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Clinical techniques in rabbits
Clinical Techniques in Rabbits Susan A. Brown, DVM
Rabbits are very popular house pets and they require the same level of veterinary care as the canine or feline patient. It is often necessary to perform a variety of clinical procedures for both routine preventative care and when attempting to diagnose and treat illness in these patients. Many practitioners are wary of handling rabbits and attempting these procedures due to their fragile nature. However, armed with a good understanding of the rabbit's behavior and physical limitations and with the use of appropriate manual or chemical restraint, the veterinarian can accomplish necessary clinical procedures without harm to the patient.
Copyright 9 1997 by W. B. Saunders Company. Key words: Rabbits, clinical techniques, venipuncture, diagnostics, restraint.
Rabbits as P a t i e n t s he domestic rabbit (Oryctolagus c u n i c u l u s ) aintains the physiology and behavior of a prey species. Even t h o u g h the patient may be a beloved pet and is handled frequently by its owners, it will respond to pain and stress like its wild ancestors. Fear in a rabbit can cause a fall in body temperature, renal ischemia, and an increase in catecholamine production that can interfere with anesthesia. ~ Rabbits that are in pain can b e c o m e anorectic, lethargic, and experience p o o r recovery from anesthesia or surgery. Severely stressed rabbits may die within a few hours despite all efforts to reverse the process. Prolonged stress can lead to gastric ulcers, cardiomyopathy, and alterations in gastrointestinal (GI) flora. 1 Despite this seemingly gloomy outlook, with careful handling, the rabbit patient can survive routine diagnostic and surgical procedures. Factors that increase success include frequent observation of the patient's behavior, gentle handling, quiet recovery and hospitalization areas, analge-
From Midwest Bird & Exotic Animal Hospital, Westchester,IL. Address reprint requests to Susan A. Brown, DVM, Midwest Bird & Exotic Animal Hospital, 1923 S. Mannheim Rd, Westchest~ IL 60154. Copyright 9 1997 by W.. B. Saunde~:sCompany. 1055-937X/97/0602-000255.00/0
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sia, and anesthesia when appropriate. It is preferable, as with most pets, to hospitalize rabbits for as short a time as possible and to return them to the h o m e environment quickly.
Restraint Before attempting to handle a rabbit, make careful observations of its behavior. Rabbits that are fearful will breathe rapidly, stare with wide open unblinking eyes and sit very still, but in a position that allows for a quick escape with the legs tucked up u n d e r the body. Angry rabbits will attempt to attack when approached, making a snorting noise and striking out with their front feet. They may also attempt to bite or strike the g r o u n d forcefully with their hind feet (thumping). Calm rabbits blink their eyes more frequently and have a relaxed appearance to the eye lids. They are curious about their surroundings and are not afraid to investigate. When a relaxed rabbit stops to rest, it stretches its body out as opposed to the tightly h u n c h e d posture of the tense rabbit. When presented with a frightened rabbit, move slowly and talk quietly. Work on the floor with the patient until it is securely restrained. Occasionally a tense rabbit will scream in fear at the slightest touch. This may be in response to an o d o r in the r o o m that is frightening (ie, a previous ferret patient who has a predatory odor) or to an extended period of exposure to alarming noises (ie, a reception area with barking dogs). Rabbits have fragile spines and powerful back legs. If handled improperly, a rabbit can fracture or luxate its lumbar vertebrae or hind limbs. Place carriers or boxes on the floor to prevent potential injuries from j u m p i n g or falling off a table. Remove a rabbit from its carrier or cage backwards to minimize fear by not exposing it immediately to excessive visual stimulation. If the rabbit is very nervous, use a towel to cover its eyes before picking it up. Once the pet is restrained, it can be moved to the examination table. As with other pets, do not have the owner
Seminars in A v i a n and Exotic Pet Medicine, Vol 6, No 2 (April), 1997: pp 86-95
Clinical Techniques in Rabbits
help with the restraint because the owner may be injured by the rabbit's sharp claws. In addition, using an inexperienced owner to help with restraint may increase the chances of injury to the rabbit's spine. Rabbits can be picked up a variety of ways. O n e m e t h o d is to grasp the loose skin over the n a p e of the neck and the shoulders and place the o t h e r hand u n d e r the rabbit's hind quarters and lift. A second m e t h o d is to grasp the thorax ventrally and place the other h a n d u n d e r the hind quarters and lift. A third m e t h o d that is particularly useful for fractious rabbits is to either grasp the thorax ventrally or by the nape of the neck with one h a n d and firmly grasp the lower l u m b a r area with the other h a n d (Fig 1). Grasp the l u m b a r area immediately cranial to the pelvis and completely enclose the lumbar vertebral column and associated musculature with the hand. This area of the spine is thus protected from a hyperextension caused by a powerful kick. Fractious rabbits usually stop trying to kick when held in this manner. If a rabbit has to be transported any distance, tuck its head u n d e r the b e n d o f the elbow and support its body along the forearm. This m e t h o d minimizes visual stimulation and keeps the pet calm. Use a towel, mat, or other material to provide a nonskid surface on the examining table. Rabbits can sustain serious injuries if they try to move quickly on a slippery table. An excellent
Figure 1. Restraint offractious rabbit byscruffofneck and lumbar grasp.
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Figure 2. First fold of the towel wrap restraint in rabbit. m e t h o d of restraint that is used for examination, medication and anesthesia is the towel restraint. Lay o p e n a bath towel on the examination table. Place the rabbit with its front feet at the edge of the center of the long side of the towel. Fold one side o f the towel over the rabbit firmly (Fig. 2), then fold the back towards the front to prevent the rabbit f r o m escaping. T h e most i m p o r t a n t part is the final fold of the towel on the remaining side. T h e final side needs to be pulled very firmly across the rabbit's neck and back enclosing the front feet. W h e n d o n e properly, only the rabbit's head should be showing, with the front feet firmly tucked into the towel and the rest of the body firmly restrained (Fig 3). T h e towel wrap facilitates examination of the eyes, ears, and teeth and can be used for administering oral medications and subcutaneous (SC) injections in the intrascapular area. T h e towel restraint can also be used when administering inhalant anesthesia by mask. Wrap the patient as described, be
Figure 3. Completed towel wrap restraint in rabbit.
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seated, and place the rabbit with its hind quarters in yotar lap and its back supported by your a b d o m e n and chest. The rabbit's head is now in an upright position, which makes it very accessible for placing a mask. Watch the rabbit's breathing pattern through the mask by observing the m o v e m e n t of the nares. A n o t h e r restraint m e t h o d is to place the rabbit on its dorsum to examine the ventrum and trim the nails. Use the lumbar grasp described above to pick the rabbit up and turn it over onto its back. Lay the patient in your lap with its head hanging slightly over your knees. Keep one hand restraining the thorax. Most rabbits will eventually relax into an almost hypnotic state, and it will be possible to complete an examination. Loud noises or pain will cause the rabbit to try to roll over quickly and escape. Trim the nails in this position with an assistant. Have the assistant grasp each forelimb at the level of the h u m e r u s in each h a n d and extend and abduct the legs slightl}: This will prevent the patient from roiling over and escaping. Train owners to restrain their pets this way. Encourage them to sit on the g r o u n d until they are confident of their technique to prevent potential injuries during escape attempts. It is a good idea for the inexperienced practitioner to initially practice this technique on or close to the ground. Rabbits are very sensitive to touch a r o u n d their eyes, ears, nose, and mouth. Make sure the rabbit is securely restrained and d o n ' t prolong the examination of this area. Once the head has been examined, remove the towel restraint and continue with tile examination. Some rabbits feel more comfortable if a section of towel is placed loosely over its head, which may give it a sense that it is hiding. Use mild sedation on rabbits that are so difficult to handle that a physical examination cannot be safely undertaken. There are many options available for sedation in the rabbit using either a single drug or a combination of agents. Diazepam at I to 2 m g / k g intramuscular (IM) or ketamine hydrochloride at 20 m g / k g IM are two possible choices.
be particular challenging, but should be examined on every rabbit because they are frequently sites of disease. An oral exam can be p e r f o r m e d without anesthesia. One option is to use an otoscope with a large cone inserted into the interdental space. The drawback to the otoscope cone is that the view is restricted and over time, the cone becomes damaged from the teeth. The frayed edges can cause abrasions of the gums. Alternately, use either a canine vaginal speculum or a h u m a n nasal speculum with a light attachm e n t (Fig 4). The metal blades are indestructible, and when spread, they allow a more complete view than an otoscope cone. This instrument can also be used when trimming molars in the awake patient. Use gentle technique when palpating the a b d o m e n of the rabbit, particularly tile stomach. Severe bruising and ruptures can occur with overzealous palpation. Use caution when inserting a rectal t h e r m o m e t e r to prevmlt tearing of the delicate rectal mucosa. Rabbit skin is very thin and tears easily. Use care when plucking matted hair particularly a r o u n d the perineum. Tears that occur in this area can be painful and have the potential for becoming infected.
Analgesia As mentioned, the success of treatment can be decreased in rabbits that are severely stressed by pain or fear. The signs of fear have been described in the Restraint section. It has been well d o c u m e n t e d that rabbits experience pain, but it may be difficult to recognize these signs2 ,3 Prey
Examination A t h o r o u g h and complete physical examination should be p e r f o r m e d on the rabbit patient as in other species. The oral cavity and teeth can
Figure 4. Canine vaginal speculum used as oral speculum for examination of rabbit molars.
