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Surgical Techniques in Small Exotic Animals
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Surgery of Pet Ferrets
Neal L. Beeber
In recent years, the domestic ferret has had a dramatic increase in popularity. In 1990, the number of pet ferrets in the United States was estimated to be more than 7 million.1 As these animals have increased in Popularity, they have become more common in veterinary practices. This discussion deals with some of the more common surgical procedures in ferrets.
Preparation and Fasting
Healthy ferrets make excellent surgical candidates, are hardy, and with attention to certain parameters they do not present any unusual anesthetic risks. The intestinal tract is short, resulting in a gastrointestinal transit time of 3 to 4 hours.2 For this reason, patients are only fasted for 4 to 5 hours before surgery, except in the case of insulinoma resection, for which the fast is 3 hours.
Sedation and Anesthesia
Isoflurane is the anesthetic of choice; however, halothane can also be used, except in critically ill patients. A nonrebreathing system is used with a flow rate of 0.6 to 1.0 L per minute. No premedication is required. In many cases, ferrets can be masked until they are sufficiently anesthetized to allow endotracheal intubation. This can usually be accomplished with a flow rate of 2 L and a 4 to 5% isoflurane concentration. The animal relaxes in 2 to 5 minutes. Because struggling or excitement is minimal, chamber induction is not usually necessary. Maintenance level of isoflurane is 1.75 to 2.5%. It is often necessary to use a small amount of lidocaine (0.1 mL) to paralyze the larynx to accomplish intubation, as in the feline species. All ferrets are intubated except for the most minor procedures. Use of 1.5 to 4.5 French endotracheal tubes is sufficient for most ferrets. If the tubes are allowed to become cold in a refrigerator, they will become stiff and more easily introduced into the trachea. Because ferrets vary in body size, several tube sizes should be available. Some breeding establishments have been importing European ferrets, which are generally larger than the American breeds and commonly weigh up to 5 to 6 lb. These ferrets need slightly larger endotracheal tubes. Whenever possible, a cuffed tube is recommended.
Ketamine and diazepam ketamine combinations can be used as a preanesthetic, for intubation, or for short procedures intra-muscularly at a dose of 10 to 20 mg/kg for ketamine and 1 to 2 mg/kg for diazepam. In my opinion, acepromazine should not be used in ferrets because of this agent’s vasodilatative properties and the possibility of heat loss. Ferrets should be placed on a warm water recirculating system to prevent heat loss, and any intravenous fluids to be administered should be warmed to 85 to 90° F. The patient’s rectal temperature should be monitored during the surgical procedure. A simple and inexpensive way to accomplish this is to use a digital outdoor thermometer available commercially for under $15 (indoor outdoor thermometer, Radio Shack catalog No. 63 854). The probe can be inserted directly into the rectum or attached to a red rubber catheter as a stylet.
Except for a routine spay, neuter, or other minor procedure, a 24 gauge intravenous catheter (Baxter Quickcath 24 gauge 1.6 cm) should be placed for all surgical procedures. The cephalic vein is the most common site for placement, but lateral saphenous, jugular, and intraosseous catheters can also be used. When a jugular catheter is necessary, a 24 gauge cephalic catheter can be placed in the jugular vein. The types of fluids administered depend on the type of surgical procedure performed and are discussed under the appropriate section. Ferrets are monitored with a pulse oximeter, which works well in this species. Recovery time depends on the animal’s condition and length of anesthesia; however, when isoflurane is used alone, recovery is remarkably fast and smooth.
General Surgical Considerations
Ferret skin is tougher than dog or cat skin, so slightly more pressure may need to be exerted. One often sees a thick subcutaneous fat layer, which should be dissected bluntly. The linea alba is readily apparent. A stab incision should be made into the abdominal cavity and extended. Care should be taken to avoid the spleen because it is often large in this species. Most common types of sutures can be used depending on the operation performed. I prefer to close the abdomen with 4-0 polydioxanone (PDS), polypropylene (Prolene), or nylon. Most nonabsorbable suture material with a cutting needle can be used for skin sutures. Ferrets rarely chew external sutures.
Most ferrets sold as pets in the United States are neutered before 6 or 7 weeks of age, so ovariohysterectomy is not a common procedure, as in dogs and cats. Ferrets should be spayed by 6 months of age, however, if they are not to be used for breeding. Ferrets are induced ovulators. If they are allowed to remain in estrus, potentially fatal bone marrow suppression may result from estrogen toxicity. Medical treatments to terminate estrus are available;3 however, spaying is recommended.
The surgical procedure is similar to that for cats. The ferret is placed in dorsal recumbency, and the abdomen is shaved and prepared. A 3 to 4 cm midline incision is made 1cm posterior to the umbilicus. Blunt dissection is used to dissect through the fat layer and subcutaneous tissue to expose the linea alba. An incision is made through the linea and is extended. Usually, a layer of fat is encountered. The uterus of ferrets is bicornate, as in cats. The uterus can be elevated by using a spay hook, or sometimes it can be seen lying just under the incision by bluntly moving the fat. The uterus of the ferret is not nearly as friable as that of the rabbit. Ferrets have a high degree of body fat, and the ovarian tissue and vessels may be obscured. The surgeon must be certain to ligate the ovarian vessels completely using 2-0 or 3-0 gut. The uterus is easily exteriorized and the suspensory ligament is readily torn. The uterus is ligated with gut and is removed. The abdomen can be closed with any of several sUi types in a simple interrupted pattern using 4 U nlonl filament absorbable or nonabsorbable material. The same suture or gut can be used for the subcutaneous or subcuticular layer. The skin can be closed with 3-0 or 4-0 nylon. Chewing of sutures has not been a problem. If the surgical procedure is performed in the morning, the ferret is released the same day. Postoperative antibiotics are not necessary. Skin sutures are removed in 7 to 10 days.
Like ovariohysterectomy, orchiectomy (castration) is usually done in young ferrets before they are sold to pet stores. For this reason, the average practitioner is not called on to perform this operation routinely. If an intact male ferret is presented, the owners should be encouraged to have the ferret castrated. In some cases, intact male ferrets are more aggressive, especially if intact females are nearby. The main objection to intact males is the heavy musky odor they produce. Many times, castration alone is enough to control odor, making descenting unnecessary. Testicular tumors have been reported, but a true incidence is difficult to estimate because most domestic ferrets arc neutered.4,5
Castration in the ferret is similar to castration in the dog. The ferret is placed in dorsal recumbency, and the prescrotal area is shaved. One prescrotal incision is made through which both testicles may be exteriorized. An open or closed method can be used. The spermatic cord and vessels are ligated with 4-0 gut, are iricised, and allowed to retract into the incision. The subcutaneous tissue is closed with gut, and the skin is closed with 4-0 nonabsorbable suture, which is removed in 7 to 10 days. Chewing the sutures has not been a problem. Alternatively, two incisions can be made in the scrotum, and the vessels can be clamped and ligated with 4-0 chromic gut. With this method, the scrotal incisions are not closed, similar to the procedure in cats. The ferrets are released the same day.
Adrenal tumors are among the most common neoplasms of ferrets. In our practice, adrenalectomy is the single most common surgical procedure performed in these animals, followed by insulinoma resection. In a retrospective study performed at our hospital and the Animal Medical Center from 1987 to 1991, the following types and frequency of biopsy results were recorded: adrenocortical adenoma, 64%; nodular adreno cortical hyperplasia, 26%; and adrenocortical carcinoma, 10%. In the patients with adreno cortical carcinoma, no gross or microscopic evidence of metastasis was seen. In addition, 70% of the cases occurred in females.6 Since this study, I have had the opportunity to operate on many more cases and have found that the biopsy percentage has shifted to adrenocortical adenoma. Hyperplasia now accounts for 95% of the cases, and the ratio of males to females has equalized. In addition, the earlier study found that 64% of ferrets had disease of the left adrenal gland, 20% had disease of the right gland, and 16% had bilateral disease. In the years after the study, my colleagues and I have seen left sided disease in 75% of patients, right sided disease in 15%, and bilateral disease in 10%.
Clinical signs, in order of decreasing frequency, are vulvar swelling, alopecia, pruritus, polydipsia, and polyuria. The diagnosis is based on clinical signs and abdominal ultrasonography. The accuracy of the ultrasound diagnosis depends on the experience of the ultrasonagrapher. Recently, a study indicated that the concentrations of certain plasma steroid hormones can be used as a marker for the disease.7 Even though the clinical signs may indicate the presence of an adrenal tumor, the clinician should obtain a presurgical ultrasound study whenever possible. This examination helps to rule out other causes of the clinical signs and indicates which adrenal gland is diseased. This distinction becomes important because removal of the right gland is technically more difficult, owing to its location under the caudate liver lobe and its proximity to the vena cava. The differential diagnosis of adrenal gland disease includes ovarian remnants, an intact female reproductive tract, pheochromocytoma, seasonal hair loss of ferrets, nutritional deficiencies,8,9 mycosis fungoides,10 and infestation by external parasites. I have also seen a ferret with cutaneous Malassezia pachydermatis infection that caused generalized hair loss. Adrenal disease in ferrets is not the same as Cushing’s disease because the clinical signs and pathologic changes are not caused by an increase in plasma cortisol concentration.
After the diagnosis is made, a complete blood screen and chemistry panel should be evaluated for each patient. Any abnormalities should be investigated and treated preoperatively. One of the most common abnormalities is hypoglycemia because many ferrets concurrently have insulin secreting tumors of the pancreas. Because these islet cell tumors are generally malignant, the prognosis should be discussed with the owners before proceeding. In addition, an in hospital blood glucose determination should be made immediately before anesthesia is induced, to make certain the blood sugar is still normal after the presurgical fasting period. Another important presurgical consideration is the possibility of underlying cardiac disease. Both hypertrophic and dilated forms of cardiomyopathy are seen in ferrets.11 At this time, clients are advised that a presurgical echocardiogram should be performed if possible. If this is not feasible, chest radiographs and careful cardiac auscultation should be performed. Every patient undergoing adrenalectomy receives an intravenous catheter. Various fluid types may be used; however, if one has any question about the presence of an insulinoma, 5% dextrose is the fluid of choice. Each patient receives a presurgical injection of antibiotics.
