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VACCINATIONS No routine vaccinations are recommended for pet rabbits.
PAIN CONTROL Pain control is important for successful recovery of the compromised rabbit (see Chapter 31). Pain assessment may be more difficult in rabbits than in other species because they typically do not vocalize when they are experiencing pain. The rabbit may sit very still in the back of the cage in a hunched position while grinding its teeth and being oblivious to its surroundings. Other clinical signs of pain in the rabbit may include decreased fecal production, head elevation, aggression, isolation, rapid shallow breathing.2 Preemptive analgesia is recommended whenever possible. Nonsteroidal anti-inflammatory drugs (NSAIDs) are estimated to be effective for 12 to 24 hours in rabbits, while the effects of opioid drugs may last for only a few hours. Buprenorphine is effective for 6 to 12 hours, while butorphanol is effective for 2 to 4 hours. Abdominal or visceral pain may respond better to opioid analgesia, while NSAIDs are generally more effective for somatic or integumentary pain. Multimodal analgesia, combining agents from different classes, should be more effective than a single agent alone and allows the agents to be used at lower dosages.
MISCELLANEOUS PROCEDURES ANESTHETIC DELIVERY There are a variety of ways to administer anesthetic agents to rabbits, including topical, injectable, inhalant, and combination protocols. Anesthesia involves many considerations, such as the stability of the patient, monitoring, anesthetic agents used, and others.9,10 Critically ill patients should be stabilized prior to anesthetic administration. Topical anesthesia may be useful prior to procedures such as catheter placement. A topical preparation containing 2.5% lidocaine and 2.5% prilocaine (lidocaine 2.5%/prilocaine 2.5% cream, Hi-Tech Pharmaceutical, Amityville, NY) enables percutaneous insertion of catheters into rabbit veins without causing any detectable pain or discomfort. After application, these topical preparations may take 45 to 60 minutes to become fully effective. A variety of injectable anesthetic/analgesic combinations have been used in rabbits (see Chapter 31). Injectable anesthetic protocols may include parasympatholytics, phenothiazines, benzodiazepines, alpha-2-adrenergic agonists, ketamine, propofol, and others. The veterinarian should be aware of the specifics of different anesthetic drugs used in rabbits; for example, the use of tiletamine/zolazepam has been associated with nephrotoxicity in rabbits. Epidural anesthesia/analgesia is becoming more commonplace with small mammals. Local anesthetics, alpha-2 adrenergic agonists, and opioid agonists have been injected into the epidural space of small mammals to control pain. The advantages of epidural anesthesia and analgesia include few or no systemic effects compared with intramuscularly or intravenously administered drugs, quicker recovery time because of decreased gas anesthetic needed, and postsurgical pain relief. Disadvantages may include inability to place a spinal needle because of
Fig. 13-14 Location of the nasolacrimal duct in a rabbit. The arrow indicates the opening of the single nasolacrimal duct medial to the lid margin in the conjunctiva of the lower eyelid.
the small size of the epidural and intervertebral space, potential for trauma to the spinal cord, and potential for death or serious complications if analgesia is administered incorrectly. Care must also be taken in calculating volumes of local anesthetic for infiltration so as to avoid toxicity. Inhalant anesthesia is the primary component of most anesthetic regimens in small mammals (see Chapter 31). Isoflurane and sevoflurane are commonly used inhalant anesthetic agents. Induction with an inhalant anesthetic is typically achieved either with an induction chamber or face mask. Rabbits, however, are generally premedicated to lower the excitatory response typically seen with induction. Restrain the animal carefully to prevent injury during this period. A variety of commercially made induction chambers and face masks are available for use with rabbits. Additionally, masks can be fashioned out of syringe cases or other materials. For maintenance of anesthesia, endotracheal intubation, using blind and direct techniques, is ideal to protect the upper airway and assist ventilation (see Chapter 31). To facilitate intubation, hyperextend the head and neck of the rabbit to allow for the alignment of the larynx and trachea with the oropharynx. Care should be taken to make sure that the rabbit is adequately premedicated and relaxed to allow for atraumatic intubation. In addition, it is important to avoid overextension of the neck, which can result in damage to the spine.
NASOLACRIMAL CANNULATION Nasolacrimal duct flushing is indicated to determine and/or restore patency of the nasolacrimal ducts when rabbits present with an ocular discharge. Common causes of nasolacrimal duct obstruction can include infectious agents and dental disease. The single nasolacrimal duct in a rabbit is located medial to the lid margin in the conjunctiva of the lower eyelid (Fig. 13-14). Most rabbits will allow flushing of the nasolacrimal duct without sedation after topical anesthetic drops are instilled. A lacrimal cannula or 24-gauge Teflon intravenous catheter can be used to flush the duct (Fig. 13-15).
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SECTION II Rabbits appropriate medical diagnostics to determine the cause of disease. In severe cases of otitis externa, rabbits may benefit from medical treatment to reduce inflammation prior to ear cleaning.
References
Fig. 13-15 Proper placement of nasolacrimal cannula for a nasolacrimal duct flush in a rabbit.
EAR CLEANING Ear cleaning may be indicated in cases of otitis externa. If a large amount of debris is present within the ear canal and the rabbit is anesthetized, insert a red rubber catheter into the ear canal and flush with warm saline. While using an otoscope, endoscope, or otoendoscope for visualization to minimize the chance of traumatizing the ear canal, an ear curette or cottontipped applicators can be used to carefully scoop out purulent debris. Alternatively, flush saline into the ear with gentle massage to soften the purulent debris, followed by removal of debris with a suction unit while visualizing the canal to avoid iatrogenic damage. Flushing should be avoided if the tympanic membrane is ruptured. Ear cleaning should be accompanied by
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