Clinical Techniques in Rabbits
species such as the rabbit tend to hide discomfort. T h e signs of pain are varied and include one or m o r e of the following: h u n c h e d appearance, tooth grinding, depression, anorexia, unresponsiveness to surroundings or to caretakers, hiding, reluctance to move or a b n o r m a l m o v e m e n t , and unusual aggressive behavior. Rabbits benefit f r o m the use of analgesics postsurgically. For m i n o r surgical procedures, one dose postoperatively may be all that is necessary. Good choices for postoperative analgesia include opioid agonist-antagonists b u p r e n o r phine (0.01 to 0.05 m g / k g intravenous [1V], SC every 8 to 12 hours) or b u t o r p h a n o l tartrate (0.1 to 0.5 m g / k g IV, SC, IM every 4 h o u r s ) 2 ,3 Naloxone (0.04 m g / k g IV, IM, SC) can reverse the sedative and depressant effects o f these drugs should it be necessary. 2 Some short-term sedative effects may be desirable to keep the patient m o r e relaxed in the hospital atmosphere. Nonsteroidal analgesics that can be used include aspirin (100 m g / k g by m o u t h every 48 hours), ibuprofen (10 to 20 m g / k g by m o u t h every 4 hours), and flunixin m e g l u m i n e (1.1 m g / k g SC every 8 hours or by m o u t h every 12 to 24 hours). These drugs are useful for pain associated with dental disease and arthritis. T h e author has used the injectable f o r m of flunixin diluted in a cherry-flavored syrup given orally with good results. The shelf-life of this preparation is at least 2 months j u d g i n g by clinical response. It is unknown if the long-term oral use of flunixin m e g l u m i n e could potentially result in gastric ulceration as in other species. T h e author does not use flunixin m e g l u m i n e for longer than 7 days consecutively without a 1- to 2-day b r e a k in dosing.
Hospitalization Hospitalize rabbits for as short a period as possible and handle only when necessary. Place hospital cages in the quietest area of the hospital. T h e sound of constantly barking dogs is extremely upsetting to the already c o m p r o m i s e d rabbit patient. Some rabbits will not defecate or urinate in their cage unless provided with a litter pan. Do not use clay cat litter material because it can be ingested resulting in a fatal gastrointestinal tract (GIT) impactiou. Use only digestible pelleted bedding. A hide box can help frighte n e d rabbits feel m o r e secure. Do not use r u b b e r
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or plastic mats on the cage floor because there is a potential for ingestion and G I T obstruction. Place debilitated rabbits on a soft, absorbent bedding such as synthetic fleece or thick p a p e r or cloth towels. Provide fresh greens such as parsley, dandelion greens, r o m a i n e lettuce, endive, beet tops, and carrot tops along with good quality grass hay and rabbit pellets. Use heavy crocks or attach food and water bowls to the cage to minimize spillage and d a m p cages. If possible, use the type of water container to which the rabbit is accustomed. Rabbits do not tolerate high humidity or high environmental temperatures. Their o p t i m m n t e m p e r a t u r e range is 40~ to 85~ with the mid to lower 70's preferred. '2 Closely m o n i t o r rabbits maintained in incubators to prevent hyperthermia.
Surgical Considerations This report is not intended to include a comprehensive discussion of surgery or anesthesia. Included in this section are a few special considerations for the rabbit patient. Use a heat source to maintain the rabbit's body t e m p e r a t u r e for major surgical procedures. T h e rabbit's body t e m p e r a t u r e can d r o p precipitously during surgery, especially during abdominal procedures. Use a rectal t h e r m o m e t e r to d e t e r m i n e if the patient needs additional heat during surgery or a heated cage during the postoperative period. It is not necessary to fast rabbits before surgery because they are unable to vomit. T h e exception is GIT surgery, where a 6- to 12-hour fast may be useful to reduce the GIT contents. Due to the nature of the rabbit GIT physiology, it is impossible to completely e m p t y the GIT contents even with fasts longer than 12 hours. Extended fasts are potentially dangerous in the rabbit and should not be attempted. In addition, fasting may p r o l o n g the a m o u n t of time it takes for the patient to resume eating postsurgically. Elevate the rabbit's h e a d and thorax when u n d e r anesthesia to minimize the pressure on the small thoracic cavity, particularly in overweight animals. Postsurgical abdominal adhesions are comm o n in the rabbit. To mir, imize this problem, wash the powder off surgic)d gloves, handle the tissues as gently and as little as possible, and use synthetic absorbable suture material (as o p p o s e d
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to chromic gut). Postsurgically use verapamil (200 m c g / k g intraperitoneal [IP] every 8 hours for nine doses) to prevent adhesions if there is extensive tissue handling. 2 Verapamil (Solo Pak Laboratories, Inc, Elk Grove Village, IL) is not necessary for uncomplicated routine procedures such as ovariohysterectomies and castrations. Rabbits can remove skin sutures with their incisors. Most rabbits will tolerate skin staples, but they are difficult to place in areas of the body where the skin is thin, such as the a b d o m e n . In addition, rabbits can occasionally pull out staples and then ingest t h e m resulting in a potential for the ingested staples to lodge in the G I T or the oral cavity. Use m o n o f i l a m e n t synthetic 4-0 suture material in a subcuticular pattern to close the skin. Subcuticular sutures are inaccessible to the patient. In addition, apply a small a m o u n t of surgical glue along the incision to bring the skin edges into apposition. Minimize the use of restraint apparatuses in the rabbit such as Elizabethan collars. They are not well tolerated and they may cause the patient to b e c o m e anorectic and depressed. In addition, the rabbit may not be able to access its nutritionally i m p o r t a n t cecotropes. Use several layers of bandaging to protect catheters or splints and reapply as needed. Use large tags of adhesive tape attached to the bandage to attract the rabbit to chew the tags instead of the bandage. Rabbits usually ignore bitter-tasting substances applied to areas where chewing is to be discouraged.
Medication Techniques Oral Medication Administering medications to a rabbit patient can be challenging. Rabbits will accept oral preparations if they are in liquid form, sweetened, and of a small volume. Use the towel restraint and place the rabbit in your lap with the head slightly elevated. Place the syringe tip into the interdental space between the incisors and premolars and give the medication slowly. Some rabbits learn to take their medication o f f a spoon when mixed with substances such as fruit j a m (strawberry is a favorite), p e a n u t butter, cherry-, orange-, or banana-flavored syrups, fruit juice, molasses, m a s h e d banana, blenderized greens, peanuts (use only two or three dry roasted unsalted, not raw), or dried fruit (raisins, apricots). Be patient when syringe feeding large vol-
umes of material by syringe. Do not a t t e m p t to syringe feed a severely depressed or debilitated rabbit. Use a nasoesophageal (NE) tube for these cases and for rabbits that are either too fractious to handle or those that refuse to take the syringe feeding. T h e r e are m a n y successful syringe feeding formulas for rabbits. For shortt e r m feeding, use canned p u m p k i n (not p u m p kin pie filling) which is high in fiber and easy to administer. A n o t h e r possibility for m o r e longterm feeding consists of one part g r o u n d rabbit pellets or alfalfa powder (found in health food stores) to one part blenderized fresh green leafy vegetables or j a r r e d strained vegetable baby food. Add to this mixture e n o u g h oral electrolyte solution to make a cream soup consistency. Administer at approximately 30 m L p e r 2 kg body weight every 8 to 12 hours. Use a syringe with a catheter tip to avoid the n e e d to strain the mixture. Administer pills to rabbits by crushing t h e m and mixing them with one of the palatable substances described above. Alternatively, give the pill directly to the patient by using the index finger to push it through the interdental space and as far back into the m o u t h as possible.
NE Feeding Tube A NE tube is invaluable in rabbit patients that are either too debilitated or fractious to take oral medications or feedings. ~ Rabbits tolerate NE tubes well and they can be left in place for several weeks if there are no complications (Figs 5 and 6). Although complications of this p r o c e d u r e are u n c o m m o n , they include rhinitis, epistaxis, dacryocystitis and esophageal reflux. 4 Use a 5 to 8 French feeding tube. The clear h u m a n pediatric tubes have the advantage of being radiopaque, allowing visualization of the contents of the tube and having an attached cap on the end to prevent tube contamination. Make additional holes in the caudal aspect of the tube before placement. General anesthesia is not necessary to place the tube. Use the towel restraint and elevate the rabbit's head. Instill two or three drops of an ophthalmic anesthetic or diluted lidocaine into the nostril. 4,5 Alternatively, lubricate the tube with lidocaine jelly before passage. Mlow a few minutes for the anesthetic to take effect. Measure the tube length f r o m the nares to the caudal aspect of the s t e r n u m a n d m a r k with tape or marking pen. Lubricate the tube and place it in the ventral meatus directing
Clinical Techniques in Rabbits
Figure 5. Nasoesophageal tube in rabbit (frontal view). it medially and ventrally. 4 If the nasal cavity was adequately anesthetized, any discomfort will be minimal and transient. The tube should move without resistance. Put the rabbit's head in a normal flexed position as the pharynx is approached to encourage tube passage into the esophagus. Always assess p r o p e r placement by radiograph. The distal end of the tube can be located in the esophagus or the stomach. Sterile saline introduced through the tube may not elicit a cough response when the placement is in the trachea. Once the placement is assessed, suture the tube in place over the bridge of the nose and at the top of the head between the ears with tape butterflies. Rabbits rarely need any form of restraint to prevent the removal of the tube. Pass the tube-feeding formula through a fine
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strainer to prevent blockage of the tube. Flush the tube with saline or water before and after each feeding or medication. One possible formula that can be used for long-term maintenance of debilitated rabbits with NE tubes is two parts alfalfa powder (available in health food stores, high in fiber, and finer in consistency than ground rabbit pellets) plus one part blenderized green leafy vegetables or j a r r e d strained vegetable baby food plus one part Deliver 2.0 (Mead Johnson, Evansville, IN) plus e n o u g h oral electrolyte solution to make the formula into a thin consistency. Strain the mixture t h r o u g h a fine strainer. Use 30 mL per 2 kg body weight every 6 to 12 hours. Subsequent stools are formed, but small in size. Consider vitamin supplementation for patients that are debilitated for longer than a week a n d / o r are unable to p r o d u c e or eat their cecotropes. Rabbits can still eat normally with an NE tube in place. Some practitioners have had success with the use of pharyngostomy tubes in rabbits. The advantages over the NE tube is that a larger tube can be used and a thicker consistency of material can be administered. The disadvantage is that it requires a surgical procedure to place the tube. The technique for pharyngostomy tube placem e n t would be similar to the canine or feline patient.