The ferret is placed in dorsal recumbency, and the abdomen is shaved from the area of the xiphoid cartilage to the inguinal area. An incision is made starting 1 to 2 cm from the xiphoid and extending 4 to 5 cm caudally. After dissecting through the fat and subcuta¬neous tissue, a stab incision is made in the linea alba and is extended with scissors. A self retaining Gelpi retractor should be used for good exposure. As in other species, a complete abdominal exploratory operation should be performed. It is especially important to check the pancreas for the possibility of insulinoma nodules (see later). In addition, all male ferrets should be examined for the presence of paraurethral cysts (discussed later). The surgeon generally must retract the spleen and intestines toward the right side of the ferret’s body. A laparotomy pad soaked in warm saline can be used to hold structures away from the surgical site. Alternately, the spleen and small intestines can be exteriorized through the incision and placed to the right. This maneuver pulls the mesentery away from the area of the adrenal gland and affords excellent exposure. Any exteriorized tissues should be covered with a warm moist lap pad to prevent tissue drying. The left adrenal gland is located just medial and proximal to the left kidney. This gland is located within a fat pad, and if diseased, it is usually irregular in shape and readily seen. In some cases, one sees a brownish-yellow discoloration. Digital palpation reveals the presence of borders on the mass. The dissection is begun on the medial side of the gland through the fat layer using Mayo scissors and is continued bluntly with mosquito forceps and sterile cotton tipped applicators. The gland is gently elevated as the dissection is continued. The small blood vessels in the fat generally do not have to be ligated. The adrenolumbar vein runs laterally and caudally from the ventral surface of the adrenal gland. It can be seen as the gland is elevated. This vessel is ligated using 4 0 chromic gut or a surgical clip (Hemoclip). The gland is continually elevated and dissected until a suture can be placed below it. The tissue is then incised, and the gland is removed. Closure is the same as for an ovariohysterectomy. Because this is a major abdominal procedure, patients are hospitalized for 1 to 2 days postoperatively. Amoxicillin oral suspension at a dose of 10 mg/lb is dispensed for 7 days.
As mentioned previously, removal of the right adrenal gland is a technically more difficult procedure. After entry into the abdominal cavity, and a general exploratory operation, the spleen and intestines are moved to the left or are exteriorized. The right adrenal gland is located under the caudate liver lobe. The hepatorenal ligament must be incised to elevate the tip of the liver lobe. The lobe is then reflected cranially. The adrenal gland is usually directly adhered to the vena cava (Figure 45-1). One must be careful to avoid lacerating this major vessel. The surgeon begins shelling out the gland by sharp dissection of the surface furthest from the vena cava and continues around the gland using iris scissors, mosquito forceps, and sterile cotton swabs. When the gland is mostly peeled away, a Hemoclip or a ligature using 5-0 absorbable suture is placed between the gland and the vena cava. Any remaining glandular tissue is trimmed using the iris scissors. One should have available 5-0 and 7-0 suture as well as sterile sponges (Gelfoam) in the event that the vena cava is lacerated. When one is certain that all hemorrhage has been controlled, closure is as described earlier.
Bilateral Adrenal Disease
When both adrenal glands are abnormal, the surgeon removes the left entirely and debulks the right. If incised, the adrenal gland bleeds profusely. One begins Caudal venacava dissecting the gland and places a crushing suture around the part that has been freed, using 4-0 monofilament absorbable or nonabsorbable material. Iris scissors can then be used to cut above the suture. The surgeon removes 50 to 75% of the right adrenal tissue.
The most common complication of adrenalectomy is prolonged or difficult recovery resulting from hypogly cemia secondary to an undiagnosed insulin secreting tumor of the pancreas. In fact, when ferrets are referred to my practice for postsurgical problems blood glucose concentrations are frequently found. For this reason, a blood glucose determination is performed before the surgical procedure and 1 to 2 hours postoperatively. Many times, fluids containing dextrose are used as a precaution. Ferrets are encouragcd to eat after they are fully awake, and Deliver (Deliver I 2.0, Mead Johnson Nutritionals, Evansville, ‘N) is often administered orally within 3 to 4 hours postoperatively. Vomiting has not been a problem.
Another problem commonly encountered is hypothermia. Intravenous fluids should be warmed before administration. We use a warm water heating pad and heat lamp during and after surgery. Ferrets are generally hardy and are good surgical candidates. Postoperative infections appear to be rare.
Even with the removal of the left adrenal gland and part of the right, most patients do not appear to require hormonal supplementation. Vital signs in these ferrets should be monitored closely in the immediate postoperative period. If recovery is prolonged or if the patient is dping poorly, a blood glucose determination should be made. If the blood glucose level is normal, corticosteroids can be administered, and blood can be saved for a resting cortisol level to ascertain the need for continued cortisone supplementation.
One problem encountered with male ferrets with adrenal disease is the presence of paraurethral cysts. Animals with this condition present with dysuria or total blockage along with other signs of adrenal disease. The cysts are thought to arise from prostatic tissue that has been stimulated by the hormones released from the adrenal gland. These cysts are present just caudal to the bladder and can usually be felt by external abdominal palpation (Figure 45-2). If the urinary tract is totally obstructed, the blockage must be relieved. This procedure can be challenging because the penis is difficult to catheterize as a result of the os penis. I have been most successful using a tomcat catheter or 3 French red rubber catheter. At the present time, the best treatment (Figure 45-2). A paraurethral cyst around the neck of the urinary bladder.) appears to be adrenalectomy. In addition, one should attempt to aspirate material from the cyst during the surgical procedure using a 22 gauge needle and a 3 mL syringe. The material in the cysts appears flocculent. Because the cysts are usually multiloculated, a few attempts should be made into different areas. Leakage from the cysts after this procedure has not been a problem. The cysts regress after the adrenal tumor has been removed. The ferret is kept on postoperative antibiotics for 14 days. In a few cases, cysts that have not regressed and that cause reobstruction need to be marsupialized. In severe cases, pre scrotal or perineal urethrostomies can be performed. In these patients, the cysts should regress after 3 to 4 weeks of antibiotic therapy.
The prognosis for patients undergoing adrenalectomy is excellent. It is the treatment of choice for this condition. In females, the swollen vulva may begin to shrink within 1 to 2 days. Hair loss takes longer to resolve. My impression is that the longer the interval between the onset of clinical signs and surgery (and usually the more extensive the alopecia), the longer the hair takes to regrow. Clinical signs return in some patients, and a second surgical procedure will be needed to remove the other gland, which has since become diseased.
As previously mentioned, insulinoma surgery is the second most common procedure performed in my practice. Signs are due to hypoglycemia and range from ferrets who begin to sleep more and seem lethargic, act nauseated, and paw at their mouths, to episodes of “vacant expressions” and staring into space, to hind limb weakness and collapse, to seizures and coma. Signs can be intermittent and can resolve quickly. Because this disease is commonly seen in ferrets over 3 years of age, early signs may be interpreted as normal aging. Diagnosis is based on the demonstration of low blood glucose concentrations and hyperinsulinemia. After a 3 hour fast, normal blood glucose should be above 80 mg/dL. Levels below 65 mg/dL suggest the diagnosis. Prolonged anorexia or starvation can produce a blood glucose level this low, but the presence of an insulinoma is much more common. Many times, the blood glucose level is below 50 mg/dL. Blood glucose levels between 65 and 80 mg/dL are suggestive of this diagnosis, and the fast should be continued for another 1 to 2 hours and an insulin level checked. An abnormally high insulin level along with low blood glucose is diagnostic.
Often, treatment begins with medical intervention. It is effective in the early course of the disease and has been used for 3 months to 2 years. Therapy is begun with prednisone at a dose of 0.5 to 2 mg/kg. The dose can be increased over time to keep clinical signs under control. Diazoxide (Proglycem), at a dose of 5 to 10 mg/kg, can be added to the prednisone regimen. It inhibits insulin release and stimulates hepatic gluconeogenesis.12 Sometimes, the dose of prednisone can be lowered when diazoxide is added. Side effects in other species include vomiting and anorexia, but these effects are rare in ferrets. The easiest form to adminis¬ter is the suspension, which is expensive.
Owners are instructed to, feed ferrets with insulinomas frequently. Owners are also instructed to avoid high sugar or carbohydrate containing supplements unless treating a hypoglycemic episode. These foods or treats can stimulate insulin secretion and can cause rebound hypoglycemia. These ferrets should be fed a high quality ferret or cat food containing an animal protein source. Brewer’s yeast should be added to the diet at a rate of one quarter teaspoon twice daily because it is a good source of chromium, which helps to stabilize blood glucose and insulin levels in humans.13 Deliver makes an excellent supplement because of its high fat content and acceptance by almost every ferret. Medical treatment is indicated in ferrets that are poor surgical ‘candidates or whose owners decline surgery for their pet. Clients should be informed that the /3 cell tumors are almost always malignant, and surgical treatment appears to slow the progression of the disease. Many ferrets become normoglycemic at least for a time after surgery. Clinical impression is that ferrets seem to do better for longer with a combination of surgical and medical treatment. In dogs with insulinomas, surgical treatment prolongs life span over medical management alone.14
Surgery is generally recommended in ferrets younger than 5 or 6 years of age. All ferrets should be carefully screened for other diseases, especially cardiac disease. As mentioned in the discussion of adrenal surgery, a cardiac ultrasound study is recommended as a presurgical screen. Often, surgery is performed concurrently for adrenal disease and insulinoma. Presurgical fasting is usually limited to 3 to 4 hours. An intravenous catheter is placed in all cases, and warmed 5% dextrose is administered during the surgical procedure.
As with adrenal surgery, a midline incision is made, and a standard exploratory operation is performed. I have seen metastasis of β cell carcinoma to the spleen and liver. In patients with splenic metastasis, splenectomy was performed. Most commonly multiple, and infrequently solitary, nodules are present. The entire pancreas should be inspected visually and also palpated. The nodules usually appear as raised areas that may be lighter in color. Sometimes, the nodules are not evident visually, yet they are firmer than the surrounding tissue, so careful digital palpation is imperative. In most cases, blunt dissection enables the surgeon to shell out the affected areas. Some minimal bleeding occurs, but it usually stops with pressure or Gelfoam application. In some cases, 5-0 or 6-0 polyglycolic acid (Dexon), polyglactin 910 (Vicryl), or polydioxanone (PDS) can be used to ligate larger vessels. If numerous nodules are present, a partial pancreatectomy can be performed. The previously listed absorbable suture can be used to ligate the pancreatic tissue in a crushing manner. If the area is small enough, one circumferential ligature can be placed around the area to be removed, or the suture can be placed in the center of the area and transfixed in both directions (Figure 45-3). In this manner, less tissue is included in each tie. Pancreatitis does not seem to be a problem after surgery.
