Reproductive medicine of rabbits and rodents

Vet Clin Exot Anim 5 (2002) 507–535

Reproductive medicine of rabbits and rodents Cynthia R. Bishop, DVMa,b,* a

Biology Department and PPHS (Pre-Professional Health Sciences), Seattle Pacific University, School of Natural and Mathematical Sciences, 3307 Third Avenue West, Seattle, WA 98119, USA b Animal Emergency and Referral Center, 19511-24th Avenue West, Lynnwood, WA 98036, USA

The small animal practitioner is likely to be called upon to help nontraditional pets such as rabbits, rodents, and ferrets [20]. Although ferrets, rabbits, guinea pigs, rats, and mice have been domesticated for centuries, much of the information about their diseases and unique anesthetic and surgical needs has only been obtained within the last 10 years [15,16]. Hamsters, gerbils, and chinchillas have been domesticated much more recently, while animals such as prairie dogs and hedgehogs are not domesticated but have recently become popular as pets [27,28]. The more information veterinarians have regarding normal anatomy, physiology, reproduction, medicine, anesthesia, and surgery of these unusual pets, the more likely they will be able to help their owners. This article presents an overview of routine and emergency reproductive concerns that the small animal practitioner may face when presented with nontraditional mammalian patients. Rabbits The veterinarian may be called upon to help with the reproductive medicine and surgery of rabbits raised for research, meat, pelt, exhibition, or as family pets. An understanding of normal reproductive anatomy and physiology (Table 1) is necessary before understanding reproductive diseases. Although the reproductive system is similar among all rabbits, the ultimate * Biology Department and PPHS (Pre-Professional Health Sciences), Seattle Pacific University, School of Natural and Mathematical Sciences, 3307 Third Avenue West, Seattle, WA 98119. E-mail address: [email protected] 1094-9194/02/$ - see front matter Ó 2002, Elsevier Science (USA). All rights reserved. PII: S 1 0 9 4 - 9 1 9 4 ( 0 2 ) 0 0 0 1 9 - 1

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Table 1 Physiological data of the rabbit Temperature Heart rate Respiratory rate Life span Male Female Babies Estrus Gestation Sexual maturity Litter size Weaning age

37.8°–38.8°C (100°–102°F) 100–300 bpm 30–60/min 5–10 years Buck Doe Kits Induced ovulator/silent estrus 31 days Ranges from 4–9 months Dwarf breeds earlier, giant breeds later 3–12 Dwarf breeds fewer, giant breeds more 6–9 weeks

Data from Donnelly T. Basic anatomy, physiology, and husbandry (rabbits). In: Hillyer E, Quesenberry K, editors. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders; 1997. p. 154–6; and Harkness J, Wagner J. The biology and medicine of rabbits and rodents, 4th edition. Baltimore: Williams and Wilkins; 1995.

goal may vary tremendously. The veterinarian who treats rabbits raised for meat production may primarily be concerned with diseases that affect production; the veterinarian who treats pet rabbit may be concerned with the safest techniques for neutering the animal. Reproductive anatomy The female rabbit, or doe, has typical ovaries and uterine horns; however, rabbits are unique in that they have two cervices that are separate from each other and lead to each uterine horn. Abundant adipose tissue is found surrounding the ovaries, uterine horns, and the associated ligaments [1,2]. This is a particular concern in obese females and can be an annoyance when routine surgery (eg, ovariohysterectomy) is performed. In emergencies where a C-section or uterine neoplasia are involved, the excessive fat can be a serious concern. Specifics regarding uterine neoplasia and reproductive surgery will be discussed later in this article. The male rabbit, or buck, has typical testes and epididymis; however, the inguinal ring remains open after the testicles descend. The testicles may be moved freely between the scrotum and the abdominal cavity [1,2]. This can be of concern during routine surgeries (eg, castration) if the surgeon was not prepared to trap the testicle outside of the inguinal ring before making the first incision. Surgery of the male reproductive tract will be discussed later in this article. Determining the sex of young rabbits can be difficult for inexperienced owners and veterinarians as well. Unlike most rodents, there is very little difference in the anogenital distance between the doe and the buck. Anterior to the anus is a second opening or ridge that conceals the vulva or penis.

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Stretching this structure will reveal a slit surrounded by the vulva in the female. Slight pressure around the area will usually cause the penis to be everted in the male. In adult males there are two obvious scrotal pouches, one on either side of the rectal area. It is more difficult to determine the sex in immature rabbits; the vulva is more attached at the anal end, making a slit, and the penis is completely round [2,3]. Reproductive physiology Rabbits, like the cat and ferret, are induced ovulators [21]. They do not have an estrus cycle like some other mammals; however, there is some rhythm to their receptivity. Receptivity may be indicated by a swollen, reddened vulva, restlessness, chin rubbing, lordosis, and allowing the buck to mount. When the doe is not receptive she will run away, vocalize, or bite the buck and refuse to allow mounting. Vaginal smears do not appear to be useful in detecting estrus [1,2]. The doe ovulates 9 to 13 hours after copulation, and gestation ranges from 30 to 33 days (average: 31 days). Does may be palpated at 10 to 14 days postcopulation for evidence of fetuses within the uterus [4]. They will be smaller than the kidneys but larger than fecal pellets (ie, marble-sized). The doe can be palpated again at 26 to 28 days to check for resorption. An ultrasound can be performed to confirm pregnancy and determine viability of the kits if palpation is inconclusive. A radiograph can be taken after the skeletons have calcified in approximately the last trimester of the pregnancy. The doe will pull hair out and make a nest within 1 week of kindling. Does that are gestating for the first time may pull inadequate amounts of hair or make an insufficient nest. In addition, they may not produce enough milk or may neglect their babies as well [1,2]. These does are frequently successful with their next litter, however. Does normally only nurse their babies once or twice a day for a short time. False pregnancies can occur and usually last about 18 days. Does may be bred back within 48 hours of parturition during a postpartum receptive period [1,2]. Domestic rabbits are hairless at birth, with closed eyelids and ear canals, and are helpless. They remain hidden in the nest for about 3 weeks. Although the doe may guard her nest, she does not stay in the nest except when nursing. The mother’s fur and brown fat (the primary organ of thermogenesis) help keep the neonates warm. The brown fat is used during the first 2 weeks of life [1,2]. This fat is also used by hibernating animals during arousal when high-energy demand occurs. This brown fat, rich in large mitochondria, produces heat rather than ATP energy during cellular metabolism. If the doe was bred back to the buck during postpartum estrus, she may abandon or even kill her first litter as she prepares for the new litter. She may not harm the kits in her first litter, but she may quit nursing them, which means they may need to be supplemented until they are old enough to wean. The weaning age varies from breed to breed. Dwarf breeds may wean as early as 4 to 6 weeks, while larger breeds may not wean until 7 to 8 weeks.

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This is a very critical and stressful time for the kits. They may lose weight and have trouble adjusting to watering systems and commercial feed. As they switch over from primarily foregut liquid digestion (milk) to hindgut, cellulose digestion, many problems can occur. Normally the kits will eat some of the doe’s feces, which may help innoculate their digestive tract with the symbiotic organisms necessary for cellulose digestion. Kits that have been raised as orphans or that have heavy parasite loads may develop severe diarrhea and die. I recommend using a probiotic or fecal slurry from a healthy adult rabbit when handfeeding the babies. Many substitutes have been used for orphan kits that cannot be fostered to another doe. Kitten milk replacers, puppy milk replacers, and goat’s milk have all been successful. I discovered that regardless of the supplement used, there should not be a sudden change to another supplement, because this can lead to severe diarrhea and death. Does produce colostrum for 2 to 3 days postpartum, after which milk quantity increases and the composition changes to approximately 10% protein, 12% fat, and 2% carbohydrate [1,2]. Reproductive surgery There are many factors that can lead to the presentation of a doe or buck for reproductive surgery. These include the prevention of pregnancy, prevention of reproductive disease, treatment of behavioral problems, and neoplasia [5]. The surgical procedure may be similar to that performed on other small mammals; however, certain special precautions are warranted. Unlike canine, feline and ferret patients, rabbits cannot vomit [6,29]. It is not necessary to fast a rabbit surgical patient overnight. Fasting for a couple of hours before surgery may decrease stomach volume and food packed in the mouth. It is not recommended to withhold water from the patient before surgery [7,8]. I recommend allowing rabbits to graze up until a couple of hours before surgery, then encouraging them to eat as soon after surgery as possible [6]. This may help maintain normal intestinal function. Part of the preanesthetic preparation should include cleaning out the mouth to remove food particles and excessive saliva. Use of anti-inflammatory agents and analgesics can help before and after surgery. Controlling stress and pain will allow the patient to recover more quickly and return to a normal appetite and activity level [6]. Adhesions, peritonitis, and ileus are potential complications in any rabbit abdominal surgery. Adhesions may be minimized by careful tissue handling, keeping tissues moist, and careful choice of suture materials. Use of anti-inflammatory drugs such as flunixine (Banamine, Schering-Plough Animal Health, Union, NJ, USA) or calcium channel-blocking agents such as verapamil (Calan, Searle Pharmaceuticals, Skokie, IL, USA) postoperatively have shown promise [7]. Peritonitis is a serious concern and is a potential complication with any abdominal surgery. Attention to sterility and lavage of the abdomen with warm, sterile saline can be very successful. Even gentle