Injectable Medications Use small gauge needles (22 to 27 gauge [G]) for most injections to minimize discomfort. Give SC injections and fluids in the dorsum of the neck or shoulder area. Minimal restraint is necessary for this procedure. Administer IM injections in the dorsal lumbar muscles cranial to the pelvis or in the cranial aspect of the quadriceps. Give IV injections in the lateral saphenous or cephalic veins. The jugular and femoral veins can also be used. Although many sources advocate the use of the marginal auricular vein or central auricular artery for injections, the practitioner should be cautious using these sites. Sloughing of the ear tip can occur. 6 This is most often due to the infusion of an irritating substance or damage to the blood vessel. These vessels should be used as a last resort for 1V injections.
Intravenous Catheters Figure 6. Nasoesophageal tube in rabbit (side view).
Use small gauge (22 to 25 G) over-the-needle catheters. Butterfly catheters do not work well
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because of their tendency to puncture the delicate wall of the vein with the slightest movement. Pluck or clip the fur over the vein and aseptically prepare the skin. Rabbit skin is thin and friable; therefore, it is unnecessary to incise the skin before placing an IV catheter. Use sufficient bandaging to prevent the removal of the catheter in the awake animal. Use IV catheters to administer warm fluids during any extensive surgical procedure. A syringe p u m p is invaluable in this small species to administer fluids at a prescribed rate and volume. Although many sources advocate the use of the marginal auricular vein or central auricular artery for IV catheterization, the practitioner is cautioned against its routine use for reasons described u n d e r Injectable Medications. The lateral saphenous vein affords a very accessible site for an indwelling catheter, particularly for use during surgery. This vein runs medial to lateral across the cranial aspect of the tarsus and has two branches that continue laterally up the leg. Place the catheter so that the injection cap will be above the plantar surface of the foot when the rabbit is sitting in a normal position. It is difficult to maintain a catheter in the lateral saphenous for extended periods in the awake animal due to the constant m o v e m e n t of the hind legs. Cephalic catheters are placed and taped in as in other species and can be maintained for extended period. Jugular catheters can be used, but sedation will be necessary in most patients for placement. Fluid administration guidelines in the rabbit are similar to the cat and dog. Maintenance fluid level for the rabbit is approximately 80 to 100 m L / k g every 24 hours. 5,7 Subcutaneous fluids are more often used due to the ease of administration. Rabbits have a large subcuticular space u n d e r the shoulders and neck. However, rabbits that are debilitated, severely stressed, or undergoing major surgical procedures should receive IV or intraosseous (IO) fluids to improve chances for survival. Subcutaneous fluids may be poorly absorbed in these cases.
Intraosseous Catheters Intraosseous catheters are invaluable in the debilitated rabbit where venous pressure is low or in the patient where IV catheterization is unsuccessful. Rabbits seem to tolerate IO cath-
eters better than IV catheters. Place IO catheters in the femur, humerus, or tibia, s The femur is most frequently used, and a catheter in this bone causes the least disruption to normal movement. Use a spinal needle (20 to 22 G 1.5 to 3 inch), if possible, due to the sturdy construction of the needle and the stylet, which prevents occlusion of the needle with bone. Alternatively, a 20- to 22G hypodermic needle can be used with a small gauge sterile surgical wire inserted in the needle bore as a stylet. Use anesthesia in all but the very docile or debilitated patient to place an IO catheter because this procedure can be painful. Use a local anesthetic in the awake patient. Clip the hair and surgically prepare the area over the head of the femur. Use sterile technique and make a small incision over the proximal aspect of the greater trochanter. Insert tile needle through the trochanteric fossa using firm pressure and gentle rotation. When the catheter has entered the marrow cavity, it should advance easily until the hub meets the skin. Remove the stylet and test patency and placement by slowly injecting 3 to 5 mL of sterile saline. If the needle is properly placed, there is little resistance to the injection, and the soft tissue a r o u n d the area does not swell. Place butterfly tape a r o u n d the needle hub and suture it to the skin. Place an injection cap on the needle and flush with heparinized saline/ Any substance that can be injected intravenously can be given intraosseously with the exception of some chemotherapy agents. The bone is not distensible; therefore, do not give fluids as a bolus. Administer medications slowly. A syringe p u m p is invaluable for slow, measured infusion into the IO catheter. IO catheters can be left in for several days. It is advisable to use systemic antibiotics while the catheter is in place and for 2 or 3 days after removal.
Blood Collection The blood volume of the rabbit is approximately 6% of the body weight. 9 Therefore, a 3,000-gm (approximately 6 pounds) rabbit has a blood volume of 180 mL. Up to 10% of this total can be collected safely in one sampling (18 mL). Up to 25% can be collected over a 2-week period. 2 For routine complete blood cell (CBC) counts and serum biochemistries, it is only necessary to obtain one half to 3 mL of blood.
Clinical Techniques in Rabbits
Figure 7. Venipuncture in rabbit using central auricular artery. Use small gauge needles (29 to 95 G) to obtain blood samples in rabbits. Do not use a toe nail clip for routine sampling. This procedure is painful, may result in infection, and may yield p o o r samples due to contamination with tissue fluids. The most accessible site to get a sample (1 to 3 mL) is the central auricular artery (Fig 7). Restrain the rabbit and apply alcohol to the ear to wet the fur. Flick the ear gently with a finger to cause the artery to become more prominent. If the rabbit is cold or frightened, warm the ear to raise the artery. Insert a 25-G needle or butterfly catheter into the artery without a syringe attached and allow the blood to flow into the collection container. In large rabbits, a vacutainer may be used, but in smaller rabbits, the suction will collapse the vessel. Once the needle is removed, apply pressure to the venipuncture site for several minutes or apply a temporary bandage to prevent h e m a t o m a formation. Blood collection from the marginal ear vein is difficult and often unsuccessful, especially in smaller breeds. The lateral saphenous vein, as previously described, is easy to access even in the awake rabbit and is of a sufficiently large size and length to be highly visible. Restrain the rabbit by scruffing it by the nape of the neck and hold it on its side using the other hand to grasp firmly a r o u n d the dorsal aspect of the quadriceps of the leg to be used for venipuncture. This occludes the vein, which allows better visualization. Gently pluck or clip the hair over a portion of the vein and apply alcohol to improve visibility. Use a 25-G needle attached to a syringe to draw the sample. Use firm pressure for at least a minute afterwards or a temporary bandage for hemostasis.
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The cephalic vein can be used, but it is very thin and fragile, particularly in smaller species. The restraint and collection m e t h o d is similar to the dog. Towel restraint is useful here. The jugular vein can be accessed in the awake animal in either sternal or dorsal recumbency. For sternal recumbency, hold the rabbit near the edge of a table with the head extended upwards and the feet extended downwards over the edge. 5 Use alcohol to help visualize the vein, which runs laterally from the thoracic inlet to the base of the ear. Place digital pressure just lateral to the thoracic inlet and visualize or palpate this superficial vein. Use a 92-G needle attached to a syringe to draw the blood. Alternatively, restrain the rabbit in a towel wrap and place it in dorsal recumbency with the neck extended. 3 Visualize or palpate the jugular vein and draw the blood as described in the previous m e t h o d of restraint. Cardiac and vena cava puncture have been described for the rabbit. It is the author's opinion that these methods have inherent risks, particularly in the hands of inexperienced phlebotomists and, therefore, are not appropriate use in for pet rabbits.
Bone Marrow Biopsy Bone marrow biopsies may be taken from a variety of sites including the long bones and pelvic bones, and the technique is similar to the cat or dog. Access the marrow cavity of the femur by using the technique previously described u n d e r Intraosseous Catheters. Attach a lO-mL syringe to the hub of the inserted needle and apply vigorous suction to retrieve bone marrow.
Cerebrospinal Fluid Collection Collect cerebrospinal fluid (CSF) by anesthetizing the rabbit, place it in lateral recumbency, and flex the head toward the sternum. Clip the hair and surgically prepare the area from the occipital protuberance to the third cervical vertebrae and laterally at least 3 cm on each side of the midline. Use the cranial margins of the wings of the atlas and the occipital protuberance as the landmarks for needle placement. Use a 22-G 1.5 inch spinal needle and place it midway between these points and direct it towards the nares. 6 Remove the styler after the dura is penetrated
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and look for spinal fluid to a p p e a r in the hub. Draw the sample and remove the needle.
Lacrimal Duct Cannulation A c o m m o n disorder of rabbits is dacryocystitis, which presents with unilateral or bilateral e p i p h o r a and facial dermatitis. Most often, this condition is treated by cannulating and flushing the lacrimal duct. This removes potential obstructive p u r u l e n t material, allows culturing of the flushed material, and improves the penetration of medication into the area. A variety of lacrimal cannulas may be used for this procedure, and either straight or curved models are acceptable. Accomplish cannulation of the lacrimal duct without a general anesthetic. Restrain the rabbit by the towel m e t h o d and apply an ophthalmic anesthetic to desensitize the affected eye. T h e r e is only one nasolacrimal canaliculus and puncturn in the rabbit located in the lower lid. Visualize the puncture by either pulling the lower lid down and rolling it outward or grasping the edge of the lower lid and pulling it directly laterally. Rabbits rarely object to this procedure. Cannulation is m o r e difficult in the presence of a persistent discharge and severe inflammation of the peripunctal area. Apply pressure below the p u n c t u r e to cause the lips of the p u n c t u m to pout, allowing better access to the duct. 7 Use 1 to 2 m L of sterile saline to gently flush the duct once the cannula is in place. Do not use e x t r e m e force to flush the duct or it may rupture. T h e saline should exit the nares if cannulation was successful. If the duct is blocked or the cannula is not placed properly, the saline will flush retrograde out the puncture. Try placing the cannula again should this occur. If the duct is ruptured, the saline will leak out into the s u r r o u n d i n g tissues, often into the retrobulbar space causing the eye to bulge laterally. If this happens, stop flushing immediately. T h e saline will be absorbed over the next 24 hours, and the eye will return to normal. If the lacrimal duct is ruptured, the healing process may r e n d e r it permanently strictured due to scar tissue formation.
