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.

Table 177-1 CAGING AND ENVIRONMENTAL REQUIREMENTS FOR POCKET PETS

Nutrition

Key Point

Feed pet rodents laboratory animal chow appropriate for their species (Table 177-2). Seed diets are deficient in protein and contain excessive fat.

Table 177-2 DIETARY INFORMATION FOR POCKET PETS

• 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.

• Supplementation of vitamin C is recommended for all guinea pigs.

• 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.

• Store food in tightly sealed containers at less than 60°F; keep food refrigerated if possible.

• 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.

Quarantine

Quarantine all newly acquired animals in a different room from current pets for a minimum of 30 days. Feed and handle quarantined animals last. Recommend that caretakers wash their hands and change clothes before handling current pets. Avoid the introduction of adult animals because this frequently results in fighting. Instead, place animals together while young and allow them to mature together. Avoid keeping more than one male per cage because this also usually leads to fighting.

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.

Table 177-3 NORMAL PHYSIOLOGIC DATA FOR POCKET PETS

Obtain the following information:

• Species, age, sex (species and sex may not be obvious or the owner may not know this information)

• Origin (e.g., private individual, breeder, pet shop, research laboratory, supply house)

• Length of ownership, owner’s previous experience

• 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

• Cagemates, other pets, quarantine history, presence of aggressive behavior

• Dates of unusual activity: breeding, parturition (Table 177-4), cannibalism, abnormal urination or defecation

• Medical history, previous weights

• Purpose for ownership: pet, breeder, display, education

• Purpose for visit: purchase examination, checkup, problem

Table 177-4 REPRODUCTIVE DATA FOR POCKET PETS

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.

Key Point

If dyspnea or severe depression is detected, warn the owner that the animal is critically ill and could die of stress brought on by an examination.

• 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.

DIAGNOSTIC Tests

Skin and Ear

Apply cellophane tape to crusted areas of the skin and view under a microscope as an aid in diagnosing ectoparasites such as lice, mites, and fleas. Skin scrapings are beneficial in detecting mites and dermatophytes. Dermatophytes are diagnosed best through culture of broken hairs or crust on dermatophyte test medium.

Use small, cotton-tipped swabs to obtain ear swabs from animals weighing more than 25 g. Mix debris with mineral oil and view under low magnification to test for ear mites, or roll onto a glass slide and Gram stain to look for bacterial or yeast infections.

Urine and Fecal Collection

Collect urine by placing the rodent in a clean meshbottomed cage with a plastic liner. After enough urine has been produced, collect it off the bottom of the cage with hematocrit tubes or a syringe and a 25-gauge needle. Perform cystocentesis on non-pregnant animals weighing more than 100 g with a 25- to 27-gauge needle.

Collect feces over several hours to provide a volume sufficient for fecal flotation. Flotation allows the detection of nematodes and some trematodes and cestodes. Cellophane tape applied to the perineal area and then viewed under a microscope often reveals oxyurid eggs. Use a fresh saline smear or fecal sedimentation to diagnose protozoal parasites. Fecal cultures are useful in diagnosing bacterial diarrhea.

Radiology

Radiology is an extremely useful tool. Machines capable of exposures as low as 40 kvp and 3 to 10 MAS effectively image mice. Most radiograph machines are capable of generating diagnostic radiographs of guinea pigs, chinchillas, and mature rats at settings used for kittens. Positioning is accomplished with masking tape or Velcro straps. Sedate unruly animals. Techniques used in cats for contrast studies of both urinary and GI systems are modified easily for use in pocket pets.

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.

• It is also possible to use the cephalic vein in guinea pigs.

• Jugular veins are good alternatives in thin individuals under sedation.

• 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.

Figure 177-4 Medial saphenous venipuncture; inset, lateral tarsal venipuncture.

Key Point

Attempt tail bleeding only as a last resort in mice, rats, gerbils, and hamsters. These techniques often are not acceptable to owners.

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.

