Chapter 177 Pet Rodents
Many small rodents are commonly kept for companionship and enjoyment. This chapter provides information needed to diagnose and treat the most frequently encountered problems of mice, rats, gerbils, hamsters, guinea pigs, and chinchillas.
HUSBANDRY
Caging and Sanitation
• Cages should be made of stainless steel, hard plastic, or glass. These materials are cleaned and sanitized easily and are resistant to gnawing or corrosion from urine and fecal matter. Minimum floor space and height requirements are listed for each species in Table 177-1. With the exception of guinea pigs, all cages need secure lids.
• Guinea pigs can be housed in open-topped enclosures with walls higher than 10 inches. Ensure that dogs, cats, wild animals, and small children do not have unsupervised access to these cages.
• Clean cages as needed, usually 1 to 3 times per week for most rodents. A scrub brush, dish soap, and water work well. If cages are not kept clean, ammonia, other irritants, moisture, and bacteria concentrations rise to harmful levels, predisposing animals to disease.
• Disinfect the cage twice a month with 1 part sodium hypochlorite (household bleach) mixed in 30 parts water. Let the bleach solution stand for at least 15 minutes. Rinse the cage well afterward.
• All solid-floored cages need to be covered in bedding. Shredded paper, non-resinous wood shavings, wood wool, and corn cobs are all acceptable. Provide at least 2 inches of bedding. Most rodents enjoy burrowing in deeper bedding when it is provided in one corner of a cage. Do not, however, fill the entire cage with deeper bedding. This usually leads to poor sanitation as a result of owners’ failure to recognize buildup of hidden wastes such as moisture from leaking water bottles, cached foods, urine, and feces.
• Wire mesh floors can be used successfully only if the dimension of the mesh is correct. Size the openings to be just large enough for an adult to retract a tarsal joint back through the mesh. Larger holes make it difficult for the animals to walk and cause pressure sores. Smaller openings may cause injuries such as tibial fractures and self-mutilation while struggling to free trapped appendages. Bedding above the wire keeps waste from dropping through the wire and therefore is not recommended. Wire bottom cages do not work well for breeding animals because neonatal rodents must be surrounded by nesting material to maintain moisture in the nest and prevent dehydration. Young rodents often cannot walk correctly on mesh sized for adult feet.
• All pet rodents require visual security. Tubes, jars, or cans made of nontoxic, nonabrasive substances work well for this purpose. Also provide objects for gnawing. Rodents possess open-rooted teeth, and constant wear is necessary to maintain normal dentition. Mice, rats, gerbils, and hamsters enjoy and benefit from exercise wheels.
• A good room temperature range for most pocket pets is 70°F to 75°F. Keep rodents with disease at 85°F to 90°F unless hyperthermia is of concern (some chinchillas).
• Provide 10 to 12 hours of darkness to 12 to 14 hours of light. This light cycle is essential if breeding is desired.
• Hamsters, guinea pigs, and chinchillas that are exposed to temperatures below 65°F may hibernate for a few days or until the ambient temperature rises. Heart rates may be less than 5 bpm during hibernation.
Nutrition
Feed pet rodents laboratory animal chow appropriate for their species (Table 177-2). Seed diets are deficient in protein and contain excessive fat.
• Seeds, as well as vegetables and other foods, may be fed as treats but not to provide more than 15% of calories. Intermittent exposure to vegetables and seeds causes mild, transient diarrhea.
• Adult chinchillas that are not obese should be fed high quality, fresh grass hay ad libitum. Obese animals may need to have the hay rationed. Chinchillas require ⅛ to ¼-cup of fresh pellets per animal each day. Feeding pellets free choice leads to obesity, and the high protein and calcium levels in these diets may predispose animals to urinary tract disease. Most pellets also do not provide sufficient fiber to maintain normal gastrointestinal (GI) motility.
• Feed all diets within 90 days of milling to ensure the highest nutritional value. Encourage owners to check dates on packages and ask pet store managers about providing dates on bulk items.
• If possible, feed pet rodents except guinea pigs from overhead racks. These devices reduce wastage and eliminate fecal contamination of food. Covered hoppers, heavy crocks, or stainless steel bowls that are attached to the side of the cage to eliminate spillage are acceptable and recommended for guinea pigs.
• Feed breeding females and their litters from the floor of their cages until the young are large enough to reach overhead feeders or crawl in and out of crocks.
• Cannibalism of neonates commonly occurs as the result of stress associated with cage cleaning. To minimize cannibalism, clean the cage and provide a 10- to 13-day food supply 1 to 2 days before parturition.