Microbiological Culture Collection Techniques for obtaining microbiological cultures in the rabbit are similar to those in the dog and cat. Howevm, three areas bear special men-
tion because they are c o m m o n sites of disease in the rabbit. Rabbits have a predisposition to develop large abscesses filled with thick tenacious p u r u l e n t material. Practitioners often make the mistake of using a glob of p u r u l e n t material as their culture sample. This practice usually leads to a negative result. This material is c o m p o s e d of dead cells and phagocytized bacteria and is not a viable choice for sampling. Collect the culture by swabbing the wall of the abscess or draining tracts after the p u r u l e n t material has b e e n removed. Negative results are still possible, but the probability of a positive culture is increased with this method. Rabbits are p r o n e to u p p e r respiratory infections caused by a variety of infectious agents including Pasteurella multocida, Staphylococcus aureus, and Bordetella bronchiseptica. A d e e p nasal culture and sensitivity is an extremely useful tool for selecting effective antibiotic therapy. T h e culture is obtained quickly and with minimal discomfort in the awake rabbit. Seat the handler with the towel-restrained rabbit's hind quarters in the handler's lap and the rabbit's back supp o r t e d by the handler's a b d o m e n and chest. T h e nares are readily visible, and the rabbit is securely restrained in this position. Use a sterile calgi swab or minitip culturette and direct it medial and caudal into the nasal sinus as far as it will go then remove it immediately. T h e rabbit may j u m p slightly as the swab enters the sinus. A small a m o u n t of blood will be present on the swab. Rarely, a rabbit will exhibit mild epistaxis for a few minutes after sampling and will sneeze. Keep the rabbit wrapped in the towel, but allow it to return to its n o r m a l sitting position until the bleeding has stopped. Advise the client that pink-tinged fluid may be seen periodically at the nares for the next h o u r or two. As m e n t i o n e d in the section on Lacrimal Duct Cannulation, dacryocystitis is a c o m m o n problena in rabbits. This disease can be caused by the same infectious agents that cause u p p e r respiratory disease. The material flushed f r o m the lacrimal duct is easily cultured. Have an assistant hold a sterile swab just ventral to the nares without touching the skin. Allow the flushed material exiting the nasal sinus to drop onto the swab. One or two drops is sufficient. Obviously, do not use an antibiotic solution to flush the lacrimal ducts when obtaining a culture.
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Malocclusion and Associated Disease As s t a t e d in t h e E x a m i n a t i o n section, t h e oral cavity a n d t e e t h a r e f r e q u e n t l y a site o f disease in the rabbit and should be thoroughly examined. H o w e v e L t h e variety a n d c o m p l e x i t y o f d e n t a l disease p r e s e n t a t i o n s , i n c l u d i n g m a l o c c l u s i o n a n d a s s o c i a t e d infections, p r e c l u d e s a t h o r o u g h d i s c u s s i o n o f d i a g n o s t i c a n d t r e a t m e n t techn i q u e s in this article. T h e r e a d e r is e n c o u r a g e d to f u r t h e r r e s e a r c h t h e s e s y n d r o m e s )
Urine Collection U r i n e can b e c o l l e c t e d in t h e r a b b i t by f r e e c a t c h i n t o a n e m p t y litter b o x , by g e n t l e e x p r e s sion o f t h e b l a d d e r in t h e awake o r a n e s t h e t i z e d p a t i e n t , by cystocentesis, o r by u r i n a r y c a t h e t e r ization. E x p r e s s t h e b l a d d e r in t h e awake a n i m a l in t h e s a m e m a n n e r as in t h e d o g o r cat. U r i n e will b e v o i d e d caudally. T a k e c a r e n o t to use excessive f o r c e o r t h e b l a d d e r can b e severely b r u i s e d o r ruptured. Cystocentesis c a n usually b e p e r f o r m e d in t h e awake a n i m a l . However, if t h e p a t i e n t is f r a c t i o u s o r e x t r e m e l y stressed, use s e d a t i o n o r a n e s t h e s i a to p r e v e n t a p o t e n t i a l l a c e r a t i o n o f t h e b l a d d e r wall. R e s t r a i n t h e awake r a b b i t by h o l d i n g t h e scruff at t h e n a p e o f t h e n e c k in o n e h a n d a n d b o t h r e a r legs in t h e o t h e r h a n d . S t r e t c h t h e r a b b i t o u t a n d p l a c e in d o r s a l r e c u m b e n c y . 6 Clip a small a r e a o f h a i r a n d p e r f o r m a sterile p r e p a r a t i o n o f tile a n t e p u b i c r e g i o n . Use a small g a u g e n e e d l e (23 to 25 G) a t t a c h e d to a n a p p r o p r i a t e - s i z e d syringe to p e n e t r a t e t h e b l a d d e r wall a n d o b t a i n t h e s a m p l e . T h e m o s t c o m m o n use for u r i n a r y c a t h e t e r i z a tion is to e n a b l e f l u s h i n g o f t h e b l a d d e r to r e m o v e excessive c a l c i u m c a r b o n a t e p r e c i p i t a t e , o f t e n r e f e r r e d to as " s l u d g e . " A n i n d w e l l i n g u r e t h r a l c a t h e t e r m a y also b e u s e d to allow h e a l i n g o f injuries to t h e u r e t h r a c a u s e d by b i t e w o u n d s o r u r e t h r a l calculi. I n a d d i t i o n , u r i n a r y c a t h e t e r i z a t i o n is essential to p e r f o r m single a n d d o u b l e c o n t r a s t studies o f t h e b l a d d e r . Use a f l e x i b l e 3.5 to 6 F r e n c h u r i n a r y c a t h e t e r o r f e e d i n g tube. Use a stylet o f wire, if necessary, o r stiffen t h e c a t h e t e r by p l a c i n g it in t h e f r e e z e r
for a s h o r t p e r i o d . D o n o t use t o m c a t c a t h e t e r s because the likelihood of injuring the delicate u r e t h r a l wall is i n c r e a s e d . A n e s t h e t i z e o r s e d a t e t h e r a b b i t a n d p l a c e it in d o r s a l r e c u m b e n c y . E x t e n d t h e p e n i s in t h e m a l e r a b b i t by a p p l y i n g t e n s i o n a n d p r e s s u r e c r a n i a l l y at its base. Use a small vaginal s p e c u l u m o r o t o s c o p e to visualize t h e u r e t h r a l o p e n i n g in t h e f e m a l e . L u b r i c a t e t h e c a t h e t e r a n d pass it i n t o t h e b l a d d e r . Use w a r m e d saline to flush t h e b l a d d e r , u s i n g small a m o u n t s frequently. A d m i n i s t e r SC o r IV fluids to e n c o u r a g e u r i n a t i o n . If t h e c a t h e t e r is to b e left in place, s u t u r e it by m e a n s o f b u t t e r f l y t a p e to t h e p e r i n e u m . R a b b i t s will c h e w o u t c a t h e t e r s unless t h e y a r e severely d e b i l i t a t e d , a n d it m a y b e n e c e s s a r y to use a c o l l a r restraint. S o m e r a b b i t s will e x p e r i e n c e u r e t h r a l s p a s m w h e n t h e cathe t e r is r e m o v e d . Use a n a n a l g e s i c a g e n t for a minimum of 4 hours postcatheterization or until t h e p a t i e n t is u r i n a t i n g n o r m a l l y .
References 1. Jenkins JR, Brown SA: A Practitioners Guide to Rabbits and Ferrets. Lakewood, CO, AAI-L~,1993, p 13 9. Harkness JE, Wagner JE: The Biology and Medicine of Rabbits and Rodents (ed 4). Media, PA, Williams & Wilkins, 1995, pp 16, 118-120 3. Laber-Laird K, Swindle MM, Flecknell P: Handbook of Rodent and Rabbit Medicine. Great Britian, Pergamon, 1996, pp 190, 234-936 4. Bennett RA: Nasogastric intubation for enteral afimentation in rabbits, in 1996 Scientific Proceedings, Tenth Annual North American Veterinary Conference, Orlando, Florida, p 848 5. Hillyer EV: Pet rabbits, in Quesenberry KE, Hillyer EV (ed): The Veterinary Clinics of North America Small Animal Practice Exotic Pet Medicine II, vol 24 (no 1). Philadelphia, PA, Saunders, 1994, pp 36-37 6. Mader DR: Clinical procedures in rabbits, in 1996 Scientific Proceedings, Tenth Annual North American Veterinary Conference, Orlando, Florida, p 863-864 7. Brown SA, Rosenthal KI.: Manson Self Assessment Colour Review of Small Mammals. London UK, Manson Publishing (in press) 8. Bennett RA: Intraosseous catheters in small mammals, in 1996 Scientific Proceedings, Tenth Annual North American Veterinary Conterence, Orlando, Florida, p 847 9. Jenkins JR: Soft Tissue surgery and dental procedures, in Hillyer EV, Quesenberry ICE (eds): Ferrets, Rabbits and Rodents Clinical Management and Surgery. Philadelphia, PA, Saunders, 1996, pp 231-934
Rabbits are very popular house pets and they require the same level of veterinary care as the canine or feline patient. It is often necessary to perform a variety of clinical procedures for both routine preventative care and when attempting to diagnose and treat illness in these patients. Many practitioners are wary of handling rabbits and attempting these procedures due to their fragile nature. However, armed with a good understanding of the rabbit's behavior and physical limitations and with the use of appropriate manual or chemical restraint, the veterinarian can accomplish necessary clinical procedures without harm to the patient.