Table 177-5 NORMAL HEMATOLOGIC VALUES FOR POCKET PETS

Table 177-6 NORMAL SERUM CHEMISTRIES FOR POCKET PETS

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.

Key Point

Take care not to place the tip into the contralateral cheek pouch, or the patient may store the medication and expel it later.

Administer medication in small amounts. Ensure that the animal swallows the medication in its mouth before more is administered. This technique is useful for force-feeding pellet gruels to anorexic pets if the caregiver is patient. Medication or food that is administered too quickly will be spit out or aspirated.

For rodents that are intractable or for administration of unpalatable substances, pass a stomach tube per os.

• 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.

Figure 177-5 Proper placement of a metal feeding tube.

Figure 177-6 Measurement and use of a flexible catheter as a feeding tube.

Key Point

Because the placement of a stomach tube is a blind procedure, administer a small volume of sterile saline into the tube before administering the medication to ensure that the tube is not in the trachea. Misplaced medications are fatal.

• 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.

Intramuscular Injections

Give IM injections in the semimembranous and semitendinous muscles. Inject only small volumes of nonirritating substances, or tissue damage with resulting self-mutilation may occur. Use the epaxial or triceps muscles if repeated injections are necessary.

Intravenous Injections

Give IV injections into any of the veins as previously described. In addition, the penile vein may be used in hamsters and guinea pigs. Placement of IV catheters is possible in animals heavier than 100 g.

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.

Intraosseous Injections

For IO injections, place a spinal needle into the proximal tibia or femur following the technique used for placing an intramedullary pin. Once the needle is seated, remove the stylet. Aspirate and check the hub of the needle for bone marrow. The tip of the needle should be in the bone marrow cavity that directly drains into the central venous system in normal bones (i.e., the cortex must be intact). Administer drugs, blood, or fluids at a rate similar to that used for IV catheters.

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.

Table 177-7 ANESTHETIC AND ANALGESIC DRUG DOSAGES FOR POCKET PETS

Monitoring

Surgical anesthesia is reached when toe, tail, and ear pinch fail to generate a withdrawal reaction.

Depth of anesthesia is best monitored by pulse and respiratory rate and character. Pulses drop to within normal ranges after induction. Further reduction, especially to less than 80% of the original stabilized value, is an indication to lighten the plane of anesthesia. Monitor the electrocardiogram (ECG) of small patients by clamping the alligator clips onto the hubs of all-metal 27-gauge needles or steel sutures placed through the skin at the usual sites. Tape cables to the table to maintain placement. Doppler units taped over the chest also provide accurate heart rates. Pulse oximeters are easier to use, more sensitive, and more expensive than the instruments mentioned previously. These instruments are easily taped to the patient’s ear, foot, or tail and provide heart rates as well as information regarding oxygenation.

Respirations are often shallow and rapid during induction. They become deep and regular as a surgical plane of anesthesia is reached.

The corneal reflex varies markedly between individuals and anesthetic agents. If the animal has a corneal reflex after induction and then loses it, reduce the anesthetic.

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.

Figure 177-7 A nose cone for anesthesia delivery can be made from a cut-down plastic syringe case and material from a latex glove. Syringe cases from 12 to 60 cc can be used, depending on the size of the patient.

All rodents induced and maintained on gas anesthesia require some form of non-rebreathing system. Usual induction is achieved between 2% and 3% for isoflurane and 2% and 4% for halothane. Maintenance for isoflurane and halothane varies from 0.25% to 2%. There is marked individual variation in the amount of anesthetic required for induction and maintenance. Use of 50% nitrous oxide in oxygen reduces anesthetic concentration requirements for other gases.

Key Point

Some chinchillas and guinea pigs hold their breath while being induced with gas anesthetics and then take deep rapid breaths. If the concentration of anesthetic gases is high enough, this behavior results in death.

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.