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manipulation of the gastrointestinal tract while performing reproductive surgery can lead to postsurgical ileus. Motility-stimulating agents such as metoclopramide (Reglan, Faulding Pharmaceutical Co., Elizabeth, NJ, USA) may be necessary. Because a rabbit’s immune response is caseous in nature, it is wise to use the least reactive suture materials that are available [7]. I prefer the monofilament synthetic absorbables (eg, PDS (Polydioxanone Suture), Ethicon, Inc., Somerville, NJ, USA). Stainless metal clips for hemostasis are also very useful (Hemoclips, Dilling Weck, Research Triangle Park, NC) [7]. In my experience, catgut causes tremendous tissue reaction and can lead to many secondary complications. Anesthesia and surgical preparation The most important piece of anesthetic equipment for an exotic animal veterinarian is an isoflurane anesthetic machine [8]. Isoflurane allows for quick induction and recovery. Many short procedures can be done using only a mask and nonrebreathing set up with isoflurane, while longer procedures may be performed using isoflurane via a face mask or intubation, when possible. I have found that preanesthetic drug use can also be helpful and less stressful for rabbits and other rodents. Many different protocols exist, and different veterinarians have their own favorite combinations. I prefer to use isoflurane alone, via a face mask, for very short procedures such as radiographic positioning, ultrasound, blood collection, intravenous (IV) catheter placement, and minor wound repair [6]. Some of the procedures mentioned do not usually require anesthesia; however, when anesthesia is required, I prefer the quick masking down with isoflurane. Rabbits and some rodents are very fearful of the face mask and object to the smell of the isoflurane gas. In those cases, I prefer to add a preanesthetic agent such as acepromazine, diazepam, butorphenol, buprenorphine, oxymorphone, or a combination. In cases where tracheal intubation is not possible, injectable anesthetics may be preferred with isoflurane as the maintenance anesthetic by mask. Combinations of acepromazine–ketamine, diazepam–ketamine, and propofol have been used successfully in our hospital with or without isoflurane for maintenance. When using a mask, before intubation or for the delivery of isoflurane for surgery, it is much less stressful for the rabbit if you deliver pure oxygen for a few minutes and then slowly increase the percentage of anesthetic gas until the righting reflex is gone. Anticholinergics such as atropine or glycopyrolate can be used to help decrease the salivary secretions and are easily incorporated into the preanesthetic protocols. Although a percentage of rabbits (30%) have the enzyme atropinesterase, it can still be useful as a preanesthetic or as an emergency drug in cases of severe bradycardia in most rabbits and in rodents [8]. As mentioned previously, I recommend thoroughly swabbing out the mouth before using the face mask or intubation. Once the animal is anesthetized, I usually examine the mouth further to identify any potential problems

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before proceeding [6]. Artificial tear ointment is applied to the eyes, and the surgical site is clipped. Rabbits have very delicate skin, which can easily be torn or cut when clipping fur with standard clippers and a number 40 blade; stretching the skin tight in front of the direction of clipper motion may help [7]. Using new or recently sharpened clipper blades that are free of animal hair or fur and that are well lubricated will also help. I clip a minimal area for surgery, and after the area is scrubbed clean I use lubricant jelly to hold back the hair or fur at the edges of the clipped area. I usually put gentle pressure in the lower abdomen to facilitate emptying of the bladder. Too much pressure should be avoided; otherwise the bladder or delicate intestinal tissue may be damaged. Surgical scrub and preparation can be performed using the same methods used for dogs and cats. Very small rodents can be more susceptible to heat loss because of their greater surface area to mass ratio [22,23]. I do not use alcohol in smaller patients because of its supercooling effect. Instead, I use a chlorhexidine solution that is diluted with warm water after the initial chlorhexidine scrub preparation. Another precaution that I use when positioning the rabbit patient on the surgical table involves loosely tying down the legs, unlike the method of more firmly tied legs for dogs and cats [6]. Rabbits have thin cortices in their bones and very strong muscles [1,8]; this combination can lead to a vertebral fracture when the animal is tied down before, during, and after surgery. Rabbits and rodents have a comparatively small thoracic cavity [1,8]. I recommend elevating the chest so that abdominal organs do not put pressure on the diaphragm and in turn make it difficult for the lungs to expand. This is the opposite technique of that used frequently in dogs and cats [6]. Some surgical tables slant cranially to shift the liver and gastrointestinal organs forward and away from the site of reproductive surgery. This type of positioning could be fatal in the rabbit or rodent. I have also learned not to recover rabbits and rodents before the procedure is 100% completed. Because rabbits and rodents are prey species and can go into shock quickly, I use an analgesic presurgically and again postsurgically [8]. I have witnessed rabbits suddenly scream and die after recovering too quickly during a minor surgical procedure. Rabbits in these types of situations have also been saved by quickly increasing the isoflurane percentage and covering their eyes to reduce the panic and pain. Ovariohysterectomy (spaying) After the rabbit has been anesthetized and positioned in dorsal recumbency and the surgical area has been prepared for surgery, the procedure can begin. A ventral, midline incision should be made starting midway between the umbilicus and pubic bones and extending caudally 1 to 3 inches. The skin in this area is usually very thin and delicate, so it does not require much digital pressure on the scalpel. I use small rat-toothed forceps to lift up the linea alba before making my stab incision into the abdominal cavity. The large cecum, bladder, stomach, or intestines are frequently lying just beneath the

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thin muscle layer and are easily nicked if the approach is too aggressive. Once inside the abdominal cavity, I do not use any instruments with sharp points to prevent inadvertent puncture of gastrointestinal organs when working around them. The uterus is usually very easy to identify in the caudal abdominal cavity. It is rarely necessary to use a snook hook (spay hook) [7]. Once the uterus is identified, the oviduct, infundibulum, and ovary can be found on each side. The ovary is frequently surrounded by fat and may be difficult to see initially. The mesometrium is also an area of fat storage and can make identification of the vessels a challenge [7]. Ligation of the ovarian pedicle with suture material or hemoclips should now be performed, followed by gentle retraction of the infundibulum and oviduct. Gently tear apart the ligaments to release the oviduct. There will be varying amounts and sizes of vessels within these ligaments depending on the age, health, and breeding status of the doe. Significantly sized vessels should be ligated or cauterized as the procedure progresses. There are two primary approaches to the removal of the uterus. The first involves ligating the bicornate uterus separately just cranial to the two cervices. Unlike a dog or a cat, a rabbit has no distinct uterine body. The second technique involves making one ligature just caudal to the cervices in the cranial vaginal area. This technique is commonly used in cases of pyometra or uterine neoplasia, whereas the first technique is used more often during routine ovariohysetectomy. If pyometra or uterine neoplasia are suspected, the entire cervices regions should be removed. Caution must be taken to prevent damage to the urethra, which enters into the proximal end of the vaginal vestibule [7]. I recommend that the vestibule be oversewn to prevent leakage of vaginal or urinary contents. I also recommend abdominal lavage with sterile saline solution that is warmed to body temperature. The uterine vessels may be large and buried in the fat. They should be double-ligated, either separately from the uterus or transfixed to the uterus [7]. When ligatures are placed, caution must be used to prevent incorporating intestinal serosa in with the reproductive tract. The abdomen is then closed in a simple interrupted pattern, again, being cautious not to inadvertently incorporate the intestinal tract into the closure. The skin can be closed using subcuticular patterns, tissue glue, skin staples or a combination of the three. Rabbits are usually very effective at removing more traditional skin sutures and can occasionally remove the skin staples as well. Orchidectomy (castration) The reproductive anatomy of a male rabbit is similar to that of dogs and cats, although the testicle is more elongated and the inguinal ring remains open throughout their life; this allows the testicles to be withdrawn into the abdominal cavity [1,2]. After the rabbit has been anesthetized and positioned in dorsal recumbency and the surgical area has been prepared for surgery, the procedure may begin.