Water
• Do not provide water in open crocks. These are contaminated or spilled easily and are a common cause of dehydration and poor sanitation.
• Sipper tubes and water bottles work well. Clean with dish soap and water daily and disinfect them weekly. Guinea pigs expel food from their mouths into their sipper tubes, so more frequent cleaning is needed.
HISTORY AND PHYSICAL EXAMINATION
A systematic history and physical examination are mandatory. Many disease syndromes are caused by poor husbandry. Pets that have been kept isolated from other rodents or acquired from a private breeder are less likely to harbor infectious disease than animals obtained from a pet store, laboratory, or wholesaler. See Table 177-3 for normal physiologic data.
Obtain the following information:
• Environment, caging (ask owner not to clean cage before coming to hospital), cleaning, temperature, humidity, photoperiod, person responsible for care
• Diet: brand, dating on packages, storage, supplements, type of feeder and waterer and method and frequency of cleaning, feeding schedule, person responsible for feeding
• Dates of unusual activity: breeding, parturition (Table 177-4), cannibalism, abnormal urination or defecation
Examination of Patient and Environment
• Become familiar with normal behavior, locomotion, haircoat, and stools of each of the pet rodent species.
• Observe the pet in its cage for mentation, activity, locomotion, dyspnea, head posture, haircoat, and any grossly observable abnormalities.
• Note respiratory and heart rates before restraint when possible. Observe the condition of cagemates.
• Handle such animals as little as possible. Initially, treat severely ill animals symptomatically, then perform a complete physical examination after the pet’s condition has stabilized.
Evaluate the Cage
• Note the type of diet and bedding as well as the level of sanitation and compare these with what was described in the history.
• Observe quantity and quality of feces and urine. Diarrhea, soft stools, absence of stools, copious urine, and discolored urine all can be signs of illness.
Restraint
• Restraint of pet rodents is easy with experience. Pets that have been handled frequently and gently by the owners require only minimal restraint. Gentle pressure directs the animal as needed. Grasp less cooperative patients (except chinchillas and guinea pigs) by the scruff over the back of the neck with thumb and forefinger (Fig. 177-1). Take care to pinch enough skin to prevent the animal from turning around, yet leave enough slack for respiration. On smaller specimens, hold the base of the tail, if present, between the fourth and fifth fingers to provide additional restraint.
• Hold docile guinea pigs with the palm of one hand supporting the chest while the other hand supports the hind quarters (Fig. 177-2). Place the thumb and forefinger of the first hand in the axillas for additional control.
• Take care to minimize damage to the fur when handling chinchillas because they lose hair easily. Grasp the animal by the tail and scoop it up into the palm of the same hand (Fig. 177-3). If necessary, grasp the thorax just behind the axillas.
• Calm uncooperative rodents by placing an appropriately sized towel over the head. Complete the physical examination by wrapping the patient in a towel and exposing only needed areas. Even oral, ophthalmic, and aural examinations can be performed with minimal effort if the animal is given the chance to relax in the towel “burrow.”
• Remove particularly aggressive patients from their cages by scooping them up in a can or bucket; then slide them out onto a slick surface and pick them up or transfer them to a holding area or scale.
Physical Examination
• Check ears for discharge, foreign bodies, and mites. Bluish discoloration of the ears is a sign of cyanosis. Bright red injected coloration is associated with septicemia. Sores behind the ears and on the neck are often a sequela of aural disease.
• Evaluate submandibular, axillary, inguinal, and popliteal lymph nodes for size and consistency. Enlargements usually indicate infectious or neoplastic disease.
• Reevaluate respirations and heart rate after the stress of handling and compare them with the resting rate noted when the animal was in the cage. Note dyspnea or respiratory sounds. Auscultate animals weighing more than 200 g. Counting every third or fourth beat and multiplying by the appropriate factor allows recording of heart rates up to 500 bpm.
• Palpate the abdomen. Pay special attention to differentiating pregnancy from the bladder, kidneys, abdominal masses, enlarged cecum, and fecal balls in the colon. While palpating the abdomen, examine the mammary chain of all female rodents for signs of mastitis, lactation, or neoplasia. Also check male mice and rats for mammary neoplasia. Mammary tissue extends from the base of the neck to the base of the tail. Gerbils typically have an elliptical sebaceous gland on their ventral midline. Do not confuse this with neoplasia or infection. Check the rectum and perineal area for signs of diarrhea, prolapse, irritation, parasites, and bite wounds. Note that coprophagy is normal in rodents.