Blood glucose should be measured postoperatively and at reasonable intervals during recovery. Some ferrets become normoglycemic 1 to 2 days postoperatively. In many cases, I observe only a slight increase in measured blood glucose, although the ferrets appear to improve clinically. Often, medical management must be continued. Some patients have Postoperative hyperglycemia, which is usually transient and reso1ves within 3 to 5 days.
The surgeon must inform clients that this is a maiignant neoplasm,15 and a cure should not be expected. One should also be sure that clients understand that medical intervention may need to be continued or resumed as the disease progresses. Even with these caveats, I believe, based on many cases, that a combination of surgical and medical management yields the best results for the longest period.
Foreign Body Surgery
Foreign body ingestion by ferrets is common, especially in young animals. The most common materials are pieces of a ferret’s plastic or rubber toys. For this reason, I recommend that owners do not provide ferrets with the soft squeaky toys commonly sold for use by ferrets or with toys made of any other material soft enough to be chewed apart. Hard nylon (Nylabone type) toys are acceptable. Other foreign bodies seen include trichobezoar, pieces of foam rubber, cork, the hard ends of shoe laces, and almost anything one could imagine.
Clinical signs include anorexia, vomiting, diarrhea, and weakness. In general, ferrets do not exhibit vomiting as often as dogs and cats, but they appear nauseated by stretching the neck and retching, salivating, and pawing at the mouth.
Diagnosis is made by history, abdominal palpation, and plain and contrast radiography. Ferrets often exhibit pain on abdominal palpation. In my experience, ferrets with trichobezoars exhibit less severe clinical signs. The most common location for foreign bodies is the small intestine, but they may also be located in the stomach or esophagus.16,17
In some cases, endoscopy may be helpful to remove esophageal and gastric’foreign bodies. I use a pediatric bronchoscope. The diameter is too large to be useful for the small intestine.
In many cases, ferrets with foreign body ingestion exhibit anorexia and are dehydrated. Therefore, adequate rehydration is important. Surgery should be considered an emergency and performed as soon as possible. A standard midline approach is used, and a complete examination of the intestinal tract is performed to check for multiple foreign bodies. In patients with esophageal or proximal duodenal obstruction, the surgeon should retropulse the material into the stomach and perform a simple gastrotomy. Gastric surgery is similar to that in the dog or cat. Closure is accomlished with a double layer simple interrupted pattern using 4-0 absorbable material.
Because of the small diameter and fragility of the intestines, gentle tissue handling is important, to mini-ize stricture of the surgical site. The incision is made jn the antimesenteric border and is closed with 4-0 or 5-0 monofilament nonreactive suture in a simple interrupted pattern. Because of its handling characterstics, I prefer 5-0 polydioxanone (PDS). If the section:of intestine appears devitalized, an intestinal resection and anastomosis should be performed. The procedure is the same as in dogs and cats. The surgical site and abdomen should be flushed completely with warmed saline solution. The omentum should be placed over the area to aid healing. Closure is routine, and as in all gastrointestinal surgery, care must be taken to avoid contamination.
Ferrets are offered water and Deliver about 12 hours postoperatively, and they are encouraged to eat solid food within 24 hours. Intravenous fluids should be continued until the patient is eating well.
- Rupprecht CE, Gilbert J, Pitts R, et al. Evaluation of an inactivated rabies virus vaccine in domestic ferrets. J Am Vet Med Assoc 1990;193:1614 1616.
- An NQ, Evans HE. Anatomy of the ferret. In: Fox JG, ed. Biology and diseases of the ferret. Philadelphia: Lea & Febiger, 1988: 100 134.
- Bernard, SL, Leathers, CW, Brobst, DF, et al. Estrogen induced bone marrow depression in ferrets. Am J Vet Res 1983;44:657.
- Meschter CL. Interstitial cell adenoma in a ferret. Lab Anim So 1989;39:353-354.
- Goad WP, Fox JG. Neoplasia in ferrets. In: Fox JG, ed. Biology and diseases of the ferret. Philadelphia: Lea & Febiger, 1988: 278 280.
- Rosenthal KL, Peterson ME, Quesenberry KE, et al. Hyperadrenocorticism associated with adrenocortical tumor or nodular hyperplasia of the adrenal gland in ferrets: 50 cases (1987-1991). J Am Vet Med Assoc 1993;203:271 275.
- Rosenthal KL, Peterson ME. Evaluation of plasma androgen and estrogen concentrations in ferrets with hyperadrenocorticism. J Am Vet Med Assoc 1996;209:1097 1102.
- Ryland LM, Bernard SL. A clinical guide to the pet ferret. Corn pend Contin Educ Pract Vet 1983;5:25 32.
- Ryland LM, Gorham JR. The ferret and its diseases. J Am Vet Med Assoc 1978;173:1154 1158.
- Rosenbaum MR, Affolter YK, Usborne AL, et al. Cutaneous epitheliotropic lymphoma in a ferret. J Am Vet Med Assoc 1996;209:1441-1444.
- Stamoulis ME, Miller MS. Cardiovascular diseases. In: Hillyer BY, Quisenberry KB, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. Philadelphia: WB Saunders, 1997:67-68.
- Feldman EC, Nelson RW. Canine and feline endocrinology and reproduction. Philadelphia: WB Saunders, 1987:259, 304-327.
- Baich JF, Balch PA. Prescription for nutritional healing. Garden Park City: Avery, NY. 1990:18 19, 211-213.
- Leifer CE, Peterson ME, Matus RE. Insulin secreting tumor: diagnosis and medical and surgical management in 55 dogs. J Am Vet Med Assoc 1986;188:60-64.
- Caplan ER, Peterson ME, Mullen HS, et a!. Surgical treatment of insulin secreting pancreatic islet cell tumors in 49 ferrets: ACVS abstract. Vet Surg l995;24:422.
- Caligiuri R, Bellah JR, Collins BR, et a!. Medical and surgical management of esophageal foreign body in a ferret. J Am Vet Med Assoc 1989;195:969-971.
- Mullen HS, Scavefli TD, Quesenberry ICE, et al. Gastrointestinal foreign body in ferrets: 25 cases (1986 1990). J Am Anim Hosp Assoc 1992;28:13-19.
Anal Sac Resection in the Ferret
James E. Creed
The ferret is a popular house pet; however, odor emitted from the anal sacs of both sexes is often objectionable. Like nearly all carnivores1 and all mustelids,2 the ferret has an anal sac on each side of the anus. The ducts open at 4 o’clock and 8 o’clock positions on the inner cutaneous zone of the anus, adjacent to the mucocutaneous junction. The sacs are interposed between the internal and external anal sphincter muscles. Material stored within the sac is secreted by a glandular complex surrounding the neck of the sac and 3 to 4 mm of the duct. This complex is evident without magnification, but a binocular loupe enhances visualization. The sebaceous gland component surrounding the distal part of the duct is covered asymmetrically by an apocrine gland component.3 Surgical removal of the anal sacs and their ducts eliminates the odor of anal sac secretions, but some odor from sebaceous and apocrine tubular glands in the perianal region typically persists.
Client request is the principal indication for performing this procedure. However, veterinarians should recommend this operation for all ferrets at 6 to 8 months of age to make them more acceptable pets. Neutering should be recommended at this age in ferrets of both sexes to reduce odor further. Neutering also prevents development of aplastic anemia in nonbreeding females, which can develop from hyperestrinism associated with prolonged estrus.3,4 The client must be made aware that anal sac resection and neutering do not eliminate all “musky” odor, because of sebaceous and apocrine glands in the ferret’s penrianal skin.
In addition to a complete physical examination, the patient’s packed cell volume of blood and total serum protein level should be determined. One study of 11 healthy male ferrets reported an average packed cell volume of 52.4% and average total serum protein of 6.0 g/dL.3 Food should be withheld for 12 hours. Anesthesia is induced with oxygen and an appropriate gaseous agent in an anesthesia chamber; it can be maintained with a mask or an endotracheal tube 2.5 mm in inner diameter (12 French outer diameter, Cole, Intermountain Veterinary Supply, N. Kansas City, MO). An alternate method is intramuscular injection of ketamine hydrochloride (26mg/kg) and acepromazine (0.2 to 0.3mg/kg).5
The ferret may be positioned for anal gland resection in dorsal or ventral recumbency. Because neutering is frequently performed and is best accomplished in dorsal recumbency, all ferrets should be positioned in this way, to provide consistent orientation of anatomic structures. The ferret is placed at the end of a table on a sandbag or similar pad to prevent loss of body heat, with its pelvic limbs pulled craniad and its tail dropped. The scrotal or ventral abdomen and perianal regions are prepared and draped for the surgical procedure. Aseptic neutering is accomplished, and then the surgical drape is shifted to expose the anal region.
A binocular loupe should be used to locate the minute opening of each anal sac duct and to aid visualization throughout the procedure. The opening of each duct and the surrounding 2 mm of skin and mucous membrane are grasped with mosquito forceps. A circumferential incision is made with a No. 15 Bard Parker scalpel blade immediately distal to the forceps tip; one must be careful not to incise too deeply. Using a gentle scraping action with the blade, skin and mucosa are reflected from the duct (Figure 45-4). The glandular complex surrounding the terminal 3 to 4mm of the duct makes dissection difficult (Figure 45-4C). This complex has a nodular surface, with skeletal muscle fibers inserting into the glandular tissue. One should not attempt to find a fascial plane at this level, and dissection should be superficial with respect to overlying tissue. Shifting the mosquito forceps to clamp them across skin, mucous membrane, and terminal duct should prevent tearing the duct as caudal traction is applied with the forceps (Figure 45-4E). Applying another forceps parallel to the first provides even more support.
A fascial plane is encountered as dissection is carried beyond the glandular complex (Figure 45-4B). The anal sac can be removed readily by reflecting sphincter muscles off the sac wall with a scraping action of the scalpel blade. Staying on the proper fascial plane not only enhances sac removal, but also minimizes hemorrhage and damage to internal and external anal sphincters. If the fascial plane is followed, little muscle will be left on the sac wall. The wall appears yellowish white; it is thin, and glandular secretions are yellow. It is easy to rupture the duct and sac, particularly if the veterinary surgeon is inexperienced. Trying to establish a fascial plane before dissecting beyond the nodular glandular complex is futile and particularly hazardous, because it is easy to cut into the duct lumen. If the duct or sac is incised, surgical extirpation can still be accomplished, but the absence of a distended sac makes the operation more tedious. Odor from an incised or ruptured sac is obnoxious, but not overwhelming.