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There are primarily three different approaches to castration of the buck [7,9]. Some surgeons make separate incisions in the ventral surface of the scrotum and exteriorize the testicles and epididymis from each incision; this technique is similar to that performed in most cats. Other surgeons make prescrotal incisions separately over each cranial scrotal region. I prefer to make one central prescrotal incision anterior to the penis and remove both testes from the same incision. This technique is similar to that used in the castration of dogs. Regardless of incision technique, the entire testicle and epididymis must be exteriorized to expose the spermatic cord with the vessels and vas deferens for ligation. The spermatic cord can be ligated as one unit in a closed fashion using suture material, hemoclips, or the overhand knot (autoligation technique) [7]. I prefer to use an open technique and double ligation of the spermatic vessels and vas deferens. This way the tunic can be ligated and the inguinal ring can be closed at the same time. The possibility of umbilical herniation and intestinal strangulation is a concern after castration. Intestinal herniation and strangulation are prevented in the intact buck by the large amount of fat present around the spermatic cord and epididymis [7]. I strongly recommend closing the neuter site rather than leaving it open, as is commonly performed on cats. When a rabbit sits down, the surgical site is touching the ground, whereas when a cat sits down the surgical site is still above the ground. The caseous type of inflammatory response in rabbits can again create a problem if the incision site is left open or if they react to the suture material. I prefer to close the skin in a subcuticular pattern and use tissue glue over the top for added insurance. If the rabbit has diarrhea, has dirty scent folds, or chews the incision open, an infection is still possible. Caesarian section (C-section) The small animal veterinarian may be called upon to perform a C-section if a doe is having trouble giving birth naturally. This could be due to fetuses that are too large or uterine inertia that is not responding to medical treatment. The procedure used can be similar to the procedure that is performed on cats and dogs. The exception would be if a spay is to be performed at the same time. If the rabbit is to be spayed, the precautions listed in the previous section on ovariohysterectomy should be observed [5]. I recommend pain medication for several days postoperatively so that the doe is more likely to allow nursing. An alternative would be to foster the babies to another nursing doe if that is possible. Reproductive diseases Uterine neoplasia Does that are not spayed have a high occurrence of uterine neoplasia. The most common neoplastic disease of rabbits is uterine adenocarcinoma [10]. Other neoplastic conditions such as adenomas, leiomyosarcomas, and

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leiomyomas can also occur. Uterine neoplasia appears to be independent of breeding history; however, certain breeds are more predisposed (eg, Tan, French silver, Havana, and Dutch) [10]. Uterine cancer, however, has been seen in rabbits of all breeds, including mixed breeds. Adenocarcinoma of the uterus is usually a slow, progressive condition with local spread to the myometrium and peritoneal cavity. Within a few years hematogenous metastasis to the lungs, liver, and bones may occur [10]. Symptoms vary but may include decreased reproductive success (eg, infertility, smaller litter size, resorption), hematuria (a thick, dark vaginal discharge), weight loss with or without a pendulous abdomen, decreased appetite, and respiratory compromise [2,10]. Diagnosis can be obtained from abdominal palpation of the uterine masses, abdominal radiographs or ultrasound, or exploratory surgery. Treatment primarily involves immediate ovariohysterectomy. Radiographs of the lungs should be taken before surgery and every 3 to 6 months postsurgery [10]. The prognosis is poor if pulmonary metastasis has already occurred before surgery [10,11]. Early spay of rabbits is considered to be the best prevention. I recommend that surgery be performed at 6 to 12 months of age. Smaller breeds sexually mature earlier than larger breeds, so they may be spayed at a slightly younger age. Benign endometrial hyperplasia Many benign conditions of the ovaries and uterus can occur. Cysts, polyps, and hyperplasia can be difficult to differentiate from cases of adenocarcinoma [10]. Hematuria, lethargy, palpably enlarged uterus, radiographically enlarged uterus, and cystic mammary glands can all occur in either benign or malignant cases [10]. Exploratory surgery, ovariohysterectomy, and histopathology are the only definitive ways to make the diagnosis. Infectious reproductive disorders Pyometra and endometritis can occur in breeding and nonbreeding does. Signs may include vaginal discharge, abdominal swelling, depression, anorexia, fever, and hypothermia. History will frequently include recent breeding or birth, infertility, and stillbirths [10]; however, does that have never been bred can still develop reproductive infections. Diagnosis may include palpation of an enlarged, doughy uterus, radiographic evidence of uterine enlargement, ultrasound confirmation of a fluid filled uterus or leukocytosis with neutrophilia [5,10]. Use caution when palpating does with suspected uterine disease, because the uterine tissue can be extremely friable, making rupture possible [5,10]. Exploratory surgery with ovariohysterectomy is necessary for a definitive diagnosis and treatment. Antibiotic therapy and intravenous fluid therapy should be started before anesthesia and surgery. A broad spectrum antibiotic can be chosen while a culture and sensitivity are pending. The two most commonly isolated organisms are Pasteurella multocida and Staphylococcus aureus [10]. The most common source of infection is transmission during mating, although hematogenous spread has also

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occurred [10]. Other organisms such as Chlamydia, Listeria monocytogenes, Moraxella bovis, Actinomyces pyogenes, Brucella melitensis, and Salmonella species have also been isolated from pyometra cases [2,5,10]. As with other sexually transmitted diseases, infectious reproductive disorders are easier to prevent than to treat. I recommend screening all breeding animals for P multocida using either cultures or serology and using virgin does and bucks when outside lines are introduced to a rabbitry [5]. Orchiitis and epididymitis can occur in breeding and pet bucks. Signs may include fever, anorexia, weight loss, swelling of the scrotal area, and prepucial discharge. The cause may be sexual transmission from a doe, fight wounds from another buck, or hematogenous spread [2,5,10]. Diagnosis and treatment usually involve castration. P multocida and Treponema cuniculi are frequently involved pathogens. T cuniculi is a spirochete that is responsible for rabbit syphilis (vent disease) [2,6,10]. Clinical signs vary but may include areas of redness and swelling in the perineal area with vesicles or scabs. The nose, face, and lips may also be infected [2,5,10]. This disease is usually self-limiting, but those that recover are frequently carriers. Diagnosis requires skin biopsies with silver staining or serology. There are two published treatment protocols: penicillin G benzathine and penicillin G procaine (42,000–84,000 IU/kg subcutaneously (SC) at 7-day intervals for three injections) or penicillin G procaine (40,000–60,000 IU/kg every 24 hours for 5–7 days) [2,10,33]. All exposed animals must be treated. Close observation for decrease in appetite, lethargy, diarrhea, or abdominal distension is necessary. Stop treatment immediately if any of the toxic effects of penicillin are seen. Feeding a high fiber diet and a probiotic supplement during treatment can be very helpful. Noninfectious reproductive disorders Noninfectious reproductive disorders can result in reduced fertility, resorption, abortion, dystocia, stillbirths, agalactia, and pregnancy toxemia [2,5,10]. Reduced fertility or infertility can be caused by the infectious agents mentioned previously or by noninfectious factors such as inadequate nutrition (protein deficiency, hypovitaminosis A, D, E or hypervitaminosis A), environmental stress, hyperthermia, and congenital anomalies. Vitamin E deficiency causes myodystrophy; a high-serum creatine phosphokinase can support this diagnosis [10]. Fetal death after less than 3 weeks of gestation usually results in resorption and may be mistaken for a breeding that was not successful [10]. Because the causes of infertility may be different from those of resorption, it may be important to differentiate between the two. If this is occurring on a large scale within a rabbitry, evaluation of diet and environment as well as screening for infectious diseases is warranted. If a doe has two consecutive pregnancies that do not produce live births, I recommend an ultrasound to confirm a true pregnancy the next time she is bred; if she doesn’t deliver, it may be a case of resorption rather than infertility. If a prize buck suddenly stops having successful litters, his sperm could be evaluated and any does