Copyright 9 1997 by W. B. Saunders Company. Key words: Rabbits, clinical techniques, venipuncture, diagnostics, restraint.
Rabbits as P a t i e n t s he domestic rabbit (Oryctolagus c u n i c u l u s ) aintains the physiology and behavior of a prey species. Even t h o u g h the patient may be a beloved pet and is handled frequently by its owners, it will respond to pain and stress like its wild ancestors. Fear in a rabbit can cause a fall in body temperature, renal ischemia, and an increase in catecholamine production that can interfere with anesthesia. ~ Rabbits that are in pain can b e c o m e anorectic, lethargic, and experience p o o r recovery from anesthesia or surgery. Severely stressed rabbits may die within a few hours despite all efforts to reverse the process. Prolonged stress can lead to gastric ulcers, cardiomyopathy, and alterations in gastrointestinal (GI) flora. 1 Despite this seemingly gloomy outlook, with careful handling, the rabbit patient can survive routine diagnostic and surgical procedures. Factors that increase success include frequent observation of the patient's behavior, gentle handling, quiet recovery and hospitalization areas, analge-
From Midwest Bird & Exotic Animal Hospital, Westchester,IL. Address reprint requests to Susan A. Brown, DVM, Midwest Bird & Exotic Animal Hospital, 1923 S. Mannheim Rd, Westchest~ IL 60154. Copyright 9 1997 by W.. B. Saunde~:sCompany. 1055-937X/97/0602-000255.00/0
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sia, and anesthesia when appropriate. It is preferable, as with most pets, to hospitalize rabbits for as short a time as possible and to return them to the h o m e environment quickly.
Restraint Before attempting to handle a rabbit, make careful observations of its behavior. Rabbits that are fearful will breathe rapidly, stare with wide open unblinking eyes and sit very still, but in a position that allows for a quick escape with the legs tucked up u n d e r the body. Angry rabbits will attempt to attack when approached, making a snorting noise and striking out with their front feet. They may also attempt to bite or strike the g r o u n d forcefully with their hind feet (thumping). Calm rabbits blink their eyes more frequently and have a relaxed appearance to the eye lids. They are curious about their surroundings and are not afraid to investigate. When a relaxed rabbit stops to rest, it stretches its body out as opposed to the tightly h u n c h e d posture of the tense rabbit. When presented with a frightened rabbit, move slowly and talk quietly. Work on the floor with the patient until it is securely restrained. Occasionally a tense rabbit will scream in fear at the slightest touch. This may be in response to an o d o r in the r o o m that is frightening (ie, a previous ferret patient who has a predatory odor) or to an extended period of exposure to alarming noises (ie, a reception area with barking dogs). Rabbits have fragile spines and powerful back legs. If handled improperly, a rabbit can fracture or luxate its lumbar vertebrae or hind limbs. Place carriers or boxes on the floor to prevent potential injuries from j u m p i n g or falling off a table. Remove a rabbit from its carrier or cage backwards to minimize fear by not exposing it immediately to excessive visual stimulation. If the rabbit is very nervous, use a towel to cover its eyes before picking it up. Once the pet is restrained, it can be moved to the examination table. As with other pets, do not have the owner
Seminars in A v i a n and Exotic Pet Medicine, Vol 6, No 2 (April), 1997: pp 86-95
Clinical Techniques in Rabbits
help with the restraint because the owner may be injured by the rabbit's sharp claws. In addition, using an inexperienced owner to help with restraint may increase the chances of injury to the rabbit's spine. Rabbits can be picked up a variety of ways. O n e m e t h o d is to grasp the loose skin over the n a p e of the neck and the shoulders and place the o t h e r hand u n d e r the rabbit's hind quarters and lift. A second m e t h o d is to grasp the thorax ventrally and place the other h a n d u n d e r the hind quarters and lift. A third m e t h o d that is particularly useful for fractious rabbits is to either grasp the thorax ventrally or by the nape of the neck with one h a n d and firmly grasp the lower l u m b a r area with the other h a n d (Fig 1). Grasp the l u m b a r area immediately cranial to the pelvis and completely enclose the lumbar vertebral column and associated musculature with the hand. This area of the spine is thus protected from a hyperextension caused by a powerful kick. Fractious rabbits usually stop trying to kick when held in this manner. If a rabbit has to be transported any distance, tuck its head u n d e r the b e n d o f the elbow and support its body along the forearm. This m e t h o d minimizes visual stimulation and keeps the pet calm. Use a towel, mat, or other material to provide a nonskid surface on the examining table. Rabbits can sustain serious injuries if they try to move quickly on a slippery table. An excellent
Figure 1. Restraint offractious rabbit byscruffofneck and lumbar grasp.
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Figure 2. First fold of the towel wrap restraint in rabbit. m e t h o d of restraint that is used for examination, medication and anesthesia is the towel restraint. Lay o p e n a bath towel on the examination table. Place the rabbit with its front feet at the edge of the center of the long side of the towel. Fold one side o f the towel over the rabbit firmly (Fig. 2), then fold the back towards the front to prevent the rabbit f r o m escaping. T h e most i m p o r t a n t part is the final fold of the towel on the remaining side. T h e final side needs to be pulled very firmly across the rabbit's neck and back enclosing the front feet. W h e n d o n e properly, only the rabbit's head should be showing, with the front feet firmly tucked into the towel and the rest of the body firmly restrained (Fig 3). T h e towel wrap facilitates examination of the eyes, ears, and teeth and can be used for administering oral medications and subcutaneous (SC) injections in the intrascapular area. T h e towel restraint can also be used when administering inhalant anesthesia by mask. Wrap the patient as described, be
Figure 3. Completed towel wrap restraint in rabbit.
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seated, and place the rabbit with its hind quarters in yotar lap and its back supported by your a b d o m e n and chest. The rabbit's head is now in an upright position, which makes it very accessible for placing a mask. Watch the rabbit's breathing pattern through the mask by observing the m o v e m e n t of the nares. A n o t h e r restraint m e t h o d is to place the rabbit on its dorsum to examine the ventrum and trim the nails. Use the lumbar grasp described above to pick the rabbit up and turn it over onto its back. Lay the patient in your lap with its head hanging slightly over your knees. Keep one hand restraining the thorax. Most rabbits will eventually relax into an almost hypnotic state, and it will be possible to complete an examination. Loud noises or pain will cause the rabbit to try to roll over quickly and escape. Trim the nails in this position with an assistant. Have the assistant grasp each forelimb at the level of the h u m e r u s in each h a n d and extend and abduct the legs slightl}: This will prevent the patient from roiling over and escaping. Train owners to restrain their pets this way. Encourage them to sit on the g r o u n d until they are confident of their technique to prevent potential injuries during escape attempts. It is a good idea for the inexperienced practitioner to initially practice this technique on or close to the ground. Rabbits are very sensitive to touch a r o u n d their eyes, ears, nose, and mouth. Make sure the rabbit is securely restrained and d o n ' t prolong the examination of this area. Once the head has been examined, remove the towel restraint and continue with tile examination. Some rabbits feel more comfortable if a section of towel is placed loosely over its head, which may give it a sense that it is hiding. Use mild sedation on rabbits that are so difficult to handle that a physical examination cannot be safely undertaken. There are many options available for sedation in the rabbit using either a single drug or a combination of agents. Diazepam at I to 2 m g / k g intramuscular (IM) or ketamine hydrochloride at 20 m g / k g IM are two possible choices.
be particular challenging, but should be examined on every rabbit because they are frequently sites of disease. An oral exam can be p e r f o r m e d without anesthesia. One option is to use an otoscope with a large cone inserted into the interdental space. The drawback to the otoscope cone is that the view is restricted and over time, the cone becomes damaged from the teeth. The frayed edges can cause abrasions of the gums. Alternately, use either a canine vaginal speculum or a h u m a n nasal speculum with a light attachm e n t (Fig 4). The metal blades are indestructible, and when spread, they allow a more complete view than an otoscope cone. This instrument can also be used when trimming molars in the awake patient. Use gentle technique when palpating the a b d o m e n of the rabbit, particularly tile stomach. Severe bruising and ruptures can occur with overzealous palpation. Use caution when inserting a rectal t h e r m o m e t e r to prevmlt tearing of the delicate rectal mucosa. Rabbit skin is very thin and tears easily. Use care when plucking matted hair particularly a r o u n d the perineum. Tears that occur in this area can be painful and have the potential for becoming infected.
Analgesia As mentioned, the success of treatment can be decreased in rabbits that are severely stressed by pain or fear. The signs of fear have been described in the Restraint section. It has been well d o c u m e n t e d that rabbits experience pain, but it may be difficult to recognize these signs2 ,3 Prey
Examination A t h o r o u g h and complete physical examination should be p e r f o r m e d on the rabbit patient as in other species. The oral cavity and teeth can
Figure 4. Canine vaginal speculum used as oral speculum for examination of rabbit molars.