Endotracheal Intubation

Most pet rodents are not intubated for anesthesia because of their small size. When necessary, as in prolonged oral and other procedures, endotracheal intubation is accomplished with the animal in dorsal or ventral recumbency, depending on the clinician’s preference. Small non-cuffed or Cole endotracheal tubes work well. A stylet usually is required to provide enough stiffness for the tube to pass the larynx. Extend the animal’s head and neck. Grasp the tongue with forceps and use gentle traction. The tip of the tube then is advanced above the tongue and just past its base. The hard palate is used to deflect the tip of the tube ventrally into the glottis. This is a blind procedure that is difficult to master. Use of a laryngoscope is helpful in larger rodents.

Another technique is to place an over-the-needle catheter in the trachea and move it up retrograde through the larynx to act as a guide. The catheter is removed after the endotracheal tube is in place.

Key Point

If endotracheal intubation is performed, it is extremely important that the tube be checked for patency. Rodents produce copious respiratory secretions, which frequently clog endotracheal tubes.

The small diameter allows these tubes to collapse or kink, resulting in asphyxiation of the patient. Check patency at least every 2 minutes by applying positive pressure ventilation at 10 to 15 cm water and watching for excursion of the chest wall. If extending the head and neck does not result in air flow, suction the tube. If this is either not successful or impossible, remove the tube and continue anesthesia with a mask or reintubate the animal with a new tube. Because of the small diameter of the trachea, endotracheal tube-induced tracheitis and subsequent swelling of the trachea may become a life-threatening situation.

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.

• Intraperitoneal injections of barbiturates provide surgical anesthesia but have a low margin of safety and a significant mortality rate. Barbiturate anesthesia can result in fatal ileus.

Euthanasia

Euthanasia is performed easily by induction of inhalant anesthetic through a mask or chamber followed by an overdose of barbiturates given intraperitoneally, IV, or intracranially. Euthanasia by IP injection of barbiturates alone causes pain in some animals.

SURGERY

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.

Abdominal Surgery

Common abdominal surgeries include cystotomy for urolith removal in guinea pigs and rats, and cesarean section (C-section) in guinea pigs and chinchillas because of dystocia.

Use a technique similar to those described for dogs and cats. Preplaced stay sutures are recommended to define incision edges for closure. Use 4-0 polyglactin 910 or polydioxanone (PDS) on a taper needle and suture in a simple continuous pattern to close the body wall. Close the skin with a subcuticular suture (absorbable) or interrupted skin monofilament, non-absorbable suture.

Fracture Fixation

Fracture fixation is accomplished best with intramedullary pinning or Kirschner apparatus. Rodents gnaw on bandages until they remove them. If they are unable to remove a splint, self-mutilation often results in self-amputation. If a cast or splint is necessary, physical restraint often is required. Healing usually takes 3 to 6 weeks.

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.

Figure 177-8 Proper technique for dental trimming.

If a tooth is abscessed, extract both it and the occlusal tooth.

• If necessary, approach cheek teeth via an incision through the cheek.

• Use a fine dental elevator to loosen the teeth.

• Patience and firm but gentle pressure are needed, or the root or surrounding bone may fracture.

• 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.

In chinchillas with dental malocclusion, the roots of the molars can become impacted, causing swelling of the mandible or exophthalmos and epiphora. These teeth are extremely difficult to extract without causing extensive bony and soft-tissue damage. Discourage breeding of animals with malocclusion, unless it was acquired as a result of trauma or infection, because this trait is hereditary.

MOUSE

Most pet and laboratory mice are derived from Mus musculus, which is the common house mouse. Mice sold in the pet trade are randomly bred and less likely to suffer from the genetic problems associated with inbred laboratory rodents. Mice possess brown fat tissues between their scapulae that also are known as hibernating glands; these are thought to provide an energy store. The spleen in male mice is 50% larger than that of females.

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.

Transmission of lice and mites occurs via direct contact. Fleas are transmitted by other household pets, such as cats and dogs.

• 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).

• Change the bedding and thoroughly clean the cage between treatments to prevent reinfestation. Occasionally, the surrounding environment needs to be treated with a premises spray used for killing fleas.

Table 177-8 PARASITICIDE DOSAGES FOR PET RODENTS

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