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bred to him could be given an ultrasound to determine if resorption is occurring. This could indicate a sexually transmitted disease from the buck to the does, which would explain the unsuccessful reproduction. Fetal death after 3 weeks usually results in abortion. Submit a fetus for bacterial culture and histopathology to determine the cause of the abortion. Infectious agents such as Listeriosis are more common in late-term abortions. Stress, trauma, hyperthermia, toxic plants, drugs, or severe dietary deficiencies can also lead to abortions [2,5,10]. In my experience, secondary uterine infections are more likely to occur in cases of abortion than in cases of resorption. Dystocia is less common in rabbits than in rodents [19]. Causes may include oversized kits, narrow pelvic canal, obesity, nutritional deficiencies, or uterine inertia [2,5,10]. Contractions and straining that produce bloody or greenish discharge but no kits may indicate dystocia. Manual assistance with removal from the vaginal canal may be necessary [10]. Application of a lubricant (eg, a water-based jelly) may help the passage of the kits. In cases of suspected uterine inertia, calcium should be administered first, followed by oxytocin if necessary. Calcium gluconate can be given either orally or by injection 30 minutes before administering the oxytocin (dosages are outlined in Box 1). Place the doe in a quiet, stress-free environment while waiting to see if treatment is successful. A C-section may be necessary if medical treatment is unsuccessful. Pregnancy toxemia is most common in the last week of gestation. Obese rabbits are more predisposed to this condition, but additional factors may include environmental stress, inadequate nutrition, hyperthermia, or extremely young or old does. Clinical signs include lethargy, decreased appetite and activity, weakness, incoordination, ketone smell to breath, abortion, convulsions, and coma. Some rabbits linger over several days and others die acutely [2,5,10]. A urinalysis will reveal acidic urine, proteinuria, and ketonuria, and the urine may be clearer than normal because of a decrease in calcium carbonate crystals. Serum chemistries may reveal acidosis, ketonemia, hyperkalemia, hyperphosphatemia, and hypocalcemia [2,10]. Hepatic lipidosis is commonly found on ultrasound or necropsy. Prevention is much more rewarding than treatment. Obese does should not be bred, and fasting, undernutrition, and stress should be avoided during pregnancy. Treatment can be attempted with intravenous or intraosseous fluids [10]. Parenteral nutrition can be accomplished with syringe feeding, orogastric tube feeding, nasogastric tube placement, or esophagostomy tube placement [5]. Formulations for assisted feeding are available (Oxbow Hay and Feed), otherwise children’s formulas may be used. Monitor blood gases for acid/base level, electrolytes, calcium and phosphorus. There are reports of other less common conditions such as uterine torsion, vaginal or uterine prolapse, hydrometra, and endometrial aneurysms [2,10]. Treatment for these requires emergency stabilization and ovariohysterectomy as soon as possible [18].

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Box 1. Drug dosages Amikacin 8–16 mg/kg (total dose; divided q8–24 h; SC, IM, IV) Atropine 0.1–0.5 mg/kg; SC, IM Buprenorphine 0.01–0.05 mg/kg q 8–12h; SC, IM, IV Butorphenol 0.1–0.5 mg/kg q2–8h; PO, SC, IM, IV Calcium gluconate (10%) 1–2 ml/kg q12–24h; PO Chloramphenicol 30–50 mg/kg q8–12h; PO, SC, IM, IV Cimetidine 5–10 mg/kg q8–12h; PO, SC, IM, IV Ciprofloxacin 5–15 mg/kg q12h; PO Cisapride 0.5 mg/kg q8–24h; PO Dexamethasone 0.1–2 mg/kg; PO, SC, IM, IV Doxycycline 2.5 mg/kg q12h; PO Enrofloxacin 5–15 mg/kg q12h; PO, SC, IM Flunixin 0.3–1.0 mg/kg q12–24h SC, IM (no more than 3 days) Metoclopramide 0.5 mg/kg q8h; PO, SC Metronidazole 20 mg/kg q12h; PO for 3–5 days Oxytocin 1–2 units per cavy or rabbit; SC, IM Penicillin G (benzathine and penicillin G procaine) 47,000–84,000 IU/kg once per week; SC, IM Prednisone 0.5–2 mg/kg; PO Trimethoprim/sulfamethoxazole 15–30 mg/kg q12h; PO Vitamin K1 1–10 mg/kg; SC as needed Data from Bishop C. Emergency medicine and surgery of rabbits and rodents. Toronto: Ontario Veterinary Medical Association; 1999.

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Mammary gland disorders Disorders of the mammary gland can interfere with normal nursing and survival of both the doe and kits [12]. Mastitis can occur in either lactating does or those in false pregnancy and is most often caused by infection. Clinical signs include firm, hot, and swollen mammary glands; lethargy; fever; and anorexia. These symptoms can lead to septicemia and death [2,5,10]. Start on broad-spectrum antibiotics and fluid therapy while culture and sensitivity of the mammary discharge or milk is pending. Hot-packing the affected glands and draining them several times a day is essential [10]. Nonsteriodal anti-inflammatory agents such as flunixin (Banamine) or analgesics such as butorphenol or buprenorphine may help during treatment and may help improve appetite and activity levels. The kits will need to be removed from the doe; however, fostering them could spread the infection to another doe and litter [10]. Cystic mastitis is a noninfectious condition seen in breeding and nonbreeding does. The affected mammary glands can be swollen and firm as well but usually have a clear or serosanguinous discharge. Look for an underlying cause such as uterine hyperplasia or neoplasia [10]. Ovariohysterectomy usually causes the mammary glands to regress and return to normal. If left untreated, these glands can undergo metaplastic changes and lead to adenocarcinoma, which can metastasize to the regional lymph nodes and lungs [10].

Guinea pigs I have bred and exhibited guinea pigs for 20 years. I have also had the pleasure of treating guinea pigs as patients in both laboratory and small animal clinical settings for the last 18 years. They are by far my favorite of all the small rodents. The scientific name for a guinea pig is Cavia porcellus, from which the name cavy is derived (most guinea pig breeders and exhibiters prefer to call guinea pigs cavies) [13]. As a small animal veterinarian I have seen guinea pig patients as part of a large breeding facility (200–500 animals), in a laboratory animal facility, in animal wholesale settings, and as individual family pets. Much of my experience in guinea pig reproductive surgery comes from the individual pet or the rescue organizations that I have worked with. Guinea pigs make wonderful pets for children and adults and are delightful to raise and exhibit. Because of their long gestation and the large size of the pups at birth, reproductive problems are more common than in rabbits. Another unique characteristic about guinea pigs is their dietary requirement for vitamin C on a daily basis. Vitamin C deficiency, or scurvy, is a common reason for presentation to the small animal veterinarian. An understanding of normal reproductive anatomy and physiology (Table 2) is necessary before understanding reproductive diseases.

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Table 2 Physiological data of the guinea pig Temperature Heart rate Respiratory rate Life span Male Female Babies Estrus

Gestation Sexual maturity Recommended Breeding onset Litter size Birth weight Weaning age

37.2°–38.8°C (99°–102°F) 250–320 bpm 90–150/min 4–6 years Boar Sow Pups Nonseasonally polyestrus 15–17 days Spontaneous ovulation Duration (1–16 hours) 59–72 days (average: 68) 5–10 weeks (male) 4–6 weeks (female) 550–700 g (3–4 mo) male 350–500 g (3–5 mo) female 1–6 (average: 2–4) 60–100 g 3–4 weeks (150–200 g)

Data from Refs. [14,16,25,30].

Reproductive anatomy The female guinea pig, or sow, has an ovary, infundibulum, and uterine horn on each side, similar to the ferret [26], dog or cat [2,13,14]. Unlike the rabbit, the guinea pig has only one central cervix that separates the uterine body from the vagina. The obese guinea pig may have abundant adipose tissue around the ovaries, uterine horns, and the associated ligaments. This can be an annoyance when routine surgery (eg, ovariohysterectomy) is performed. An even greater challenge when performing a guinea pig spay is the anatomical position of the ovaries, which are found just caudal to the kidney on either side close to the dorsal wall of the abdominal cavity. Specifics regarding uterine neoplasia and reproductive surgery will be discussed later in this article. The male guinea pig, or boar, has typical testes and epididymis as well as several accessory sex glands. Boars have a bulbourethral gland, coagulating gland, seminal vesicles, and a prostate gland. The seminal vesicles lie ventral to the urethra and extend into the abdominal cavity, which could be mistaken for uterine horns [2,13]. Boars have an os penis bone and two horny styles (slender projections off the penis) [2,13,14]. Similar to the rabbit the inguinal ring remains open after the testicles descend allowing the testicles to be moved freely between the scrotum and the abdominal cavity [2,13]. This can be of concern during routine surgeries such as castration if the surgeon was not prepared to trap the testicle outside of the inguinal ring prior to making the first incision. Surgery of the male reproductive tract will be discussed later in this article.