Clinical Techniques in Rabbits
species such as the rabbit tend to hide discomfort. T h e signs of pain are varied and include one or m o r e of the following: h u n c h e d appearance, tooth grinding, depression, anorexia, unresponsiveness to surroundings or to caretakers, hiding, reluctance to move or a b n o r m a l m o v e m e n t , and unusual aggressive behavior. Rabbits benefit f r o m the use of analgesics postsurgically. For m i n o r surgical procedures, one dose postoperatively may be all that is necessary. Good choices for postoperative analgesia include opioid agonist-antagonists b u p r e n o r phine (0.01 to 0.05 m g / k g intravenous [1V], SC every 8 to 12 hours) or b u t o r p h a n o l tartrate (0.1 to 0.5 m g / k g IV, SC, IM every 4 h o u r s ) 2 ,3 Naloxone (0.04 m g / k g IV, IM, SC) can reverse the sedative and depressant effects o f these drugs should it be necessary. 2 Some short-term sedative effects may be desirable to keep the patient m o r e relaxed in the hospital atmosphere. Nonsteroidal analgesics that can be used include aspirin (100 m g / k g by m o u t h every 48 hours), ibuprofen (10 to 20 m g / k g by m o u t h every 4 hours), and flunixin m e g l u m i n e (1.1 m g / k g SC every 8 hours or by m o u t h every 12 to 24 hours). These drugs are useful for pain associated with dental disease and arthritis. T h e author has used the injectable f o r m of flunixin diluted in a cherry-flavored syrup given orally with good results. The shelf-life of this preparation is at least 2 months j u d g i n g by clinical response. It is unknown if the long-term oral use of flunixin m e g l u m i n e could potentially result in gastric ulceration as in other species. T h e author does not use flunixin m e g l u m i n e for longer than 7 days consecutively without a 1- to 2-day b r e a k in dosing.
Hospitalization Hospitalize rabbits for as short a period as possible and handle only when necessary. Place hospital cages in the quietest area of the hospital. T h e sound of constantly barking dogs is extremely upsetting to the already c o m p r o m i s e d rabbit patient. Some rabbits will not defecate or urinate in their cage unless provided with a litter pan. Do not use clay cat litter material because it can be ingested resulting in a fatal gastrointestinal tract (GIT) impactiou. Use only digestible pelleted bedding. A hide box can help frighte n e d rabbits feel m o r e secure. Do not use r u b b e r
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or plastic mats on the cage floor because there is a potential for ingestion and G I T obstruction. Place debilitated rabbits on a soft, absorbent bedding such as synthetic fleece or thick p a p e r or cloth towels. Provide fresh greens such as parsley, dandelion greens, r o m a i n e lettuce, endive, beet tops, and carrot tops along with good quality grass hay and rabbit pellets. Use heavy crocks or attach food and water bowls to the cage to minimize spillage and d a m p cages. If possible, use the type of water container to which the rabbit is accustomed. Rabbits do not tolerate high humidity or high environmental temperatures. Their o p t i m m n t e m p e r a t u r e range is 40~ to 85~ with the mid to lower 70's preferred. '2 Closely m o n i t o r rabbits maintained in incubators to prevent hyperthermia.
Surgical Considerations This report is not intended to include a comprehensive discussion of surgery or anesthesia. Included in this section are a few special considerations for the rabbit patient. Use a heat source to maintain the rabbit's body t e m p e r a t u r e for major surgical procedures. T h e rabbit's body t e m p e r a t u r e can d r o p precipitously during surgery, especially during abdominal procedures. Use a rectal t h e r m o m e t e r to d e t e r m i n e if the patient needs additional heat during surgery or a heated cage during the postoperative period. It is not necessary to fast rabbits before surgery because they are unable to vomit. T h e exception is GIT surgery, where a 6- to 12-hour fast may be useful to reduce the GIT contents. Due to the nature of the rabbit GIT physiology, it is impossible to completely e m p t y the GIT contents even with fasts longer than 12 hours. Extended fasts are potentially dangerous in the rabbit and should not be attempted. In addition, fasting may p r o l o n g the a m o u n t of time it takes for the patient to resume eating postsurgically. Elevate the rabbit's h e a d and thorax when u n d e r anesthesia to minimize the pressure on the small thoracic cavity, particularly in overweight animals. Postsurgical abdominal adhesions are comm o n in the rabbit. To mir, imize this problem, wash the powder off surgic)d gloves, handle the tissues as gently and as little as possible, and use synthetic absorbable suture material (as o p p o s e d
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to chromic gut). Postsurgically use verapamil (200 m c g / k g intraperitoneal [IP] every 8 hours for nine doses) to prevent adhesions if there is extensive tissue handling. 2 Verapamil (Solo Pak Laboratories, Inc, Elk Grove Village, IL) is not necessary for uncomplicated routine procedures such as ovariohysterectomies and castrations. Rabbits can remove skin sutures with their incisors. Most rabbits will tolerate skin staples, but they are difficult to place in areas of the body where the skin is thin, such as the a b d o m e n . In addition, rabbits can occasionally pull out staples and then ingest t h e m resulting in a potential for the ingested staples to lodge in the G I T or the oral cavity. Use m o n o f i l a m e n t synthetic 4-0 suture material in a subcuticular pattern to close the skin. Subcuticular sutures are inaccessible to the patient. In addition, apply a small a m o u n t of surgical glue along the incision to bring the skin edges into apposition. Minimize the use of restraint apparatuses in the rabbit such as Elizabethan collars. They are not well tolerated and they may cause the patient to b e c o m e anorectic and depressed. In addition, the rabbit may not be able to access its nutritionally i m p o r t a n t cecotropes. Use several layers of bandaging to protect catheters or splints and reapply as needed. Use large tags of adhesive tape attached to the bandage to attract the rabbit to chew the tags instead of the bandage. Rabbits usually ignore bitter-tasting substances applied to areas where chewing is to be discouraged.
Medication Techniques Oral Medication Administering medications to a rabbit patient can be challenging. Rabbits will accept oral preparations if they are in liquid form, sweetened, and of a small volume. Use the towel restraint and place the rabbit in your lap with the head slightly elevated. Place the syringe tip into the interdental space between the incisors and premolars and give the medication slowly. Some rabbits learn to take their medication o f f a spoon when mixed with substances such as fruit j a m (strawberry is a favorite), p e a n u t butter, cherry-, orange-, or banana-flavored syrups, fruit juice, molasses, m a s h e d banana, blenderized greens, peanuts (use only two or three dry roasted unsalted, not raw), or dried fruit (raisins, apricots). Be patient when syringe feeding large vol-
umes of material by syringe. Do not a t t e m p t to syringe feed a severely depressed or debilitated rabbit. Use a nasoesophageal (NE) tube for these cases and for rabbits that are either too fractious to handle or those that refuse to take the syringe feeding. T h e r e are m a n y successful syringe feeding formulas for rabbits. For shortt e r m feeding, use canned p u m p k i n (not p u m p kin pie filling) which is high in fiber and easy to administer. A n o t h e r possibility for m o r e longterm feeding consists of one part g r o u n d rabbit pellets or alfalfa powder (found in health food stores) to one part blenderized fresh green leafy vegetables or j a r r e d strained vegetable baby food. Add to this mixture e n o u g h oral electrolyte solution to make a cream soup consistency. Administer at approximately 30 m L p e r 2 kg body weight every 8 to 12 hours. Use a syringe with a catheter tip to avoid the n e e d to strain the mixture. Administer pills to rabbits by crushing t h e m and mixing them with one of the palatable substances described above. Alternatively, give the pill directly to the patient by using the index finger to push it through the interdental space and as far back into the m o u t h as possible.
NE Feeding Tube A NE tube is invaluable in rabbit patients that are either too debilitated or fractious to take oral medications or feedings. ~ Rabbits tolerate NE tubes well and they can be left in place for several weeks if there are no complications (Figs 5 and 6). Although complications of this p r o c e d u r e are u n c o m m o n , they include rhinitis, epistaxis, dacryocystitis and esophageal reflux. 4 Use a 5 to 8 French feeding tube. The clear h u m a n pediatric tubes have the advantage of being radiopaque, allowing visualization of the contents of the tube and having an attached cap on the end to prevent tube contamination. Make additional holes in the caudal aspect of the tube before placement. General anesthesia is not necessary to place the tube. Use the towel restraint and elevate the rabbit's head. Instill two or three drops of an ophthalmic anesthetic or diluted lidocaine into the nostril. 4,5 Alternatively, lubricate the tube with lidocaine jelly before passage. Mlow a few minutes for the anesthetic to take effect. Measure the tube length f r o m the nares to the caudal aspect of the s t e r n u m a n d m a r k with tape or marking pen. Lubricate the tube and place it in the ventral meatus directing
Clinical Techniques in Rabbits
Figure 5. Nasoesophageal tube in rabbit (frontal view). it medially and ventrally. 4 If the nasal cavity was adequately anesthetized, any discomfort will be minimal and transient. The tube should move without resistance. Put the rabbit's head in a normal flexed position as the pharynx is approached to encourage tube passage into the esophagus. Always assess p r o p e r placement by radiograph. The distal end of the tube can be located in the esophagus or the stomach. Sterile saline introduced through the tube may not elicit a cough response when the placement is in the trachea. Once the placement is assessed, suture the tube in place over the bridge of the nose and at the top of the head between the ears with tape butterflies. Rabbits rarely need any form of restraint to prevent the removal of the tube. Pass the tube-feeding formula through a fine
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strainer to prevent blockage of the tube. Flush the tube with saline or water before and after each feeding or medication. One possible formula that can be used for long-term maintenance of debilitated rabbits with NE tubes is two parts alfalfa powder (available in health food stores, high in fiber, and finer in consistency than ground rabbit pellets) plus one part blenderized green leafy vegetables or j a r r e d strained vegetable baby food plus one part Deliver 2.0 (Mead Johnson, Evansville, IN) plus e n o u g h oral electrolyte solution to make the formula into a thin consistency. Strain the mixture t h r o u g h a fine strainer. Use 30 mL per 2 kg body weight every 6 to 12 hours. Subsequent stools are formed, but small in size. Consider vitamin supplementation for patients that are debilitated for longer than a week a n d / o r are unable to p r o d u c e or eat their cecotropes. Rabbits can still eat normally with an NE tube in place. Some practitioners have had success with the use of pharyngostomy tubes in rabbits. The advantages over the NE tube is that a larger tube can be used and a thicker consistency of material can be administered. The disadvantage is that it requires a surgical procedure to place the tube. The technique for pharyngostomy tube placem e n t would be similar to the canine or feline patient.