• Evaluate the urogenital tract for signs of inflammation, foreign bodies, urine scalding, and vaginal discharge. Locate and palpate the testicles in males. The easiest method of determining sex in pet rodents is to compare the anogenital distance, which is twice as long in males as in females. Other characteristics that allow the determination of sex are as follows:
• Visualization or palpation of testicles or extrusion of the penis from the prepuce indicates a male.
• Examine the skin and fur for conditions such as crusts, alopecia, masses, herniations, and wounds.
• Check tail and feet for swellings, coloration, sores, length of toenails, and condition of footpads.
DIAGNOSTIC Tests
Blood Samples
• Use lateral or medial saphenous veins to obtain samples in animals heavier than 100 g. Liberally clip the area to allow exposure of the vessel before attempting venipuncture. Place a 25- to 27-gauge needle in the vein and collect blood into microtainers or hematocrit tubes as it drips from the hub of the needle (Fig. 177-4). Take extreme care not to collapse and lacerate the vein with overzealous aspiration if a syringe is attached to the needle.
• An alternative technique that is useful in smaller animals is to coat the skin over the vein with a thin layer of petroleum jelly and then to puncture the vessel. Blood is collected with a hematocrit tube as it exits the wound. Samples up to 1% of the animal’s weight are considered safe, even in stressed animals.
To bleed the tail, warm the tail with water or compresses to dilate the tail vessels. In large rats, perform venipuncture with a needle and obtain blood in the usual fashion. In smaller animals, lacerate the tip of the tail. Blood from the wound is collected as described previously. See Tables 177-5 and 177-6 for hematology and chemistry values.
ROUTINE PROCEDURES
Oral Medications: Nutritional Support
Incorporate oral medications into a treat, or administer them in liquid form. If the medication is palatable, administer it by placing the tip of a dosing syringe into the diastema.
• Metal feeding needles, red rubber urinary catheters, or infant feeding tubes work well. Selection is based on the size of the animal and individual preference. Metal feeding needles with ball tips frequently are used in patients weighing less than 100 g (Fig. 177-5). The metal provides the necessary stiffness to pass a tube of small diameter. The ball at the end of the needle makes it difficult (but not impossible) to pass the tube into the trachea. These tubes have the potential to create esophageal tears with improper restraint or when excessive force is applied.
• Measure the length from the tip of the nose to the last rib. Ventroflex the head slightly, and place the tip of the tube through the diastema and over the tongue. If the tube does not pass easily down the esophagus to the premeasured distance, check the tube size and/or reposition the tube before attempting further advancement. The needle is easily palpable percutaneously if it is placed correctly. It is usually safe to administer up to 3 ml/100 g body weight.
• A flexible catheter is ideal for use as a stomach tube in larger rodents (Fig. 177-6). Use a speculum to prevent chewing on the tube. An otoscope head, avian speculum, or piece of wood or plastic with a hole drilled in the center works well. Measure and mark the tube for the distance from the tip of the nose to the last rib. Place the speculum in the mouth and over the tongue. Pass the tube while holding the speculum in place and slightly ventroflexing the head. Resistance is encountered if the tube is malpositioned or is an inappropriate size. The tube must pass over the tongue before it can be advanced down the esophagus. This is difficult in some animals. Palpate the throat to confirm the presence of the feeding tube in the esophagus.
• This method is also useful for administration of nutrition to anorexic patients. Place a pharyngostomy tube if repeated dosing is necessary, using the technique for cats. Flush pharyngostomy tubes with water at least every 4 to 6 hours. Nasogastric tubes are not recommended because they are difficult to place and maintain patency because of their small size. Securely suture all tubes to the skin. Place a tube collar made of radiographic film or use rear leg hobbles to prevent removal of tubes.
• Nutritional support is critical in rodents because of their high metabolic rate. Provide supplements in animals that are anorexic for longer than 12 hours. If the GI tract is capable of digestion, use a slurry of pellets mixed with a high-calorie supplement. If the tube diameter is too small for this mixture, use avian hand-feeding formula or a mixture of vegetable and cereal baby foods in place of the pellets. If the ability of the GI tract to tolerate enteral feeding is questionable, first try isotonic electrolyte or dextrose solutions. Parenteral nutrition is used successfully in research animals and may be feasible in select pet cases.
Subcutaneous Injections
• Avoid irritating substances in rats and mice because their mammary tissue extends into these areas. The resulting inflammatory response is thought to increase the occurrence of mammary neoplasia.