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Determining the sex of young guinea pigs can be difficult for inexperienced owners and veterinarians alike. Unlike other rodents, there is little difference in the anogenital distance between the sow and boar. The sow has an anogenital region that looks like a ‘‘Y’’. The arms of the Y are grooves that surround the urethral opening. The straight portion of the Y runs from the vaginal orifice to the anus. The vaginal orifice is closed by a membrane except during estrus and parturition [2,13–15]. The boar has an anogenital region that looks like a dotted ‘‘i’’ The dotted top is the prepucial opening that encloses the penis. The straight line separates the two scrotal pouches. The base of the ‘‘i’’ is the anus. Slight pressure around the area will usually cause the penis to be everted in the male. In adult males there are two obvious scrotal pouches with large testicles present at a relatively young age [2,13–15]. Guinea pigs have two mammary glands, and two nipples, located in the groin, present in both sexes [13]. Reproductive physiology Guinea pigs are induced polyestrus and can breed all year in the proper setting [2,13–16]. The estrous cycle lasts 15 to 17 days (range: 13–21 days) [2,13–16]. The estrus period itself lasts between 24 and 48 hours; however, the sow usually accepts the boar for 6 to 11 hours [2,13–17]. Spontaneous ovulation usually occurs about 10 hours after the onset of estrus [13]. There is a fertile postpartum estrus in most sows between 2 and 25 hours after parturition. Sows bred at this postpartum estrus have a 60% to 80% pregnancy rate. Behavioral signs of estrus may include lordosis, mounting other sows, accepting a boar mounting, swollen external genitalia, and perforation or thinning of the vaginal membrane [2,13–16]. Once mating has occurred, the vaginal canal is sealed with a plug of ejaculate called a copulatory plug. Rodent copulatory plugs are very hard and waxy. Their function is not known, but it has been postulated that they store sperm, prevent sperm leakage, induce pseudopregnancy, affect sperm transport, or prevent fertilization of the female by subsequent males [14]. The copulatory plug remains in place up to 48 hours and then drops out. By this time, the vaginal membrane has resealed and will not open again until parturition [2,13,14]. The guinea pig placenta is similar to that of humans (hemochorial), with the trophoblasts in contact with maternal blood [2,5,14]. Gestation ranges from 59 to 72 days (average: 68 days). If the litter is large, the gestation is likely to be shorter with smaller pups. If the litter is small, the gestation is longer with larger pups and there is a greater possibility of dystocia [2,13–16]. First-time mother sows that are extremely young or elderly are more likely to have only one pup or smaller litters. Postpartum estrus conceptions are more likely to yield large litters [24]. Superfetation is a phenomenon involving a sow carrying litters from two separate fertilizations (this will be discussed later in the article) [13,15]. Sows may be palpated as early as

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15 days postcopulation for evidence of fetuses within the uterus [2,14,15]. They will be smaller than the kidneys but larger than fecal pellets (ie, marble-sized). An ultrasound can be performed approximately 20 days and beyond to confirm pregnancy and determine viability of the pups if palpation is inconclusive. Fetal movements may be felt as early as 42 days gestation [13]. A radiograph can be taken after the skeletons have calcified in approximately the last trimester of the pregnancy (>45–50 days gestation). False pregnancies can occur and usually last about 17 days [13]. Guinea pigs do not build nests and give very few signs of impending birth. The fibrocartilagenous pubic symphysis can separate dramatically under the influence of the hormone relaxin [2,14,15]. When the pubic symphysis has separated by 2.5 cm (approximately a two-finger width), parturition should follow within 24 hours. Sows that are older than 7 months when bred for the first time or those on poor diets may have a symphysis that cannot separate enough to allow passage of the fetuses; in these cases a C-section will be necessary [2,14]. Normal parturition is very rapid, with pups being delivered within minutes of each other [2,13–16]. I have observed pups coming out of the birth canal so close to one another that the sow cannot get the membranes off of the faces quick enough; this commonly results in suffocation. It is advisable to leave the boar or another sow in the cage with the pregnant sow to help her with the cleaning of the babies. The sow will eat the placental membranes and clean up the babies after the birthing. Newborn guinea pigs are very precocious and are soon running around the cage and nibbling at the food. They are born fully furred, with their eyes open and their teeth erupted [2,13–15]. The sow may not produce enough milk or may neglect her babies; however, these sows are frequently successful with their next litter. If the sow is neglecting her pups, she should be placed in a small cage so that she cannot run away from the pups. If she doesn’t have milk, an attempt should be made to foster the pups to a sow that is nursing. Many substitutes have been used for orphan pups that cannot be fostered to another sow. Kitten milk replacers, puppy milk replacers, and goat’s milk have all been successful. I have discovered that regardless of the supplement choice, a sudden change to another supplement should be avoided, because this can lead to severe diarrhea and death. Because guinea pig pups are so precocious, they are usually eating food within 3 days of birth [2,14]. Some breeders have had good success with placing wheat bread soaked with replacement formula in a shallow dish. The orphan pups usually learn quickly how to eat the soaked bread and drink out of a water bottle. They should be supplemented with vitamin C if they are not nursing and are not yet eating pellets that have vitamin C added to them. Weaning can be a stressful time for orphan pups. As they switch over from primarily foregut liquid digestion (milk) to hindgut, cellulose digestion, many problems can occur. Normally the pups will eat some of the sow’s feces, which may help innoculate their digestive tract with the symbiotic organisms necessary for cellulose digestion. Pups that have been raised

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as orphans or that have heavy parasite loads may develop severe diarrhea and die. I recommend using a probiotic or fecal slurry from a healthy adult guinea pig when handfeeding the babies. Guinea pig milk is approximately 4% fat, 8% protein, and 3% lactose (carbohydrate). Even sows with plenty of milk production tend to be passive nursers; they sit up, allowing the pups to get underneath them to nurse. They do not seem to notice if one pup is not getting enough milk, so the smaller, weaker pups suffer. If two sows and their litters are housed together, the pups will frequently nurse off of either sow, which can be to the detriment of the younger pups. Guinea pig pups naturally wean between 3 to 4 weeks of age. Of interest is the finding that males make better breeders as adults if they remain with their mother and siblings until they are at least 30 days of age [2]. Reproductive surgery There are many factors that could lead to the presentation of a sow or boar for reproductive surgery. These may include the prevention of pregnancy, prevention of reproductive disease, treatment of behavioral problems, and neoplasia [5]. The surgical procedure may be similar to other small mammals; however, certain special precautions are warranted. Unlike canine and feline patients, rodents cannot vomit [6]. It is not necessary to fast a guinea pig surgical patient overnight. Fasting them for a few hours before surgery may decrease the stomach volume and food packed in the mouth. It is not recommended to withhold water from the patient before surgery [7,8]. I recommend allowing guinea pigs to graze up until a couple of hours before surgery, then encouraging them to eat as soon after surgery as possible [6]; this may help maintain normal intestinal function. Part of the preanesthetic preparation should include cleaning out the mouth to remove food particles and excessive saliva. Use of anti-inflammatory agents and analgesics can help before and after surgery. Controlling stress and pain will allow the patient to recover more quickly and return to a normal appetite and activity level [6]. Adhesions, peritonitis, and ileus are potential complications in any guinea pig abdominal surgery. Adhesions may be minimized by careful tissue handling, keeping tissues moist and careful choice of suture materials. Use of antiinflammatory drugs such as flunixine (Banamine) or calcium channel–blocking agents such as verapamil (Calan) postoperatively have shown promise [7]. Peritonitis is a very serious concern and potential complication with any abdominal surgery. Attention to sterility and lavage of the abdomen with warm, sterile saline can be very successful. Even gentle manipulation of the gastrointestinal tract while performing reproductive surgery can lead to postsurgical ileus. Motility-stimulating agents such as cisapride (Propulsid) or metoclopramide (Reglan) may be necessary. To minimize adhesions, it is best to use the least reactive suture materials. I prefer the monofilament synthetic absorbables (eg, PDS). Stainless metal clips for hemostasis are also