Injectable Medications Use small gauge needles (22 to 27 gauge [G]) for most injections to minimize discomfort. Give SC injections and fluids in the dorsum of the neck or shoulder area. Minimal restraint is necessary for this procedure. Administer IM injections in the dorsal lumbar muscles cranial to the pelvis or in the cranial aspect of the quadriceps. Give IV injections in the lateral saphenous or cephalic veins. The jugular and femoral veins can also be used. Although many sources advocate the use of the marginal auricular vein or central auricular artery for injections, the practitioner should be cautious using these sites. Sloughing of the ear tip can occur. 6 This is most often due to the infusion of an irritating substance or damage to the blood vessel. These vessels should be used as a last resort for 1V injections.
Intravenous Catheters Figure 6. Nasoesophageal tube in rabbit (side view).
Use small gauge (22 to 25 G) over-the-needle catheters. Butterfly catheters do not work well
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because of their tendency to puncture the delicate wall of the vein with the slightest movement. Pluck or clip the fur over the vein and aseptically prepare the skin. Rabbit skin is thin and friable; therefore, it is unnecessary to incise the skin before placing an IV catheter. Use sufficient bandaging to prevent the removal of the catheter in the awake animal. Use IV catheters to administer warm fluids during any extensive surgical procedure. A syringe p u m p is invaluable in this small species to administer fluids at a prescribed rate and volume. Although many sources advocate the use of the marginal auricular vein or central auricular artery for IV catheterization, the practitioner is cautioned against its routine use for reasons described u n d e r Injectable Medications. The lateral saphenous vein affords a very accessible site for an indwelling catheter, particularly for use during surgery. This vein runs medial to lateral across the cranial aspect of the tarsus and has two branches that continue laterally up the leg. Place the catheter so that the injection cap will be above the plantar surface of the foot when the rabbit is sitting in a normal position. It is difficult to maintain a catheter in the lateral saphenous for extended periods in the awake animal due to the constant m o v e m e n t of the hind legs. Cephalic catheters are placed and taped in as in other species and can be maintained for extended period. Jugular catheters can be used, but sedation will be necessary in most patients for placement. Fluid administration guidelines in the rabbit are similar to the cat and dog. Maintenance fluid level for the rabbit is approximately 80 to 100 m L / k g every 24 hours. 5,7 Subcutaneous fluids are more often used due to the ease of administration. Rabbits have a large subcuticular space u n d e r the shoulders and neck. However, rabbits that are debilitated, severely stressed, or undergoing major surgical procedures should receive IV or intraosseous (IO) fluids to improve chances for survival. Subcutaneous fluids may be poorly absorbed in these cases.
Intraosseous Catheters Intraosseous catheters are invaluable in the debilitated rabbit where venous pressure is low or in the patient where IV catheterization is unsuccessful. Rabbits seem to tolerate IO cath-
eters better than IV catheters. Place IO catheters in the femur, humerus, or tibia, s The femur is most frequently used, and a catheter in this bone causes the least disruption to normal movement. Use a spinal needle (20 to 22 G 1.5 to 3 inch), if possible, due to the sturdy construction of the needle and the stylet, which prevents occlusion of the needle with bone. Alternatively, a 20- to 22G hypodermic needle can be used with a small gauge sterile surgical wire inserted in the needle bore as a stylet. Use anesthesia in all but the very docile or debilitated patient to place an IO catheter because this procedure can be painful. Use a local anesthetic in the awake patient. Clip the hair and surgically prepare the area over the head of the femur. Use sterile technique and make a small incision over the proximal aspect of the greater trochanter. Insert tile needle through the trochanteric fossa using firm pressure and gentle rotation. When the catheter has entered the marrow cavity, it should advance easily until the hub meets the skin. Remove the stylet and test patency and placement by slowly injecting 3 to 5 mL of sterile saline. If the needle is properly placed, there is little resistance to the injection, and the soft tissue a r o u n d the area does not swell. Place butterfly tape a r o u n d the needle hub and suture it to the skin. Place an injection cap on the needle and flush with heparinized saline/ Any substance that can be injected intravenously can be given intraosseously with the exception of some chemotherapy agents. The bone is not distensible; therefore, do not give fluids as a bolus. Administer medications slowly. A syringe p u m p is invaluable for slow, measured infusion into the IO catheter. IO catheters can be left in for several days. It is advisable to use systemic antibiotics while the catheter is in place and for 2 or 3 days after removal.
Blood Collection The blood volume of the rabbit is approximately 6% of the body weight. 9 Therefore, a 3,000-gm (approximately 6 pounds) rabbit has a blood volume of 180 mL. Up to 10% of this total can be collected safely in one sampling (18 mL). Up to 25% can be collected over a 2-week period. 2 For routine complete blood cell (CBC) counts and serum biochemistries, it is only necessary to obtain one half to 3 mL of blood.
Clinical Techniques in Rabbits
Figure 7. Venipuncture in rabbit using central auricular artery. Use small gauge needles (29 to 95 G) to obtain blood samples in rabbits. Do not use a toe nail clip for routine sampling. This procedure is painful, may result in infection, and may yield p o o r samples due to contamination with tissue fluids. The most accessible site to get a sample (1 to 3 mL) is the central auricular artery (Fig 7). Restrain the rabbit and apply alcohol to the ear to wet the fur. Flick the ear gently with a finger to cause the artery to become more prominent. If the rabbit is cold or frightened, warm the ear to raise the artery. Insert a 25-G needle or butterfly catheter into the artery without a syringe attached and allow the blood to flow into the collection container. In large rabbits, a vacutainer may be used, but in smaller rabbits, the suction will collapse the vessel. Once the needle is removed, apply pressure to the venipuncture site for several minutes or apply a temporary bandage to prevent h e m a t o m a formation. Blood collection from the marginal ear vein is difficult and often unsuccessful, especially in smaller breeds. The lateral saphenous vein, as previously described, is easy to access even in the awake rabbit and is of a sufficiently large size and length to be highly visible. Restrain the rabbit by scruffing it by the nape of the neck and hold it on its side using the other hand to grasp firmly a r o u n d the dorsal aspect of the quadriceps of the leg to be used for venipuncture. This occludes the vein, which allows better visualization. Gently pluck or clip the hair over a portion of the vein and apply alcohol to improve visibility. Use a 25-G needle attached to a syringe to draw the sample. Use firm pressure for at least a minute afterwards or a temporary bandage for hemostasis.
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The cephalic vein can be used, but it is very thin and fragile, particularly in smaller species. The restraint and collection m e t h o d is similar to the dog. Towel restraint is useful here. The jugular vein can be accessed in the awake animal in either sternal or dorsal recumbency. For sternal recumbency, hold the rabbit near the edge of a table with the head extended upwards and the feet extended downwards over the edge. 5 Use alcohol to help visualize the vein, which runs laterally from the thoracic inlet to the base of the ear. Place digital pressure just lateral to the thoracic inlet and visualize or palpate this superficial vein. Use a 92-G needle attached to a syringe to draw the blood. Alternatively, restrain the rabbit in a towel wrap and place it in dorsal recumbency with the neck extended. 3 Visualize or palpate the jugular vein and draw the blood as described in the previous m e t h o d of restraint. Cardiac and vena cava puncture have been described for the rabbit. It is the author's opinion that these methods have inherent risks, particularly in the hands of inexperienced phlebotomists and, therefore, are not appropriate use in for pet rabbits.
Bone Marrow Biopsy Bone marrow biopsies may be taken from a variety of sites including the long bones and pelvic bones, and the technique is similar to the cat or dog. Access the marrow cavity of the femur by using the technique previously described u n d e r Intraosseous Catheters. Attach a lO-mL syringe to the hub of the inserted needle and apply vigorous suction to retrieve bone marrow.
Cerebrospinal Fluid Collection Collect cerebrospinal fluid (CSF) by anesthetizing the rabbit, place it in lateral recumbency, and flex the head toward the sternum. Clip the hair and surgically prepare the area from the occipital protuberance to the third cervical vertebrae and laterally at least 3 cm on each side of the midline. Use the cranial margins of the wings of the atlas and the occipital protuberance as the landmarks for needle placement. Use a 22-G 1.5 inch spinal needle and place it midway between these points and direct it towards the nares. 6 Remove the styler after the dura is penetrated
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and look for spinal fluid to a p p e a r in the hub. Draw the sample and remove the needle.
Lacrimal Duct Cannulation A c o m m o n disorder of rabbits is dacryocystitis, which presents with unilateral or bilateral e p i p h o r a and facial dermatitis. Most often, this condition is treated by cannulating and flushing the lacrimal duct. This removes potential obstructive p u r u l e n t material, allows culturing of the flushed material, and improves the penetration of medication into the area. A variety of lacrimal cannulas may be used for this procedure, and either straight or curved models are acceptable. Accomplish cannulation of the lacrimal duct without a general anesthetic. Restrain the rabbit by the towel m e t h o d and apply an ophthalmic anesthetic to desensitize the affected eye. T h e r e is only one nasolacrimal canaliculus and puncturn in the rabbit located in the lower lid. Visualize the puncture by either pulling the lower lid down and rolling it outward or grasping the edge of the lower lid and pulling it directly laterally. Rabbits rarely object to this procedure. Cannulation is m o r e difficult in the presence of a persistent discharge and severe inflammation of the peripunctal area. Apply pressure below the p u n c t u r e to cause the lips of the p u n c t u m to pout, allowing better access to the duct. 7 Use 1 to 2 m L of sterile saline to gently flush the duct once the cannula is in place. Do not use e x t r e m e force to flush the duct or it may rupture. T h e saline should exit the nares if cannulation was successful. If the duct is blocked or the cannula is not placed properly, the saline will flush retrograde out the puncture. Try placing the cannula again should this occur. If the duct is ruptured, the saline will leak out into the s u r r o u n d i n g tissues, often into the retrobulbar space causing the eye to bulge laterally. If this happens, stop flushing immediately. T h e saline will be absorbed over the next 24 hours, and the eye will return to normal. If the lacrimal duct is ruptured, the healing process may r e n d e r it permanently strictured due to scar tissue formation.