Intraoperative hemorrhage is negligible, although sterile cotton tipped applicator sticks work well to clear oozing blood from the surgical field. Placement of sutures and administration of local or systemic antibiotics are not required.
The patient is normally discharged when recovery from anesthesia is complete. Although no serious postoperative sequelae have been observed, complications can occur. Persistent minor hemorrhage may develop postoperatively, but this ceases spontaneously. Potential complications include prolapsed rectum and fecal incontinence if trauma to the anal sphincter muscles is excessive. Staying on the proper fascial plane minimizes trauma and the possibility of these serious sequelae.
- Ewer RF. The carnivores. Ithaca, NY: Cornell University Press, 1973:95.
- Ryland LM, Gorham JR. The ferret and its diseases. J Am Vet Med Assoc 1978;173:l154.
- Creed JE, Kainer RA. Surgical extirpation and related anatomy of anal sacs of the ferret. J Am Vet Med Assoc 1981;179:575.
- Kociba GJ, Caputo CA. Aplastic anemia associated with estrus in pet ferrets. J Am Vet Med Assoc 198l;178:1293.
- Muir WW Ill, Hubbell JAB. Handbook of veterinary anesthesia. 2nd ed. Philadelphia: Mosby, 1995:368.
Soft Tissue Surgery in Reptiles
Steve J. Mehler and R. Avery Bennett
In recent years reptiles have become increasingly popular as pets. Veterinarians are called upon to perform a variety of medical and surgical procedures on these animals.1,2 The anatomy and physiology of reptiles differs from the more familiar mammalian patients and the surgeon must be familiar with these differences. Skin incisions are generally made between scales in the thin softer tissue between them. It is assumed that healing in this skin is more rapid than when an incision is made through the tough scales. A number 11 scalpel blade is particularly useful for skin incision as its fine tip allows the surgeon to incise with more precision in the zig-zag pattern required to cut between scales (Figure 45-5). In a retrospective report there was no difference in healing when the incision for celiotomy in snakes was made through the scutes (large ventral scales) on the midline compared with a lateral incision between scales.3
The incised skin of most reptiles has a tendency to invert. Because of this, an everting skin closure pattern, such as an interrupted horizontal mattress, is commonly used. Alternatively, skin staples are designed to slightly evert the skin edges when applied and serve nicely for skin closure in reptiles. Reptiles have very little subcutaneous tissue and most incisions are closed with sutures in the deep tissues and skin only. The skin of reptiles is very tough and is considered the holding layer for wound security. For example, when closing a celiotomy in an iguana, the body wall muscle is very thin and does not hold suture well. No distinct fascia is identified and the muscle does not easily separate from the skin. A two layer closure is used with a simple continuous pattern in the body wall and an everting pattern in the skin recognizing that the skin is the holding layer. Sutures are tightened to gently appose the skin edges. Sutures tightened excessively will cause necrosis of the skin within the suture and dehiscence of the incision. Reptiles do not traumatize their skin incisions or remove sutures.
The tissue reaction to 8 types of suture material and cyanoacrylate tissue adhesive placed between the skin edges in ball pythons (Python regis) has recently been reported. One centimeter skin incisions were made and a piece of suture placed in the wound. The same suture material was used to close the skin over each piece of implanted suture. One wound was left to heal by second intention and served as a control. Cyanoacrylate was not significantly different from the control wound. All suture materials induced an inflammatory response. No suture material was was absorbed at 90 days. In many instances, the inflammatory response progressed with time and in some wounds suture had been or was in the process of being extruded from the tissues. Materials that are more rapidly absorbed such as poligliecaprone 25 and chromic catgut were in the process of being absorbed at 90 days while other sutures with longer absorption times such as polydioxanone may take years to be completely absorbed. In another study polyglactin 910, poliglecaprone 25, and polyglyconate were evaluated in the skin of juvenile loggerhead sea turtles (Caretta caretta). Polyglactin 910 induced statistically significant more panniculus inflammation in juvenile loggerhead sea turtles and poliglecaprone 25 and polyglyconate caused the least cutaneous tissue reaction.5 Chromic catgut has been observed to induce granuloma formation and was not absorbed 12 weeks postoperative in some reptiles6 but appears to be less inflammatory in some chelonians.5
Given the variety of suture materials on the market that induce less inflammation and have more predictable absorption rates, the use of chromic gut is not advised. Synthetic absorbable monofilament suture materials are preferred but absorption appears to be prolonged in reptiles compared to mammals. If absorbable suture materials are placed in the skin, it should be anticipated that they will require removal.
Skin suture removal is generally not attempted for at least 4 weeks postoperative. At that time incisional healing is assessed by gently teasing the incision edges to determine wound security. Often only every other suture is removed at 4 weeks and the remaining sutures removed 2 to 3 weeks later. Ecdysis (skin shedding) is considered to speed wound healing as during this time the epidermis is metabolically active. Because of this many surgeons prefer to wait for suture removal until after the subsequent ecdysis. Environmental temperature has been shown to have an effect on wound healing in reptiles.7 The patient is maintained at the upper end of its preferred optimum temperature range during the recovery period to promote healing. Following suture removal, the skin will frequently stick to the incisional scar for several sheds (ecdysis) but this eventually resolves.
Prior to undertaking a surgical procedure in a reptile patient, the surgeon must become familiar with the unique anatomy of the particular family of reptiles to which the patient belongs.1-3 There is variation in anatomy among families of reptiles; for example, crocodilians are considered to have a 4 chambered heart while squamates (lizards and snakes) and chelonians (turtles and tortoises) have a 3 chambered heart.8-11 There is also variation within a family of reptiles. In green iguanas, the kidneys are normally located within the pelvic canal while in monitor lizards they are within the coelomic cavity.
Some features are relatively consistent across species of reptiles. In general, reptiles do not have a muscular diaphragm and, as such, have a ceolomic cavity rather than thoracic and abdominal cavities; however, crocodilians do have a relatively well developed septum between the thoracic viscera and the abdominal viscera. Reptiles do not have lymph nodes. They do not store fat in the subcutaneous tissue but have discrete fat bodies within the coelom. In some species the spleen and pancreas are intimately associated with each other forming a splenopancreas.
The urinary system of reptiles is substantially different from mammals. Reptiles have a renal portal system such that, when the portal vein is open, blood from the caudal half of the body passes through the kidney prior to reaching the systemic circulation. Urine leaves the kidneys through the ureters which empty into the cloaca, not the urinary bladder. Urine then travels from the cloaca into the bladder of those species with a urinary bladder (chelonians and some lizards) or into the colon in those species without a bladder (snakes, crocodilians, and some lizards) where water absorption and ion exchange occur.8-11 Urine does not flow through the reproductive system and the short urethra only connects the bladder to the cloaca.
The cloaca receives excretions from the ureters, colon and urinary bladder in those species with a bladder, and the reproductive system. Chelonians and crocodilians have a single copulatory organ (penis) while squamates have paired copulatory organs called hemipenes (hemipenis, singular). The copulatory organs do not contain tubular structures such as a urethra. Semen travels along a groove in the hemipenis into the cloaca of the female. The female reproductive tract is bilateral in reptiles with each oviduct having a separate opening into the cloaca.
The approach for celiotomy in reptiles varies with the family of reptile. Because reptiles lack a diaphragm celiotomy can allow access to both thoracic and abdominal viscera.
Lizards and crocodilians have a body structure more similar to mammals than chelonians and snakes. A paramedian incision is recommended in these species because of the ventral abdominal vein. This vein receives blood from the caudal abdominal wall and courses along the ventral midline 2 to 3 mm inside the body wall. It is located between the umbilical scar and the pubic bones and is suspended by a short mesovasorum. Some surgeons prefer a midline approach using meticulous dissection to avoid damaging this rather large vein.3 Making a paramedian incision 2 to 4 mm lateral to midline minimizes the risk of lacerating this vessel. It has been reported that this vein may be ligated without consequence.2,3
Closure is accomplished using a simple continuous pattern with a synthetic absorbable material on a fine, atraumatic swaged-on needle. Because the muscle of the body wall is thin and tightly adhered to the skin, care must be taken with suture placement and tension on the suture or tearing through the muscle will occur. Suturing the body wall will pull the skin edges into apposition. Skin staples or an everting pattern of a nonabsorbable material maintain skin apposition.
In laterally compressed lizards, such as chameleons, an intercostal or paracostal approach is more appropriate. The ribs in the species extend caudally close to the hip and femur so there is little space to enter through this approach alone. Exposure can be improved by combining the intercostal or paracostal approach with a ventral midline approach creating a flap on one side or the other. For closure, suture the apex of the triangle of body wall created to the cranial aspect of the ventral midline incision. Then suture the paracostal or intercostal body wall followed by closure of the ventral midline incision. The skin is closed routinely.
Snakes have organs arranged in a linear configuration. In most cases, the specific organ being approached must be identified preoperative as celiotomy will not allow access to all of the viscera. It is essential to know the location of the specific organ being approached.12
The coelomic membrane may be closed as a separate layer or incorporated in the body wall closure. The body wall is a thin pale muscle that is tightly adhered to the skin. The coelomic membrane is not attached to the skin. The muscle of the body wall is closed with a simple continous pattern using a synthetic absorbable material on a fine atraumatic needle which will also approximate the skin edges. The skin is closed with either skin staples or an everting pattern such as a horizontal mattress.
Chelonians present a unique challenge for celiotomy because of their shell. For most procedures a plastron osteotomy is required. In species with a small plastron, such as snapping turtles and sea turtles, some procedures can be accomplished through a flank incision. Some procedures, such as cystotomy, can be accomplished through this approach in other chelonians.3
The pelvic bones are avoided during plastron osteotomy to avoid injury to the appendicular skeleton. Radiographs are helpful in assessing the location and extent of the pelvic bones. In most species, osteotomy through the femoral and abdominal epidermal shields (Figure 45-6) will allow access to coelomic viscera while avoiding injury to the appendicular skeleton and heart. The osteotomy must be large enough to allow the procedure to be accomplished and located in a position to allow access to the target organ.