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very useful (Hemoclips, Dilling Weck, Research Triangle Park, NC) [7]. In my experience, catgut causes tremendous tissue reaction and can lead to many secondary complications. Anesthesia and surgical preparation The most important piece of anesthetic equipment for an exotic animal veterinarian is an isoflurane anesthetic machine [8]. Isoflurane allows for quick induction and recovery. Many potential protocols are discussed in the rabbit section under anesthesia and surgical preparation. These apply to guinea pigs as well as rabbits. When using a mask for isoflurane administration, it is much less stressful for the guinea pig if pure oxygen is delivered for a few minutes, followed by a gradual increase in the percentage of anesthetic gas until the righting reflex is gone. Anticholinergics such as atropine or glycopyrolate can be used to help decrease the salivary secretions and are easily incorporated into the preanesthetic protocols. As mentioned previously, I recommend thoroughly swabbing out the mouth before using the face mask. Once the animal is anesthetized I usually examine the mouth further to identify any potential problems before proceeding [6]. Artificial tear ointment is applied to the eyes and the surgical site is clipped. I usually apply gentle pressure to the lower abdomen to facilitate emptying of the bladder; too much pressure should be avoided, because this could damage the bladder or delicate intestinal tissue. Surgical scrub and preparation can be performed using the same methods used for dogs and cats. Very small rodents are more susceptible to heat loss because of their greater surface area to mass ratio [30]. In the smaller patients I do not use alcohol because of its supercooling effect; instead, I use a chlorhexidine solution that is diluted with warm water after the initial chlorhexidine scrub preparation. Although guinea pigs are not as fragile as rabbits when it comes to positioning, care should be taken to avoid tying their legs down too tightly. Rabbits and rodents have a comparatively small thoracic cavity [1,8]. I recommend elevating the chest so that the abdominal organs do not put pressure on the diaphragm, which in turn makes it difficult for the lungs to expand. This is the opposite technique of that used frequently in dogs and cats [6]. Some surgical tables slant cranially to shift the liver and gastrointestinal organs forward and away from the site of reproductive surgery; however, this type of positioning could be fatal in rodents. I have also learned not to recover rabbits and rodents before the procedure is completed. Because rabbits and rodents are prey species and can go into shock quickly, I use an analgesic presurgically and again postsurgically [8]. Ovariohysterectomy (spaying) After the guinea pig has been anesthetized, positioned in dorsal recumbency, and the surgical area prepared for surgery you may proceed. A ventral, midline incision is made starting just caudal to the umbilicus and extending caudally 1/2 to 2/3 of the way to the pubic bones (1–2 inches). The

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skin in this area is usually very delicate; therefore, not much digital pressure is required on the scalpel. I use small, rat-toothed forceps to lift up the linea alba before making my stab incision into the abdominal cavity. The large cecum, bladder, stomach, or intestines are frequently lying just beneath the thin muscle layer and can be easily nicked if too aggressive an approach is taken. Once inside the abdominal cavity, I do not use any instruments with sharp points to prevent inadvertent puncture of gastrointestinal organs when working around them. The uterus is sometimes immediately visible; however, in other cases a snook hook (spay hook) may be needed. Care should be taken to avoid mistaking a portion of the small intestine or colon for the uterine horns. Once the uterus has been identified, the oviduct, infundibulum, and ovary can be found on each side [16]. The ovary is frequently surrounded by fat and may be difficult to see initially. Elevating the ovary away from the kidney is often difficult, and caution must be used to prevent tearing the delicate ligament and the associated ovarian vessels. It can also be challenging to ligate the ovarian pedicle without incorporating intestinal contents into the ligature. The importance of reducing any handling of the gastrointestinal tract while performing the rabbit spay was mentioned earlier, and it is equally important to be gentle and avoid handling the intestines in the guinea pig; however, it may be impossible to see and ligate the ovaries and uterine horns without pushing aside or even pulling some of the abdominal contents out of the abdominal cavity. When it is necessary to remove organs to gain access to the ovaries, I recommend laparotomy sponges that are soaked with warm, sterile saline to keep the organs moist and protected. Ovarian cysts and tumors as well as uterine masses can make this procedure even more challenging. Ligation of the ovarian pedicle with suture material or hemoclips should be performed, followed by gentle retraction of the infundibulum and oviduct. Gently tear apart the ligaments to release the oviduct. There will be varying amounts and sizes of vessels within these ligaments depending on the age, health, and breeding status of the sow. Any significantly sized vessels should be ligated or cauterized as the procedure progresses. The uterine vessels should be double-ligated either separately from the uterus or transfixed to the uterus. When ligatures are placed, caution must be used to prevent incorporating intestinal serosa in with the reproductive tract. I usually lavage the abdomen with a small amount of warm, sterile, saline solution before closing the muscle layer. The abdomen is then closed in a simple interrupted pattern, again, being cautious not to inadvertently incorporate intestinal tract into the closure. This can be a real challenge because the cecum, in particular, continues to poke out of the incision while the abdomen is being closed. I will use the flat handle of the spay hook to help keep the intestines in the abdomen while I am stitching it closed. The skin can be closed using subcuticular patterns, tissue glue, skin staples or a combination of the three. Rodents are also very effective at removing more traditional skin sutures and can occasionally remove the skin staples as well.

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Orchidectomy (castration) The reproductive anatomy of a male guinea pig is similar to that of dogs and cats, although the testicle is more elongated and the inguinal ring remains open throughout their life; this allows the testicles to be withdrawn into the abdominal cavity [1,2]. After the guinea pig has been anesthetized and positioned in dorsal recumbency and the surgical area has been prepared for surgery, the procedure can begin. There are primarily two different approaches to castration of the boar [13,14]. Some surgeons make separate incisions in the ventral surface of the scrotum and exteriorize the testicles and epididymis from each incision, which is similar to the technique performed in most cats. I prefer a prescrotal incision separately over each cranial scrotal region on either side of the body of the penis. Regardless of the incision technique, the entire testicle and epididymis must be exteriorized to expose the spermatic cord with the vessels and vas deferens for ligation. The spermatic cord can be ligated as one unit in a closed fashion using suture material, hemoclips, or the overhand knot (autoligation technique) [13,14]. I prefer to use an open technique and double ligation of the spermatic vessels and vas deferens. This allows me to see the vessels, which can be hidden in the huge fat pad next to the spermatic cord. I can then ligate the tunic and close the inguinal ring at the same time. The possibility of umbilical herniation and intestinal strangulation is a concern after castration. Intestinal herniation and strangulation are prevented in the intact boar by the large amount of fat present around the spermatic cord [14]. I strongly recommend closing the neuter site rather than leaving it open as is commonly done in cats. A guinea pig sits and walks low to the ground which would allow bedding and feces to gain access to the surgical site. I prefer to close the skin in a subcuticular pattern and use tissue glue over the top for added insurance. In very large boars I will also use skin staples if needed. If the guinea pig has diarrhea, impaction of the rectal pouch or chews the incision open an infection is still possible. Cesarian section (C-section) The small animal veterinarian may be called upon to perform a C-section if the sow is having trouble giving birth naturally. This could be due to fetuses that are too large, incomplete separation of the pubic symphysis, or uterine inertia that is not responding to medical treatment. The procedure used can be similar to the procedure that is performed on cats and dogs. The exception would be if a spay is to be performed at the same time. If the sow is to be spayed, the precautions listed in the previous section on ovariohysterectomy should be observed [5]. It is easier to spay a pregnant sow than it is to spay a nonpregnant sow because the heavy, gravid uterus has already pushed the gastrointestinal tract away from the ventral abdomen. It is very important to elevate the thoracic cavity during surgery so that the distended abdomen does not put too much pressure on the lungs. I recommend pain medication for several days postoperatively so that the sow is more likely

