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.
Caging and Sanitation
Feed pet rodents laboratory animal chow appropriate for their species (Table 177-2). Seed diets are deficient in protein and contain excessive fat.
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.
Obtain the following information:
Examination of Patient and Environment
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.
Evaluate the Cage
Examine the Animal
Once the animal is restrained properly, examine the head. Assess the cranial nerves. Check the nose for presence and character of discharge. Examine the mouth for ptyalism, swelling, overgrown incisors, or discharges. To inspect the oral cavity, place an avian speculum across the mouth just caudal to the incisors. Use a light source and a pair of hemostats as retractors to improve access. Alternatively, use an otoscope with a pediatric head to examine the premolars and molars of guinea pigs and chinchillas for overgrowth. Examine the cheek pouches of hamsters for swelling, impaction, or discharge. An ophthalmic examination, including a fundic examination, is important.
Gerbils, rats, and mice produce red tears (chromodacryorrhea) with stress or disease. Do not confuse them with hemorrhage.
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 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.
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.
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.
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.
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.
Administer SC injectable medications or fluids over the shoulder blades or in folds of skin on the flank.
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.
Use intraperitoneal (IP) injections only as a last resort for large volumes of fluids or for irritating injections that cannot be administered via an IV or IO route.
Express the bladder and aseptically prepare the abdomen.
Restrain the rodent with its head down to move the abdominal organs cranially. Give the injection 0.5 to 2 cm lateral to the midline in the caudal abdomen. Aspirate before injecting to ensure that the injection is not being given into the bladder or bowel. Never use this technique in pregnant animals.
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.
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.
Premedication and Patient Preparation
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.
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.
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.
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.
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.
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.
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.
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.
The most common surgeries are laceration repair and removal of dermal or SC masses.
Castration is a common procedure in guinea pigs. This usually is performed when owners want to house more than one male together or do not wish to breed their female any longer.
Some surgeons partially suture the inguinal rings for extra security. This technique is also applicable to other rodent species.
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 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.
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.
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.
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.
Ectoparasites usually are found in new acquisitions.
Transmission of lice and mites occurs via direct contact. Fleas are transmitted by other household pets, such as cats and dogs.