Plastron osteotomy is performed using a power or pneumatic bone saw, or a sterile motorized wood working tool with a fine circular saw blade. Standard bur bits are not recommended because they cut an excessively wide osteotomy which will delay bone healing. Standard surgical preparation is performed and the surface of the plastron must be completely free of keratin debris and soil. This requires a surgical scrub brush. Alcohol, ether, or acetone is used to remove grease from the surface of the plastron to allow a better bond to form between the keratin and the epoxy resin that will be used to stabilize the plastron osteotomy postoperative.
The plastron is dermal bone and efforts are made to improve the environment for bone healing. The osteotomy cut is beveled slightly and the blade should be as thin as possible so when the segment of plastron is replaced, bone-to-bone contact will be achieved (Figure 45-7). The blade is irrigated while performing the osteotomy to dissipate heat and control bone dust. It is best to make a 3 sided osteotomy in species with a hinge (e.g. box turtles). An osteotomy is made on both sides as well as the caudal margin of the proposed flap. The segment of plastron is then reflected craniad based on the intact hinge which will provide blood supply to the segment of bone. For those species without a hinge (most tortoises), the segment is cut along the cranial or caudal border and the two sides. The fourth side is partially cut with the saw and then, as the section of bone is elevated, it is cracked along the remaining border to preserve some blood supply as well as some stability.
After the bone has been osteotomized, a periosteal elevator is used to dissect the body wall off the plastron preserving the attachments of the pelvic or pectoral musculature. It may be difficult to bend the segment beyond 90 degrees and it may require an assistant to hold the segment up and out of the surgeon’s field while the procedure is performed. There are two large venous sinuses within the coelomic membrane located paramedian on each side between the midline and the bridge (junction of the plastron with the carapace). These are generally obvious during the intial approach but once manipulated, undergo vasospasm and become relatively imperceptible. Care is taken not to damage these vessels so when they dilate following closure, hemorrhage does not occur. It has been reported that they can be ligated without consequences.2,3 The incision into the coelom is made along the ventral midline. The membrane is thin and transparent in the central region where there is no muscle.
Closure is accomplished using a synthetic absorbable material in the coelomic membrane and body wall. The bone flap is replaced and secured using epoxy resin and fiberglass cloth.
Epoxy is mixed and applied 2 to 3 cm around the periphery of the plastron osteotomy and over the entire bone segment leaving a 3 to 4 mm border around the osteotomy on both sides to prevent the resin from flowing into the osteotomy which would delay healing. A sterile autoclaved piece of fiberglass cloth is placed over the plastron flap with a 2 to 3 cm border extending over the osteotomy onto the plastron. The epoxy already on the plastron is gently worked into the cloth being careful not to allow the resin to seep into the osteotomy. The epoxy is allowed to cure and a second layer is applied over the entire patch. This layer should be thin enough that the resin does not soak through the cloth and into the osteotomy. Enough layers of epoxy are applied to create a completely smooth surface with no texture from the cloth remaining. During the final curing process, a piece of plastic sheeting or wax paper is applied to the patch to prevent paper or soil from adhering to the resin. This will not stick to the epoxy and is removed the following day. Within 24 hrs the resin is completely cured and the turtle can resume normal activity, including swimming. Some surgeons prefer to apply a thin layer of antibiotic cream along the osteotomy site to prevent resin from entering and provide some antibacterial activity.
Healing of a plastron ostetomy requires 1 to 2 years.3 Patches have remained viable for over 5 years and are generally not removed. Often, the patch will fall off on its own; however if the borders become elevated from the plastron, the patch can be pried off. In young growing chelonians, the patch is cut at the growth rings after bone healing is complete to allow for shell growth. Because the epoxy is potentially carcinogenic, the cuts are best made under a hood or, at least, in a well-ventilated area using a respirator mask. Copious irrigation will help prevent aerosolization of the toxic dust.
Flank celiotomy is used in chelonians with a small plastron or in tortoises with small cystic calculi or a small intestinal foreign body.3 With the animal in dorsal recumbency, the left hindlimb is pulled caudally exposing the inguinal depression. The skin is incised in either a longitudinal or transverse manner and the muscles are bluntly separated until the coelomic membrane is identified. The membrane is grasped with tissue forceps and incised to allow access to the coelomic cavity. Through this approach the left lobe of the bladder can be accessed and with a digital exploratory, small intestinal foreign bodies can be exteriorized. This approach has not been adequate for access to the entire female reproductive tract for ovariosalpingectomy; however, focal oviductal lesions may be approached through the flank. A two or three layer closure is performed with the coelomic membrane and muscle sutured either as separate layers or together.
Surgery of the Female Reproductive Tract
Female reptiles have a bilateral reproductive tract but their reproductive physiology varies considerably. Some reptiles lay eggs (crocodilians, chelonians, and some squamates) while others deliver live babies (some lizards and some snakes). Dystocia and prevention of reproduction are the major indications for surgery of the female reproductive tract. Surgical management of dystocia is indicated when husbandry changes and medical management have failed to relieve the dystocia or if there is evidence (such as radiographic) that the eggs are unable to pass because they are too large or of an abnormal shape. Ovariosalpingectomy is performed to treat dystocia or to prevent future problems related to the reproductive tract such as yolk coelomitis, dystocia, and salpingitis.
Preovulatory egg stasis is characterized by the development of yolks on the ovary that are not subsequently released. Postovulatory stasis occurs when the eggs or feti are within the oviduct but do not pass normally. In either case, it is recommended that the ovaries as well as the oviducts be removed.3 It appears that if the oviduct is removed without removing the ovary, yolks will be released into the coelom potentially inducing yolk coelomitis. If the ovaries are removed and the oviducts left, they simply atrophy and are unlikey to cause problems in the future. Removal of one side of the reproductive tract (unilateral ovariosalpingectomy) for treatment of reproductive disease allows the patient to remain reproductively viable which may be important for herpetoculturists.
The female reproductive tract is relatively mobile within the coelom. In lizards and chelonians it is readily accessible through a standard celiotomy approach. In snakes, the tract is very long and if the entire oviduct contains eggs or feti that must be removed, it is often necessary to make several celiotomy approaches. Generally, 3-5 eggs can be manipulated out of a single salpingotomy incision.
When reproductively active, the blood vessels supplying the ovary and oviduct become engorged and hypertrophied making surgical removal more challenging. For this reason, in pet reptile species with a high incidence of dystocia, prepubertal elective ovariosalpingectomy should be considered. The procedure is much easier when the vessels, ovaries and oviducts are small and the patient is in good metabolic condition.
The oviduct wall is very thin and transparent. When there is salpingitis the wall becomes thicker but more friable making it a challenge to suture closed. Cultures and biopsies should be obtained from the oviduct for diagnostic purposes to guide the postoperative management of the case and determine the prognosis for future reproductive capability. Once the oviduct is identified, an incision is made over an egg or fetus approximately the length of the egg/fetus. If the salpingotomy incision is too small the oviduct will tear while the eggs/feti are manipulated through the incision. The first egg/fetus is generally removed without much effort. Eggs/feti that have been in place a long period of time adhere to the oviduct wall. A 20 ga catheter on a 20 cc syringe filled with saline is inserted between the egg/ fetus and oviduct wall and saline is injected to separate the wall from the egg/fetus. This will not only free the egg/fetus from its adhesions to the oviduct but also provide some lubrication. After the first egg/fetus is removed, adjacent eggs/feti are massaged toward the salpingotomy using saline injection, finger dilation, and digital manipulation to separate adhesions between the egg/ fetus and oviduct, and to extrude the egg/fetus from the salpingotomy. Once all the eggs/feti have been removed the salpingotomy is closed using a fine (6-0 to 8-0) monofilament, synthetic absorbable material on a fine atraumatic needle in a two layer inverting pattern or a simple continuous oversewn with an inverting pattern. Following a properly performed salpingotomy the prognosis for reproductive viability is good.
In cases where there is irreparable damage to the reproductive tract or where the owner desires to prevent future episodes of dystocia, ovariosalpingectomy is performed. The following discussion applies primarily to green iguanas. Other lizards and chelonians will have some variation in anatomy but the procedure is similar. In snakes with their longitudinal configuration, the ovary is cranial to the oviduct and must be approached through a separate incision or by extending the celiotomy craniad until the ovary is identified.
In iguanas, the right ovary is very close to the right external iliac vein, while the left is more loosely attached with the left adrenal gland interposed between the left external iliac vein and the ovary (Figure 45-8). When the ovary is active, as with preovulatory egg stasis, the ligament is stretched out and it is easy to apply hemostatic clips to the vessels supplying the ovary. Two clips are applied to each vessel and the vessel is transected between the clips. The process is continued until all vessels are clipped and the ovary with its multitude of yolk follicles is removed.
When the ovary is not active, removal is more challenging. Removal of the right ovary is accomplished by gently elevating the ovary, applying one or two clips between the right ovary and the right external iliac vein, then transecting the tissue distal to the clip to allow removal of the ovary. The left ovary is removed in a similar manner with the clips applied between the ovary and the left adrenal gland. The tissue distal to the clips is transected allowing removal of the ovary without damaging the adjacent adrenal gland.
Following removal of the ovaries, the oviducts are removed. Dissection is initiated at the infundibulum and continued to the cloaca. With preovulatory egg binding, the oviduct is empty and vessels are easily controlled either with hemostatic clips or bipolar cautery. One or two clips are applied to the base of each oviduct at the cloaca prior to their transection and removal.
In cases of postovulatory egg binding where the oviducts are full of eggs, the ovaries are relatively small and inactive as they have already released their yolks. The oviducts full of eggs will obscure visualization of the ovaries and are removed prior to ovariectomy. The vessels to the oviducts are generally engorged and numerous. Each vessel is identified, two hemostatic clips are applied, and the vessel is transected between them. Dissection is initiated at the ovaries and continued caudad until the oviducts can be ligated or clipped at the cloaca prior to transection. After the oviducts are removed the ovaries are visualized as described above. The ovaries are removed as described previously. Closure is routine.
Postoperative care is supportive. Most patients will have been anorectic for 2 to 4 weeks prior to surgical intervention. Fluid therapy is administered through an intravenous or intraosseous catheter. Antibiotics are indicated in the management of bacterial salpingitis. Again, the patient should be maintained at the upper end of its preferred temperature range for proper function of the immune system and the digestive system.