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to eat and allow nursing. An alternative would be to foster the babies to another nursing sow, if possible. Reproductive diseases Infectious reproductive disorders Pyometra, vaginitis, and metritis can occur in breeding and nonbreeding sows. Signs may include vaginal discharge, impacted vaginal area, abdominal swelling, depression, anorexia, fever, polydipsia, and hypothermia. History will frequently include recent breeding or birth, infertility, and stillbirths [10]; however, sows that have never been bred can still develop reproductive infections. Certain types of bedding may predispose sows to vaginitis; these include wood chips, sawdust, or other types that can become impacted into the vaginal area. Diagnosis may include palpation of an enlarged, doughy uterus; a vaginal area impacted with bedding material; radiographic evidence of uterine enlargement; ultrasound confirmation of a fluid filled uterus; or leukocytosis with neutrophilia [5,10]. Caution should be used when palpating sows with suspected uterine disease, because the uterine tissue can be extremely friable and rupture is possible [5,10]. Exploratory surgery with ovariohysterectomy is necessary for a definitive diagnosis and treatment of pyometra. Antibiotic therapy and intravenous fluid therapy should be initiated before anesthesia and surgery. A broad spectrum antibiotic can be chosen while a culture and sensitivity are pending. The two most commonly isolated organisms are Bordatella bronchiseptica and hemolytic Streptococcus spp [2,13,14]. The most common sources of infection are transmission during mating, contaminated or irritating bedding material, and hematogenous spread [2,14]. Other possible pathogens include Escherichia coli, Corynebacterium pyogenes, Salmonella, Staphylococcus, and Streptococcus spp [2,13,14]. The protozoal parasite Toxoplasma gondii has also been reported to infect guinea pigs and cause abortion. The cat is the definitive host, and the guinea pig becomes infected by ingesting feed or grass that is contaminated with cat feces containing the oocyst stage of the parasite [13]. As with other sexually transmitted diseases, infectious reproductive disorders are easier to prevent than to treat. I recommend using virgin sows and boars when introducing outside lines into a caviary [5]. Cleanliness and careful choice of bedding material cannot be overemphasized as effective ways to prevent these infections. Having a clean cage in which the sow can deliver her pups will help prevent postpartum infections. Treatment of vaginitis consists of thorough examination of the vagina for foreign matter, gentle cleansing of the area using a dilute chlorhexidine solution, and therapy with a broad-spectrum, systemic antibiotic while a culture and sensitivity is pending. Antibiotics that have been useful in such cases include trimethoprim/sulfas, enrofloxacin, chloramphenicol, and tetracyclines [5,14]. Vaginoscopy is more easily accomplished during estrus, when the vaginal membrane is relaxed [2,13,14].

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Orchiitis and epididymitis can occur in breeding and pet boars. Signs may include fever, anorexia, weight loss, swelling of the scrotal area, and prepucial discharge. The cause may be sexual transmission from a sow, fight wounds from another boar, or hematogenous spread [2,5,13,14]. Treatment usually involves systemic antibiotics and sometimes requires castration. Bordatella and hemolytic Streptococcus spp are frequently involved pathogens. Many of the same organisms that cause reproductive infections in sows can also lead to infection in the boar. Clinical signs vary but may include areas of redness and swelling in the perineal area with prepucial discharge. The disease may resolve, but those that recover may become carriers. Diagnosis requires culture and sensitivity. Scrotal plugs may become secondarily infected and may be treated similar to sows with vaginitis. Scrotal plugs will be discussed under noninfectious reproductive diseases. Prevention of Bordatella infection can be achieved using an injectable autogenous, formalin-killed, bacterin (Bronchine, Biocor Animal Health Inc, Omaha, Nebraska). The vaccine protocol involves inoculating the guinea pig with 0.2 ml intramuscularly and repeating 2 to 3 weeks later. The vaccine is approved for subcutaneous injection, however, the only research available on the efficacy of this vaccine was done using the intramuscular method of administration [2,5,13]. There are conflicting reports regarding the effectiveness of intranasal administration (ie, whether or not it is helpful or harmful to the guinea pig) [2,13,14]. I advise against intranasal administration until further research has been conducted. A booster vaccination is recommended every 6 months for guinea pigs that would be at risk of continued exposure [2,13,14]. Several veterinarians have reported success of vaccination during outbreaks and apparent elimination of the carrier state [2,5,13]. Noninfectious reproductive disorders Noninfectious reproductive disorders can result in reduced fertility, resorption, abortion, dystocia, stillbirths, agalactia, and pregnancy toxemia [2,5,13–15]. Reduced fertility or infertility can be caused by the infectious agents mentioned previously or noninfectious causes such as inadequate nutrition (protein deficiency, hypovitaminosis A, C, D, E or hypervitaminosis A), environmental stress, hyperthermia, behavioral problems, and congenital anomalies. Vitamin E deficiency causes myodystrophy; a highserum creatine phosphokinase can support this diagnosis [10]. Fetal death in the first trimester of gestation usually results in resorption and may be mistaken for an unsuccessful breeding [2,13–15]. Because the causes of infertility may be different from those of resorption, it may be important to differentiate between the two. If this is occurring on a large scale in a caviary, evaluation of diet and environment as well as screening for infectious diseases is warranted. Feed designed for small rodents is not sufficient for guinea pigs [2]. In addition, rabbit formulations are insufficient for guinea pig reproduction. Rabbit feed contains no ascorbic acid (vitamin C) and has

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excessive levels of vitamin D (too much for guinea pigs) [2]. Guinea pigs also have higher requirements for folic acid than do rabbits or other rodents [2]. If a sow has two consecutive pregnancies that do not produce live births, I recommend an ultrasound to confirm a true pregnancy the next time she is bred; if she doesn’t deliver, it may be a case of resorption rather than infertility. If a prize boar suddenly stops having successful litters, his sperm could be evaluated and any sows bred to him could be given an ultrasound to determine if resorption is occurring. This could indicate a sexually transmitted disease from the boar to the sows, which can lead to unsuccessful reproduction. Fetal death after the first trimester usually results in abortion. The aborted fetus should be submitted for bacterial culture and histopathology to determine the cause of the abortion. Stress, trauma, hyperthermia, toxic plants, drugs, or severe dietary deficiencies can also lead to abortions [2,5,13–15]. In my experience, secondary uterine infections are more likely to occur in cases of abortion than in resorption. Behavioral or psychological reasons for infertility may also be involved in guinea pigs. Dominant adult sows may not allow more submissive boars to mount them. Males that are not allowed to mate until they are over 1 year of age may have decreased libido, leading to functional infertility [13]. This is why it is advisable to place young boars into breeding at about 4 months of age [13]. Some researchers have found that boars make better breeders if they are left with their mother and siblings until they are at least 30 days of age [2]. I have witnessed several prize boars that have become infertile secondary to heat stroke or fever due to some infectious process. On a few occasions, fertility returned after several months of recovery; in others, fertility never returned. Premature births can occur when sows are housed together. A sow may go into labor prematurely soon after her cagemate has her litter. Many theories have been proposed by lay people and veterinarians as to the cause of this problem. There could be a pheromone released at parturition that stimulates the second sow to go into labor. The pups from the first litter could nurse on the pregnant sow and stimulate the release of oxytocin, which could lead to premature labor. Some breeders suggest that premature birth occurs if the pregnant sow eats the placenta from the first sow. The pups born prematurely will rarely survive. Sows that have ovarian cysts may become infertile (ovarian cysts will be discussed as a separate condition). Dystocia is more common in guinea pigs than in rabbits or other rodents. Causes may include oversized kits, narrow pelvic canal, inability of the pubic symphysis to separate, uterine torsion, obesity [25], nutritional deficiencies, or uterine inertia [2,5,13–15]. Because the guinea pig pups are so precocious and large at birth and the pubic symphysis must separate (usually 2.5 cm) it is not uncommon to have dystocia problems in guinea pigs that are older when bred for the first time. Most veterinarians agree that if the sow is older than 8 to 12 months when bred for the first time it can

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be very dangerous and likely to end with dystocia [2,5,13–15]; however, many guinea pig breeders disagree with this statement, claiming that it is just an old wives’ tale. It is possible that hypovitaminosis C may also play a role in inadequate pubic symphysis separation. Whatever the cause may be, it is very serious and must be monitored closely. Contractions and straining that produce bloody or greenish discharge but no pups may indicate dystocia. Manual assistance with removal from the vaginal canal may be necessary in these instances [2,5,13–15]. Application of a lubricant (eg, a water-based jelly) may help the passage of the pups. In cases of suspected uterine inertia, calcium should be administered first, followed by oxytocin if necessary. Calcium gluconate can be administered either orally or by injection, and oxytocin can be administered by injection SC or IM (dosages are outlined in Box 1). The sow should be placed in a quiet, stress-free environment while waiting to see if treatment is successful. A C-section may be necessary if medical treatment is unsuccessful. Pregnancy toxemia is commonly seen in the breeding guinea pig. It is most common in the last 2 weeks of gestation and the first 1 to 2 weeks post partum [31,32]. It has been my experience that there are actually two forms of pregnancy toxemia. Guinea pig pregnancy toxemia (ketosis) is very similar to that observed in sheep (twin lamb disease) [2,5,13–16]. The other form of pregnancy toxemia may be similar to that observed in pregnant women [2,5,16]. In the latter type of toxemia, the heavy, gravid uterus may compress its own vascular supply or that of the kidneys or gastrointestinal vessels, leading to ischemia of tissues and hypertension. This may lead to the release of thromboplastin and initiate disseminated intravascular coagulation (DIC) [5,14–16]. This may or may not explain another syndrome that is observed occasionally: hemorrhagic syndrome. Hemorrhagic syndrome occurs during or shortly after parturition and usually results in fatal hemorrhage [13,15,16]. Different proposed causes include vitamin K deficiency due to poor quality pellets or hay, pressure from the gravid uterus on the liver leading to congestion and dysfunction, calcium deficiency and its effect on the clotting cascade, or the initiation of DIC [2,5,13–16]. One source explains that the pressure of the gravid uterus upon the liver can cause bile blockage and hepatic dysfunction, resulting in a vitamin K-deficient hemorrhagic syndrome [2]. This can cause complications during C-sections and even following vaginal deliveries [2]. Pressure from the gravid uterus or difficult birthing can also lead to temporary paresis or paralysis of the rear legs. Sows will usually regain use of their hind limbs without any intervention as long as they are kept in a clean environment with easy access to food and water. The symptoms, prevention, and treatment of the two types of toxemia will be discussed separately. Pregnancy ketosis involves a sow that either stops eating or significantly reduces food and water intake. The sow is usually inactive, depressed, uncoordinated, and occasionally has dyspnea. The condition may progress to