Microbiological Culture Collection Techniques for obtaining microbiological cultures in the rabbit are similar to those in the dog and cat. Howevm, three areas bear special men-
tion because they are c o m m o n sites of disease in the rabbit. Rabbits have a predisposition to develop large abscesses filled with thick tenacious p u r u l e n t material. Practitioners often make the mistake of using a glob of p u r u l e n t material as their culture sample. This practice usually leads to a negative result. This material is c o m p o s e d of dead cells and phagocytized bacteria and is not a viable choice for sampling. Collect the culture by swabbing the wall of the abscess or draining tracts after the p u r u l e n t material has b e e n removed. Negative results are still possible, but the probability of a positive culture is increased with this method. Rabbits are p r o n e to u p p e r respiratory infections caused by a variety of infectious agents including Pasteurella multocida, Staphylococcus aureus, and Bordetella bronchiseptica. A d e e p nasal culture and sensitivity is an extremely useful tool for selecting effective antibiotic therapy. T h e culture is obtained quickly and with minimal discomfort in the awake rabbit. Seat the handler with the towel-restrained rabbit's hind quarters in the handler's lap and the rabbit's back supp o r t e d by the handler's a b d o m e n and chest. T h e nares are readily visible, and the rabbit is securely restrained in this position. Use a sterile calgi swab or minitip culturette and direct it medial and caudal into the nasal sinus as far as it will go then remove it immediately. T h e rabbit may j u m p slightly as the swab enters the sinus. A small a m o u n t of blood will be present on the swab. Rarely, a rabbit will exhibit mild epistaxis for a few minutes after sampling and will sneeze. Keep the rabbit wrapped in the towel, but allow it to return to its n o r m a l sitting position until the bleeding has stopped. Advise the client that pink-tinged fluid may be seen periodically at the nares for the next h o u r or two. As m e n t i o n e d in the section on Lacrimal Duct Cannulation, dacryocystitis is a c o m m o n problena in rabbits. This disease can be caused by the same infectious agents that cause u p p e r respiratory disease. The material flushed f r o m the lacrimal duct is easily cultured. Have an assistant hold a sterile swab just ventral to the nares without touching the skin. Allow the flushed material exiting the nasal sinus to drop onto the swab. One or two drops is sufficient. Obviously, do not use an antibiotic solution to flush the lacrimal ducts when obtaining a culture.
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Clinical Techniques in Rabbits
Malocclusion and Associated Disease As s t a t e d in t h e E x a m i n a t i o n section, t h e oral cavity a n d t e e t h a r e f r e q u e n t l y a site o f disease in the rabbit and should be thoroughly examined. H o w e v e L t h e variety a n d c o m p l e x i t y o f d e n t a l disease p r e s e n t a t i o n s , i n c l u d i n g m a l o c c l u s i o n a n d a s s o c i a t e d infections, p r e c l u d e s a t h o r o u g h d i s c u s s i o n o f d i a g n o s t i c a n d t r e a t m e n t techn i q u e s in this article. T h e r e a d e r is e n c o u r a g e d to f u r t h e r r e s e a r c h t h e s e s y n d r o m e s )
Urine Collection U r i n e can b e c o l l e c t e d in t h e r a b b i t by f r e e c a t c h i n t o a n e m p t y litter b o x , by g e n t l e e x p r e s sion o f t h e b l a d d e r in t h e awake o r a n e s t h e t i z e d p a t i e n t , by cystocentesis, o r by u r i n a r y c a t h e t e r ization. E x p r e s s t h e b l a d d e r in t h e awake a n i m a l in t h e s a m e m a n n e r as in t h e d o g o r cat. U r i n e will b e v o i d e d caudally. T a k e c a r e n o t to use excessive f o r c e o r t h e b l a d d e r can b e severely b r u i s e d o r ruptured. Cystocentesis c a n usually b e p e r f o r m e d in t h e awake a n i m a l . However, if t h e p a t i e n t is f r a c t i o u s o r e x t r e m e l y stressed, use s e d a t i o n o r a n e s t h e s i a to p r e v e n t a p o t e n t i a l l a c e r a t i o n o f t h e b l a d d e r wall. R e s t r a i n t h e awake r a b b i t by h o l d i n g t h e scruff at t h e n a p e o f t h e n e c k in o n e h a n d a n d b o t h r e a r legs in t h e o t h e r h a n d . S t r e t c h t h e r a b b i t o u t a n d p l a c e in d o r s a l r e c u m b e n c y . 6 Clip a small a r e a o f h a i r a n d p e r f o r m a sterile p r e p a r a t i o n o f tile a n t e p u b i c r e g i o n . Use a small g a u g e n e e d l e (23 to 25 G) a t t a c h e d to a n a p p r o p r i a t e - s i z e d syringe to p e n e t r a t e t h e b l a d d e r wall a n d o b t a i n t h e s a m p l e . T h e m o s t c o m m o n use for u r i n a r y c a t h e t e r i z a tion is to e n a b l e f l u s h i n g o f t h e b l a d d e r to r e m o v e excessive c a l c i u m c a r b o n a t e p r e c i p i t a t e , o f t e n r e f e r r e d to as " s l u d g e . " A n i n d w e l l i n g u r e t h r a l c a t h e t e r m a y also b e u s e d to allow h e a l i n g o f injuries to t h e u r e t h r a c a u s e d by b i t e w o u n d s o r u r e t h r a l calculi. I n a d d i t i o n , u r i n a r y c a t h e t e r i z a t i o n is essential to p e r f o r m single a n d d o u b l e c o n t r a s t studies o f t h e b l a d d e r . Use a f l e x i b l e 3.5 to 6 F r e n c h u r i n a r y c a t h e t e r o r f e e d i n g tube. Use a stylet o f wire, if necessary, o r stiffen t h e c a t h e t e r by p l a c i n g it in t h e f r e e z e r
for a s h o r t p e r i o d . D o n o t use t o m c a t c a t h e t e r s because the likelihood of injuring the delicate u r e t h r a l wall is i n c r e a s e d . A n e s t h e t i z e o r s e d a t e t h e r a b b i t a n d p l a c e it in d o r s a l r e c u m b e n c y . E x t e n d t h e p e n i s in t h e m a l e r a b b i t by a p p l y i n g t e n s i o n a n d p r e s s u r e c r a n i a l l y at its base. Use a small vaginal s p e c u l u m o r o t o s c o p e to visualize t h e u r e t h r a l o p e n i n g in t h e f e m a l e . L u b r i c a t e t h e c a t h e t e r a n d pass it i n t o t h e b l a d d e r . Use w a r m e d saline to flush t h e b l a d d e r , u s i n g small a m o u n t s frequently. A d m i n i s t e r SC o r IV fluids to e n c o u r a g e u r i n a t i o n . If t h e c a t h e t e r is to b e left in place, s u t u r e it by m e a n s o f b u t t e r f l y t a p e to t h e p e r i n e u m . R a b b i t s will c h e w o u t c a t h e t e r s unless t h e y a r e severely d e b i l i t a t e d , a n d it m a y b e n e c e s s a r y to use a c o l l a r restraint. S o m e r a b b i t s will e x p e r i e n c e u r e t h r a l s p a s m w h e n t h e cathe t e r is r e m o v e d . Use a n a n a l g e s i c a g e n t for a minimum of 4 hours postcatheterization or until t h e p a t i e n t is u r i n a t i n g n o r m a l l y .
References 1. Jenkins JR, Brown SA: A Practitioners Guide to Rabbits and Ferrets. Lakewood, CO, AAI-L~,1993, p 13 9. Harkness JE, Wagner JE: The Biology and Medicine of Rabbits and Rodents (ed 4). Media, PA, Williams & Wilkins, 1995, pp 16, 118-120 3. Laber-Laird K, Swindle MM, Flecknell P: Handbook of Rodent and Rabbit Medicine. Great Britian, Pergamon, 1996, pp 190, 234-936 4. Bennett RA: Nasogastric intubation for enteral afimentation in rabbits, in 1996 Scientific Proceedings, Tenth Annual North American Veterinary Conference, Orlando, Florida, p 848 5. Hillyer EV: Pet rabbits, in Quesenberry KE, Hillyer EV (ed): The Veterinary Clinics of North America Small Animal Practice Exotic Pet Medicine II, vol 24 (no 1). Philadelphia, PA, Saunders, 1994, pp 36-37 6. Mader DR: Clinical procedures in rabbits, in 1996 Scientific Proceedings, Tenth Annual North American Veterinary Conference, Orlando, Florida, p 863-864 7. Brown SA, Rosenthal KI.: Manson Self Assessment Colour Review of Small Mammals. London UK, Manson Publishing (in press) 8. Bennett RA: Intraosseous catheters in small mammals, in 1996 Scientific Proceedings, Tenth Annual North American Veterinary Conterence, Orlando, Florida, p 847 9. Jenkins JR: Soft Tissue surgery and dental procedures, in Hillyer EV, Quesenberry ICE (eds): Ferrets, Rabbits and Rodents Clinical Management and Surgery. Philadelphia, PA, Saunders, 1996, pp 231-934