Castration is primarily performed in male green iguanas that have become aggressive toward their owner.3 Castration has been shown to decrease testosterone levels and sexually aggressive behaviors in other lizard species.13-15 Most commonly, orchidectomy is performed in iguanas after the aggressive behavior has developed and it may be more appropriate to perform the procedure in prepubertal iguanas before the inappropriate behaviors have developed. When performed in an aggressive animal, it appears that the aggression is not ameliorated until the following breeding season. The prognosis for attenuation of the behavior has anecdotally been reported to be around 50% following orchidectomy.3
Orchidectomy is performed through a standard celiotomy. As with the ovaries, the right testicle is more closely attached to the right external iliac vein by its short, vascular mesorchium. The right adrenal gland is located on the other side of the external iliac vein. The left testicle is more loosely attached to the left external iliac vein and the left adrenal gland is located between the left testicle and the external iliac vein (Figure 45-9). The adrenals are elongated, granular, pink glands easily distinguished from the smooth, white testicles. The testicles are covered by a capsule that can be ruptured during aggressive manipulation. Rupture of the capsule does not result in hemorrhage but the contents flow out making it difficult to continue with the dissection.
The testicles are removed in a manner similar to that described for removal of inactive ovaries. The right testicle is gently elevated and one or two hemostatic clips are applied between the testicle and the external iliac vein. The tissue distal to the clips is transected allowing removal of the testicle. The left testicle is removed following application of hemostatic clips between the left adrenal gland and the testicle. If hemorrhage from the external iliac vein occurs, one or two hemostatic clip are applied longitudinally along the damaged side of the vessel to control hemorrhage (Figure 45-10). Partial occlusion of the external iliac vein has not been associated with clinical disease; however, if over half of the diameter of the external iliac vein is attenuated, signs of vascular obstruction might be anticipated.
Urinary calculi can develop in any species of reptile that has a urinary bladder but seem to occur most frequently in desert tortoises (Gopherus agassizzii) and green iguanas. Improper nutrition and inadequate access to water or dehydration have been suggested as initiating causes.3 Clinical signs of cystic calculi include anorexia, depression, constipation from occlusion of the colon, dystocia from occlusion of the oviduct, cloacal prolapse from tenesmus, and paraparesis secondary to compressive injury to the pelvic nerves.3 A definitive diagnosis is made based on radiographs or palpation. Calcium urate calculi are radiopaque while ammonium urate calculi can be very difficult to visualize radiographically.
In chelonians, cystic calculi are palpated in the left inguinal fossa. The urinary bladder of chelonians is bilobed and the right liver lobe lays over the right lobe of the urinary bladder. Because the right portion of the bladder is compressed by the right liver lobe, most cystic calculi are present in the left lobe of the bladder. A finger is inserted into the fossa with the chelonian in a sternal recumbency. With the finger left in place, the tortoise is tipped to verticle (90 degrees) and the stone is felt hitting the finger as it falls to the dependent portion of the bladder. In lizards, cystic calculi are easily identified by abdominal palpation.
Cystotomy is performed through a standard celiotomy approach. The bladder is large and easily identified when a calculus is present. The bladder wall is very thin and transparent but becomes somewhat thicker because of the cystitis usually associated with a calculus. The bladder is isolated with moist gauze sponges or laparotomy pads prior to making the cystotomy to minimize coelomic contamination. The urine of reptiles contains mucus and urates giving it a thick, cloudy appearance which may not be easily aspirated through small suction tips. Following removal of the calculus, the bladder is irrigated to remove residual debris. Closure is accomplished using a fine (5-0 to 7-0) monofilament, absorbable material on a small, swaged-on, atraumatic needle in a simple continuous appositional pattern oversewn with an inverting pattern. Celiotomy closure is routine.
Because dehydration may cause dessication of urates within the bladder initiating calculus formation attention must be paid to maintaining adequate hydration. Antibiotics are indicated if bacterial cystitis is present. Husbandry changes (nutrition, temperature, access to water) are made where appropriate.
Reproductive Organ Prolapse
The cloaca of reptiles consists of three compartments: the coprodeum, the urodeum, and the proctodeum. Each of these compartments and their associated structures can, theoretically, prolapse; however, this has never been reported. The coprodeum is the most cranial compartment of the cloaca and is where the rectum enters.13 This compartment receives urinary and fecal wastes from the terminal colon. The urodeum is the middle section of the cloaca and is where the ureters and the reproductive systems terminate. Urinary wastes of reptiles pass into the urodeum and then into the urinary bladder (chelonians and most lizards), or into the terminal cloaca (snakes and some lizards) where water absorption occurs.13,14 The proctodeum is the caudal compartment of the cloaca and is a reservoir for fecal and urinary wastes prior to their excretion.
The anatomy and location of the male copulatory organ varies among reptile orders. Squamate reptiles (most lizards and snakes) have hemipenes (paired copulatory organs). Hemipenes in these reptiles are hollow organs that are inverted within the tail. Crocodilians, chelonians, and some lizards have a single phallus or penis. The penis of these reptiles is within the cloaca or coelomic cavity and is a solid organ. It is directed cranially within the cloaca and is everted during copulation. Neither the reptile penis or hemipenis contains a urethra. These organs are not for urination but strictly for the transport of semen.14
Paraphimosis occurs more commonly in chelonians than in squamate reptiles. Causes include excitement, stress, infection or inflammation, neurologic deficits, cloacal impaction, trauma to the exposed organ from cage mates or the enclosure substrate, forced separation during copulation, and iatrogenic trauma secondary to probing for sex determination.13,15
The prolapsed organ is often edematous from venous engorgement, lacerations from cage mates or the substrate, and may be infected, necrotic, and covered with inflammatory exudates.13,15 If the tissue is very edematous and necrotic it may be difficult to determine if the prolapsed tissue is penis/hemipenes or another structure. It is simple to ascertain the nature of the tissue in most squamates. If the base of the prolapsed tissue is coming from the caudal aspect of the vent (i.e. coming from the tail) it is most likely a hemipenis or hemipenes. In crocodilians and chelonians, with severely damaged prolapsed tissue, it may be difficult to determine the origin of the tissue without entering the coelomic cavity. The penis/hemipenis is solid and has no lumen, while prolapsed intestine is hollow and has a lumen. If the sex is unknown, the oviduct is also hollow and often contains striations, unlike the intestine.
The reptile is sedated or placed under general anesthesia and the prolapsed organ is cleaned and lubricated. If lacerations are present, attempts are made to suture them, but usually edematous tissues will not hold sutures well. The tissue is then replaced into the tail in squamate reptiles or into the cloaca in chelonians and crocodilians. Moistened cotton-tip applicators are useful in reducing the prolapsed tissue.13 If the prolapase does not reduce, application of a cold compress or hygroscopic fluids (glycerin or concentrated sugar solution) may help. In addition, stay sutures can be placed in the center of the vent (the opening of the cloaca), both proximal and distal, to help with traction. The vent can also be incised laterally on one or both sides. Once the prolapse is reduced it is kept in place with a purse string or transverse sutures in the vent. Transverse vent sutures have the benefit of allowing fecal and urinary wastes to be passed more easily than through a purse string suture. In squamate reptiles, a purse string can be placed in the vent at the base of the tail (Figure 45-11). It is best to place a stent into the vent to prevent over-tightening allowing urine and feces to pass while keeping the tissue in place. This technique allows for normal cloacal function.13 Regardless of the technique used the sutures are removed in 2 to 3 weeks.
If the tissue is necrotic or infected it should be amputated. Amputation of the penis, hemipenis or both hemipenes will not compromise urination. In snakes and lizards, amputation of one hemipenis still allows reproductive viability.13 Mattress sutures or encircling sutures are placed around the base of the prolapsed tissue, and the organ is amputated distal to the suture (Figure 45-12). The mucosa of the stump is sutured with a simple continuous pattern, and the stump is replaced into its normal anatomic location.
Prolapse of the oviducts has occurred in female reptiles.15 In some cases it is possible to reduce the prolapsed tissue; however, the viability of the tissue and assessment of damage to the suspensory ligament of the oviduct is limited.13 Amputation of the exposed tissue has been performed but celiotomy for complete assessment of the prolapsed tissue and repair or removal of the reproductive tract is recommended. If only one side of the reproductive tract is removed, the contralateral side allows for reproductive viability.
A variety of surgical procedures such as enterotomy for removal of foreign bodies may be performed in reptile patients once the surgeon is familiar with the unique anatomy of and surgical approaches used in reptile patients. Once the approach to the celomic cavity is made, most procedures are analogous to those performed in domestic animal surgery.
- Bennett RA: Reptilian surgery. Part I. Basic principles. Compendium on Continuing Education Pract Vet 1989;11:10-20.
- Bennett RA: Reptilian Surgery. Part II. Management of surgical diseases. Compendium on Continuing Education Pract Vet 1989;11:122-133.
- Mader DR, Bennett RA, Funk RS, Fitzgerald KT, et al. Surgery. In: Mader DR. Reptile Medicine and Surgery 2nd edition. Elsevier, St. Louis, Missouri; 581-630, 2006.
- McFadden MS, Bennett, RA, Kinsel MJ, Mitchell MA. Evaluation of the histologic reactions to commonly used suture materials in the skin and muscle of ball pythons (Python regis). Am J Vet Res 72 (10); 1397-1406, 2011.
- Govett PD, Harms CA, Linder KE, et al. Effects of four different suture materials on the surgical wound healing of loggerhead sea turtles,Caretta caretta. Journal of Herpetological Medicine and Surgery;14,6-10, 2004.
- Millichamp NJ, Lawrence K, Jacobson ER, et al: Egg retention in snakes. Journal of the American Veterinary Medical Association 1983;183:1213-1218.
- Smith DA, Barker IK: Preliminary observations on the effects of ambient temperature on cutaneous wound healing in snakes. Proceedings of the American Association of Zoo Veterinarians. 1983, 210-211.
- Funk RS. Snakes. In: Mader DR. Reptile Medicine and Surgery 2nd edition. Elsevier, St. Louis, Missouri; 42-58, 2006.
- Barten SL. Lizards. In: Mader DR. Reptile Medicine and Surgery 2nd edition. Elsevier, St. Louis, Missouri; 59-77, 2006.