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muscle spasms, paralysis, and death. The breath may smell of ketones, which is attributable to ketonemia. Other clinical and laboratory findings include ketonuria, proteinuria, aciduria, hypoglycemia, acidosis, hyperlipemia, and hyperkalemia [2,5,13–16]. Subclinical hypocalcemia may predispose a sow to either or both forms of toxemia; therefore, calcium supplementation is recommended in these cases. Hepatic lipidosis is commonly found on ultrasound or necropsy. The pathophysiology of this form of toxemia appears to involve a negative energy balance in the sow because of the heavy demand of her growing fetuses. She is unable to supply sufficient glucose either to the developing fetuses or to herself; therefore, she catabolizes her own body fat and protein stores. This occurs most often in obese sows and can even occur in obese boars that undergo anorexia or severe stress [2,5,13–16]. Death can occur acutely, or the guinea pig may linger and progressively deteriorate over several days [14,15]. Predisposing factors include obesity, change in diet, environment, lack of exercise, large fetal loads, heat stress, and primiparity [2,5,13–16]. Treatment may not be effective, so prevention is the key. Stress, obesity, and changes in diet or environment in late pregnancy should be avoided. Offering supplements to the regular diet that are high in carbohydrates in the last 2 weeks of gestation, calcium supplementation, and close observation may help. Exercise to lose weight before breeding may prevent toxemia, and placing the food dish and water on opposite sides of the cage in early pregnancy may encourage exercise in pregnant sows. Food and water should be made easily accessible in the last few days of gestation, however, because the fetal mass may weigh more than the mother at this time, making it difficult for her to move [2,14]. Once ketosis occurs, treatment may include intravenous or intraosseous fluids (LRS with dextrose), oral glucose, calcium gluconate (orally or by injection), magnesium sulfate, and short-acting corticosteriods (if the sow is in shock) [2,5,13–16]. Some veterinarians recommend 1 to 2 ml 50% dextrose in 3 to 5 ml saline IV or Interosseous (IO) [2]. Parenteral nutrition can be accomplished with syringe feeding, orogastric tube feeding, nasogastric tube placement or esophagostomy tube placement [5]. Formulations for assisted feeding are available (Oxbow Hay and Feed); otherwise children’s formulas may be used. Monitor blood gases for acid/base level, electrolytes, calcium, and phosphorus. The second form of pregnancy toxemia involves the heavy gravid uterus compressing blood vessels or nerves, which may lead to severe vascular and neurologic dysfunction [2,5,14,16]. If the blood pressure can be measured, it is helpful to determine whether hypertension (eg, compression of the renal vessels) or hypotension (shock) is involved. If hypertension is involved, emergency stabilization and a C-section are required [5]. Vasodilators may be indicated initially and close monitoring of blood pressure during and after surgery is necessary [5]. If hypotension is involved, then treatment for shock with short-acting corticosteroids, colloids (such as hetastarch), and intravenous or intraosseous fluid therapy are indicated [5]. Most likely a

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C-section is required in either case to save the sow. If ischemia occurs, the release of thromboplastin may initiate disseminated intravascular coagulation, which is usually a fatal condition. Administration of flunixin (Banamine) and heparin may combat the potential for the thromboplastin effect and DIC. I have used the same protocols that are published for dogs and cats successfully in a couple of cases [5]. I am not aware of any specific ways to prevent this form of toxemia; however, because postpartum breedings tend to produce the largest litters, they may predispose the sow to greater risk. Ovarian cysts are very common in reproducing and nonreproducing sows. They have been identified in 76% of female guinea pigs between 1.5 and 5 years of age [14]. They may occur in one or both ovaries, but they occur most commonly in the right ovary. The ovary may have single or multiple cysts that range in size from 0.5 to 7 cm (average: 3 cm) [13,14]. The cysts are filled with clear liquid, appear to develop spontaneously, and increase in size as the animal ages [13,14]. Clinical signs may include; decreased fertility, bilateral symmetrical alopecia of the flanks, abdominal swelling, pain induced anorexia, and enteritis (secondary ileus) [2,5,13,14]. Diagnosis may be made by abdominal palpation, radiography, or ultrasound. Treatment must include analgesics with or without motility stimulating agents. Ovariohysterectomy is the best way to prevent and treat ovarian cysts. If surgery is not an option or considered too risky, it is possible to attempt percutaneous drainage of the cysts; ultrasound-guided drainage would be the safest alternative. Superfetation is a phenomenon that has been recorded in guinea pigs. This occurs when the sow carries fetuses from two separate fertilizations, occurring at two different estrus cycles [13,15]. There are two known situations. The first and most common involves a second fertile mating at the next estrus (approximately 16 days after the first mating). The second situation is less common and involves a fertile mating two estrus cycles later producing a separate litter approximately 35 days after the first litter [13,15]. The uterus has two separate horns, and each litter is carried in a different horn [13]. In the first situation the pups are only about 16 days apart in age, and when labor is initiated both litters are usually born. The smaller, premature pups usually do not survive. In the second situation, it has been reported that two separate parturition events have occurred, and full-term pups were delivered each time with both litters surviving [13], though I have never witnessed this event personally. I have seen many examples of the litters that are 16 days apart and have not seen one premature pup survive. Scrotal plugs are common in mature boars [2,13–15]. They may accumulate a mass of sebaceous material in the area between the penis and anus, where folds of skin act to hold the material in. These can be confused with rectal impactions and they may sometimes have fecal material or bedding material packed into them [2,13–15]. Treatment involves gentle cleansing of the area with dilute soapy water (eg, chlorhexiderm scrub) and rinsing

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well. Prevention may involve changing the bedding material or neutering the boar. Breeding boars need to be checked for this condition on a regular basis as it may lead to infertility and secondary infections. Reproductive neoplasia Tumors of the reproductive tract are also reported in guinea pigs. Leiomyomas, leiomyosarcomas, cystic endometrial hyperplasia of the uterus, and fibroadenomas and adenocarcinomas of the mammary glands have all been reported [2,5,13–15]. Surgical removal and histopathology are the recommended treatment of choice. Ovariohysterectomy of the young sow should prevent all of the neoplasms mentioned above. Testicular tumors have been reported in boars as well [2,13,14]. Castration is the best form of prevention and is also the recommended treatment. Mastitis in guinea pigs is usually of an infectious nature; however, several husbandry problems can be involved. Several organisms have been reported, including Pasteurella spp, Klebsiella spp, E coli, Staphylococcus spp, Streptococcus spp, and Pseudomonas spp [2,13–15]. Some of the husbandry problems that may predispose a sow to mastitis include wet, dirty cages; sharp objects; wire cage bottoms; and trauma by pups [2,13–15]. Organisms may enter through the teat canal or through bite wound to the teats. Clinical signs may include swollen, inflamed glands that are warm; pain; anorexia; and refusal to let pups nurse. Later in the course of infection, the glands may become cool and cyanotic [2,5,13–15]. There may be a mucopurrulent discharge from the teat or the milk may be bloody. If the infection spreads and septicemia occurs, the sow and pups may die [14]. Culture and sensitivity of the discharge or milk is necessary to determine the causative organisms. Treatment can begin with a broad-spectrum antibiotic, such as trimethoprim/sulfa, enrofloxacin, chloramphenicol or aminoglycosides [5,14]. Use of nonsteroidal anti-inflammatory drugs (eg, flunixin) and analgesics (eg, butorphenol) may help improve the sow’s appetite and activity level. Other suggestions include hot-packing the glands, keeping the environment clean, and weaning the pups early. If the glands do not improve, they can be surgically resected [2,5,13–15].

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