Intravenous Injections
For small rodents, give a bolus of fluids every 2-4 hours, followed by a diluted heparin flush. A pediatric IV pump is used for continuous infusion of fluids to larger animals. Maintenance of catheters in active animals is extremely difficult.
ANESTHESIA
Premedication and Patient Preparation
• In chinchillas and guinea pigs, withhold food for 6 hours before anesthesia. Withhold food from smaller, mature rodents for 2 hours. Withhold food from immature animals for up to ½ hour depending on age and condition.
• Use heat lamps and heating pads to prevent hypothermia. Have a prewarmed incubator available for recovery. Preoperative or intra-operative warmed SC or IV fluids are strongly recommended. Place IV or IO catheters whenever possible.
• Administer atropine preoperatively to reduce airway secretions. Acepromazine, diazepam, or midazolam work well as premedications for other anesthetics. Avoid acepromazine in gerbils because it potentiates seizures. See Table 177-7 for anesthetic drugs and dosages.
Inhalation Anesthesia
Induction
Induce gas anesthesia using small face masks purchased from laboratory supply houses or make them from syringe cases and latex gloves (Fig. 177-7). Induction in an anesthetic chamber is also possible.
The risk of this behavior is reduced by premedication with tranquilizers, initial induction with nitrous oxide with later addition of primary anesthetic gas after relaxation, and low induction settings. Changes in respirations, especially erratic or apneustic patterns and decreased respiratory rates, indicate deepening anesthesia.
Injectable Anesthesia
Doses and routes for injectable anesthetics are listed in Table 177-7. Needed doses for injectable anesthetics are tremendously variable among species and individuals. Most injectable anesthetics provide safe sedation for minor procedures, but very few induce a safe surgical plane of anesthesia on a consistent basis.
• Ketamine in combination with diazepam is easily obtainable, is given intramuscularly, and has a wide margin of safety, but it does not provide good analgesia.
SURGERY
Hemostasis
• If necessary, give fresh blood transfusions drawn from a donor of the same species and mixed with sodium heparin (1000 IU/ml) at a rate of 0.005 ml/1 ml of blood directly into an IV or IO line.
Pain Management
Administer postoperative analgesics to all rodents undergoing surgical or dental procedures. Common analgesics include buprenorphine, butorphanol, ketoprofen, carprofen and meloxicam. See Table 177-7 for dosages.
Common Procedures
The most common surgeries are laceration repair and removal of dermal or SC masses.
Castration
Technique
1. Make a skin incision over each testicle or one incision on the midline just cranial to the prepuce.
2. Remove the testicles via a closed technique (i.e., do not open the vaginal tunics). All rodents possess open inguinal rings and can eviscerate if a closed technique is not used. Ligate the testicular vessels and vas deferens with absorbable suture
Dental Procedures
Incisors can be trimmed with nail trimmers, but this technique often fractures the tooth, causing abscesses of the root. Instead, use a high-speed dental burr or a flat cutting disk on a Dremel hand tool. Trim molars with a high-speed drill or pediatric rongeurs. A mouth speculum that deflects the tongue and other soft tissues is essential to prevent lacerations and provide working space (Fig. 177-8). Intubate the trachea to prevent aspiration pneumonia when working on molars.
If a tooth is abscessed, extract both it and the occlusal tooth.
• The roots of the maxillary incisors curve dramatically back into the head. Take care to follow the curve of the tooth.
• Packing an infected tooth socket with a calcium hydroxide paste may decrease the occurrence of persistent infection. Remove the paste in 3 to 4 days.
• Administer meloxicam, carprofen or ketoprofen postoperatively to control pain. See Table 177-7 for dosages.
MOUSE
Dermatology
Ectoparasites
Ectoparasites usually are found in new acquisitions.
• Alopecia and pruritus, especially on the back of the head and dorsal midline, usually are associated with lice (Polyplax serrata), mites, or fleas.
• Mite infestation (e.g., Mycoptes musculinus, Myobia musculini, Radfordia affinis) often causes a greasy haircoat and folliculitis.
• Diagnosis is based on clinical signs, history, visualization of parasite, skin scrape, and cellophane tape test.
• Treatment of fleas and lice is with Pyrethrin powder. Ivermectin is recommended for treatment of mites (see Table 177-8).
Dermatophytosis
• Alopecia also may be the result of dermatophytes (see Chapter 42). Lesions are often hyperkeratotic.
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