- Boyer TH, Boyer DM. Turtles, tortoises, and terrapins. In: Mader DR. Reptile Medicine and Surgery 2nd edition. Elsevier, St. Louis, Missouri; 78-99, 2006.
- Lane T. Crocodilians. In: Mader DR. Reptile Medicine and Surgery 2nd edition. Elsevier, St. Louis, Missouri;100-117, 2006.
- McCracken HE. Organ location in snakes for diagnostic and surgical evaluation. In: Fowler ME, Miller RE, editors. Zoo and Wild Animal Medicine Current Therapy 4. WB Saunders, Philadelphia; 243-248,1999
- Moore MC: Castration affects territorial and sexual behavior of free-living male lizards, Sceloporus jarrovi. Animal Behavior 1987;35:1193-1199.
- Cooper WE, Mendonca MT, Vitt LJ: Induction of orange head coloration and activation of courtship and aggression by testosterone in the male broad-headed skink (Eumeces laticeps). Journal of Herpetology 21:96-101, 1987.
- Mason P, Adkins EK: Hormones and social behavior in the lizard, Anolis carolinesis. Hormone Behavior 1976;7:75-86.
- Lock BA. Reproductive surgery in reptiles. In: Bennett RA. Soft Tissue Surgery. Veterinary Clinics of North America: Exotic Animal Practice:3, 733-752, 2000.
- Bennett RA, Mader DR. Cloacal prolapse. In: Mader DR. Reptile Medicine and Surgery 2nd edition. Elsevier, St. Louis, Missouri; 751-755, 2006.
- Barten SL. Penile prolapse. In: Mader DR. Reptile Medicine and Surgery 2nd edition. Elsevier, St. Louis, Missouri; 862-864, 2006.
Abdominal Surgery of Pet Rabbits
Robert F. Hoyt, Jr., and Cathy A. Johnson-Delaney
Because of their relative ease of care and their docile dispositions, rabbits are becoming more and more popular as pets in today’s transient society. They are now estimated to be found in more than 1% of households in the United States, and represent some 4 million animals. As such, requests to small animal practitioners to provide rabbits with both medical and surgical care are increasing. In many areas, practitioners are responding to the increase in popularity of this animal species, with resulting supplemental income to many practices. In some cases, rabbits comprise a significant percentage of the total number of patients, and the result is that some practices are devoted exclusively to exotic animals. Unfortunately, some clinicians are failing to take advantage of emerging “pocket pet” clientele and to incorporate these patients into their practices. Practitioners’ reluctance to provide such veterinary care, especially surgery on rabbits, may, in part, be due to a lack of formal training in the species during their veterinary education and training. It may also be due to a lack of confidence based on clinical experience with the species; no mentor may have been available for guidance. Reference texts recommended for practitioners considering treating rabbits in their practices should include Ferrets, Rabbits, and Rodents Clinical Medicine and Surgery, Quesenberry K and Carpenter J editors, 3rd edition, Elsdevier, 2012, BSAVA Manual of Rabbit Surgery, Dentistry, and Imaging. Francis Harcourt-Brown and John Citty editors. 2013, the current edition of James W. Carpenter’s Exotic Animal Formulary, and the 3rd edition of Ferrets, Rabbits, and Rodents by Drs Quesenberry and Carpenter.
The intent of this chapter is to provide veterinary practitioners with basic information necessary to safely perform common abdominal surgical procedures in rabbits. Included are a general overview of anatomy, indications for each type of surgical procedure, and detailed descriptions of commonly performed procedures: gastrotomy, cystotomy, ovariohysterectomy, orchiectomy (castration), and vasectomy. Because each procedure occurs within the abdominal cavity (castration may also be performed outside the abdominal cavity), several areas are common to all and warrant discussion beforehand: a review of the unique properties of rabbit skin, preoperative considerations, guidelines for preparation of the surgical area, general surgical principles particular to the rabbit, and useful suture patterns for closure.
Anatomy of the Skin
Except for some of the heavy skinned rabbit breeds whose pelts are used in the fur trade, a rabbit’s skin is thin relative to body size.1 The full thickness of the skin, including the hypodermis and panniculus carnosus, is generally only 1.0 to 2.0 mm thick. Except for the tip of the nose and the inguinal region (in both sexes) and a small area on the scrotum in bucks, a rabbit’s skin is covered with fine textured hair, and both underfur and guard hairs are present. Rabbits generally molt their hair coats annually, with hair loss starting on the shoulders and moving caudally. Frequently, patterns and rates of hair growth and regrowth (where the hair has been clipped for a surgical procedure) do not appear uniform; this is often a concern of clients. After the rabbit’s hair has been clipped, it may not begin to grow back uniformly and may look patchy, with some areas of hair longer and appearing to grow faster than others. This unusual, seemingly abnormal skin coat can be more pronounced in young, white animals when the hair on the animal’s flank has been removed. It does, however, represent normal skin responses to variations in rabbit hair growth cycles. The raised, blotchy patches are areas of active hair growth. Beginning with the second coat of hair, waves of hair growth periodically move caudally and ventrally from the neck region. These “growth waves” occur in areas of the skin where all the hair follicles are simultaneously in an active growth cycle. Owners should be informed of this when the rabbit is discharged from the veterinary clinic.
The hair growth cycle has been divided into three main phases: anagen, catagen, and telogen. The anagen, or growing, phase, is the time when the germ cells undergo a burst of mitotic activity, leading to the formation of the sheathed hair bulb and papillary cavity and emergence from the skin surface. The catagen, or transition, phase is a brief period in which mitotic activity slows and the follicle shortens. The hair then passes into the telogen phase, which is a resting period. Changes in the vascularity and thickness of the skin are associated with these phases of the hair growth cycle. The skin thickness is approximately 1.0 mm during the telogen phase and may become 2.0 mm thick during the anagen period. As rabbits become older, the waves become less frequent and more patchy in their distribution.
Normal rabbit behavior and activity are typically sedentary, and stoic, but nervous or wary of their environment when compared with dogs and cats. As such, they may have preexisting health problems that may not manifest themselves clinically to their owners and can be easily overlooked preoperatively in today’s busy veterinary practices. This is typical of species that are prey and they will hold off showing illness. The rabbit should be examined in a quiet setting and therefore, each animal must have a complete presurgical workup including physical examination, history, and, if possible, complete blood count and urinalysis. Diet, eating habits, and volume or consistency of fecal production are important items to be addressed. Clinical or subclinical problems, such as dehydration or emerging septicemia, should be corrected before any surgical procedure, to maximize the potential for a successful outcome. Because rabbits cannot vomit, withholding of food and water before the surgical procedure is not necessary, although 2 to 3 hours of fasting will clear the oral cavity, and may decrease the ingesta within the stomach, intestines, and in particular, the cecum. Assessment of pain requires astute observation. Signs of pain include reluctance to move, sitting in a stiff, hunched posture, rapid, shallow respiration, and tensing upon palpation. An elevated rectal temperature may indicate pain, stress, inflammation or infection.
As in other species, prophylactic antibiotics may be used in rabbits undergoing surgical procedures. Because of the predominance of gram-positive bacterial flora in the rabbit gastrointestinal tract, especially the cecum, any antibiotic that affects those populations, such as oral penicillins, cephalasporins, macrolides, and tylosin, should be avoided. Antibiotics, such as trimethoprim sulfa combinations, or fluorinated quinolones such as enrofloxacin, given either individually or together, can be used effectively with minimal side effects. These drugs are generally started the day before surgery, or they are administered at induction of anesthesia and are maintained for 3 days to ensure adequate blood levels should unexpected contamination occur during the surgical procedure. Clostridial overgrowths can occur in rabbit ceca and large intestine if the diet has been high in carbohydrates and sugars when using fluoroquinolone antibiotics alone. Fluoroquinolones are not effective against clostridial infections. If clostridial populations are suspected due to diet or detection of spores on fecal gram stains, metronidazole may be added to the antibiotic regimen. In the healthy rabbit, cecoliths are ingested and a few make it through the acidic stomach to the cecum to replenish the cecal flora. During illness or antibiotic therapy, this process is interrupted. Orally administered probiotics may not survive transit through the stomach and small intestine. The colon and large intestine of the rabbit sorts materials by particle size. Particles greater than 2 mm are passed into the colon and rectum to form fecal pellets. Smaller particles are moved back to the cecum via retroperistalsis. Thus probiotics administered in small quantities rectally may actually be moved back to the cecum to enable the rabbit to reestablish normal flora. An enema of healthy rabbit cecoliths is advantageous when administered to rabbits with anorexia and diarrhea post operatively. In addition, administration of fluids and gastrointestinal motility stimulants such as metoclopromide may be indicated to enhance intestinal tract motility postoperatively. Since anesthesia and opiate analgesics may slow gastrointestinal motility, use of motility stimulants will often speed recovery of normal intestinal motility. The rabbit should be encouraged to eat soon after surgery.
Preparation of the Surgical Site
For all procedures described in this chapter, the rabbit is placed in dorsal recumbency on heated water blankets with all four limbs fully extended. The animals are preanesthetized with a combination of ketamine hydrochloride (35 mg/kg intramuscularly), xylazine (5 mg/kg intramuscularly), and glycopyrrolate (0.1 mg/kg subcutaneously); they are carefully intubated and maintained on isoflurane and oxygen during the procedure. Alternative regimens include the use of glycopyrrolate, and ketamine at 20 to 30mg/kg IM with diazepam at 1 to 3 mg/kg IM. A number of different anesthetic regimens have been published and should be considered depending on the procedure to be performed. It is important to minimize stress to the rabbit during drug administration and the anesthetic induction phase. The reader is referred to the literature for additional information on anesthetic protocols used in rabbits.2,3 Intravenous access is established with a 24 gauge catheter (Angiocath) placed in the marginal ear or cephalic vein and maintained by the slow administration of an isotonic crystalloid fluid. As the rabbit has a large and heavy abdomen in comparison with the thoracic cavity, care should be taken to slightly elevate the thorax during surgery and to not tilt the rabbit’s head and thorax downward as is sometimes performed with carnivore abdominal surgery. The reader is referred to the literature for additional information on anesthetic protocols used in rabbits.2,3
Preparing the rabbit’s skin for surgery can be a challenge to a practitioner inexperienced with the species. Removing hair at and around the incision site without damaging the skin can be frustrating. Although many clinicians use traditional clinic clippers with a No. 40 dipper clipper blade to remove hair, variable high speed clippers (e.g., Double K Industries, Inc., Model 401) specifically designed for animals with fine hair, such as rabbits and rodents are recommended. These clippers make hair removal easier and reduce the incidence of accidental cutting or burning of the skin. If possible, hair should be removed at least 5 to 10 cm in every direction from the incision site.
Rabbit skin can be sensitive to alcohol. Care should be taken to avoid excessive scrubbing when preparing the skin for surgery. After clipping the hair, the surgical site is vacuumed to remove any remaining hair and is wiped with a saline soaked gauze. The skin is then surgically prepared using alternating applications of povidone iodine soap and either alcohol or sterile saline for a total of three applications each. Each application begins at the center of the surgical site and works outward in larger and increasing sized circles. Povidone iodine solution should be sprayed over the entire surgical site and allowed to dry. Chlorhexidine surgical scrub has also been used rather than povidone iodine. The author has found that using cosmetic-grade soft cotton rather than gauze sponges causes less irritation to sensitive rabbit skin, particularly the scrotum when preparing the site for castrations. The author performs local blocks of all incisional sites using 2% lidocaine 1:10 dilution with sterile water prior to skin incisions.
General Surgical Principles
The rabbit intestine occupies most of the abdominal cavity. Handling and manipulating the intestinal tract must be minimized as the tissue is fragile. Injury to the intestine may precipitate bacterial migration which may lead to fatal peritonitis and clostridial toxicosis. Rabbits also are highly prone to form adhesions and generous lavage with warm saline should be performed prior to closure of the abdominal wall. Adhesions occur due to multiple microhemorrhages which may occur during tissue handling. Postoperative ileus or intestinal gas formation may be fatal to the rabbit because the intestinal flora is disrupted.
Taking steps to minimize inclusion of foreign materials such as gauze lint, surgical glove powder, and talc also decreases the opportunity for adhesion formation. After donning a surgical gown and gloves and before beginning the procedure, the surgeon should wash the powder and talc off surgical gloves with sterile saline soaked gauze sponges. In addition, any gauze sponges with frayed ends should be removed from the surgical tray because these sponge fragments may fall into the rabbit’s abdomen and become a nidus for adhesion formation.5
Another important practice for avoiding adhesion formation is performed during closure of the abdomen. The two cut peritoneal surfaces from the incision must be brought into apposition when closing the muscle fascia layer. This maneuver reestablishes continuity of the nonadherent peritoneal surface, which directly contacts the underlying organs. Failure to restore the peritoneal barrier often results in adhesions involving the muscle fascia layer with one or more abdominal organs, often with adverse outcomes.
Various suture materials can be used to close wounds in rabbits, including absorbable and nonabsorbable materials. Chromic or plain catgut or suture material that increases the inflammatory response should not be used in rabbits. Long-acting synthetic absorbable suture material that is broken down by enzymatic hydrolysis such as polyglactin 910 or polydioxanone suture are preferred. Nonabsorbable sutures for skin include monofilament synthetics and stainless steel. As many rabbits will remove externally placed sutures, it is recommended that skin closure be performed using a subcuticular pattern. Elizabethan collars may be necessary short-term to prevent the rabbit from opening the surgical site, however with adequate post-operative analgesia, the use of physical barriers to the incision site may be minimized. Closure of the abdomen in most rabbits is difficult in more than two layers. The muscle-fascia layer (with underlying peritoneum) is the primary strength layer of the abdomen. As such, this layer should be closed using a tapered swaged on needle and a simple interrupted suture pattern. These sutures should be placed close together. Elevating the muscle fascia layer with towel clamps placed at each end of the incision during the closure is useful and reduces suturing time. The added visualization helps to avoid accidental suturing of an underlying organs and at the same time ensures approximation of the two edges of “glistening” peritoneum with each stitch. When completed, the suture line should be evaluated for potential herniation using Brown Adson thumb forceps. The tips of the forceps are held together, and the surgeon gently probes between each suture. If the forceps can easily enter the abdomen, a potential for abdominal organ herniation exists, and additional sutures should be placed. This process is continued until sufficient sutures are placed to prevent forcep entry.
The skin may be closed using various different suture patterns and materials. I prefer to close the skin using either absorbable suture material, as mentioned earlier, on a swaged on, reverse cutting needle in a subcuticular pattern or by using surgical staples. Surgical staples are usually reserved for linear skin incisions on flat surfaces. The site and nature of the skin lesion generally dictate which method to use. Surgical glue can be used to close any gaps remaining in the skin after subcuticular closure.
Adjustable Cervical Collars (“Scratch Guards”)
Rabbits have an almost compulsive desire to keep themselves well groomed. This behavior is often exaggerated by surgery to a level which, unless they are inhibited, the animals literally lick and bite themselves down to muscle and bone, removing skin, sutures, and anything else in an attempt to eliminate the incision site pain. Post-operative pain control using a combination of opiate and an NSAID, will help to decrease self-trauma to the surgical site. Unfortunately, rabbits usually do not tolerate the traditional Elizabethan collar well postoperatively. Although this device does keep them from removing the sutures or traumatizing the incision, the animals seem frightened and frequently do not eat, drink, or move around. Often, they just stay in one location with their heads down. One soft, adjustable type of cervical collar overcomes many of the problems associated with Elizabethan collars.6 It can be easily constructed using available clinic materials and is reusable. Preparation begins with an initial circular ring made from a roll of gauze or flexible anesthetic gas tubing approximately 14 inches in circumference. Four by four gauze sponges are next wrapped around the ring to provide both padding to the animal’s neck and external diameter enlargement to the collar. The gauze is secured in place by wrapping over it with both surgical adhesive tape and Vetwrap (3M, Minneapolis, MN) applied sequentially. This not only reinforces the gauze ring, but provides for a consistent collar diameter and water resistance. The finished collar is simply placed over the animal’s head, and the slack in the ring is compressed until it’s snug. The collar can then be secured by adhesive tape so it resembles a yoke (Figure 45-13). It was coined “scratch guard” by staff members because the animals did not scratch the surgical site. When this collar is used, the affected animals seem distracted from surgical site discomfort and appear to resume normal activity and eating and drinking.
In a report of more than 1500 abdominal surgical procedures using the scratch-guard only 2.5% of animals exhibited self mutilation episodes, and those episodes occurred primarily because of the rabbit’s ability to remove the scratch guard,6 Some rabbits however will be depressed with any type of collar restraint. A belly-band wrap using Vetwrap, and taped circumferentially cranially and caudally may be needed for several days. A gauze sponge with a topical anesthetic cream or gel can be placed over the incision. A layer of gauze bandage is wrapped loosely around the rabbit’s mid-section, and finished with 2 to 4 layers of Vetwrap, then taped. The bandage needs to be loose enough to not constrict the abdomen, but layered enough to prevent the rabbit from chewing through it.
Postoperative care plays an important part in successful surgery in rabbits. Postoperative care can be broken down into two time periods: the first 24 hours after the surgical procedure and the next 13 days. After surgery, the animal is allowed to recover in a warmed, intensive care cage where the endotracheal tube is removed (on return of the animal’s gag reflex). Once the rabbit is fully conscious, a scratch guard cervical collar is fitted, and food and water are offered. Close monitoring of the animal is important during the first 24 hours postoperatively, and, therefore; the animal should remain where it can be observed frequently by staff members. During this period, the animals should have complete health checks a minimum of twice a day. This examination includes rectal temperature, pulse count, thoracic auscultation, monitoring of fluid and water intake, monitoring of urination (volume) and defecation (amount and consistency), and monitoring of the incision line and the animal’s behavior (activity and body language). In addition, the animals are continually evaluated for signs of pain, which may be obvious, such as vocalizations, to subtle, including reluctance to move, abnormal (hunched) postures, anorexia, grinding of teeth, elevated body temperature, increased respiratory rate, and unexpected aggression. If pain is present, pain relief can be provided (buprenorpbine, 0.05 mg/kg subcutaneously twice daily) as needed. NSAIDS such as meloxicam (Metacam, Boerhinger Ingleheim) at 0.2 to 0.5 mg/kg SQ or PO q 24 h or Carprofen (Rimadyl, Pfizer) at 2 mg/kg PO or SQ q 12 h or 4 mg/kg SQ q 24 h can be used with the opiate and greatly enhance the rabbit’s return to normal activity. Minimal to no fecal production suggests potential cecal stasis. A “cow patty” stool may indicate bacterial enteritis. The rabbit must resume eating and drinking as soon as possible following surgery. Rabbits normally have a small amount of ileus in the postoperative period, and the return to voluntary food and water consumption helps to prevent this occurrence from becoming deleterious. Offering the animals hay, either grass or timothy, usually stimulates reluctant animals to eat immediately. Critical Care (Oxbow Pet Products, Murdock NE) can be used to encourage eating, by assist-feeding it directly orally. Many pet rabbits will begin to eat when hand-fed and encouraged by nursing care. If the animal returns to near normal behavior, especially regarding food and water consumption, and normal urination, and defecation within 24 hours of the procedure the surgeon can send it home. To help facilitate a successful outcome, the owner should perform as many health monitoring techniques as possible, especially monitoring body temperature, incision site, food and water intake, urination and defecation (volume and consistency), movement, and overall attitude. Owners appreciate participating in the rabbit’s postoperative care and in becoming more aware of their pet’s health. A return progress visit should be scheduled for 3 to 5 days following major surgery.
Common Surgical Procedures
The common surgical procedures performed in the peritoneal cavity are discussed in this section. The techniques presented focus on procedures that I believe can be easily learned and are usually successful. No attempt is made to discuss all available surgical techniques. In addition, the description of each technique begins as if the surgeon had already opened the abdomen as discussed previously.
Rabbits are hindgut fermenters and have a simple, glandular stomach. The stomach serves as a reservoir for most of the ingested food, and it is never completely empty in a healthy animal. The stomach acids in the rabbit are among the most acidic of those of any species, with a pH of 1.2 to 1.5. This high acidity enables rabbits to use plant proteins more efficiently than most mammals and normally minimizes problems with ingested hair.
Unlike other species with incessant grooming behaviors, such as cats, rabbits physiologically cannot vomit.Consequently, ingested foreign materials, especially hair, which would normally induce a protective emetic reflex in other species, have the potential to become life threatening obstructions, unless sufficient roughage is present from the diet, and the rabbit is active, continually well-hydrated, and maintains normal gut motility. Additional predisposing factors in creating an obstructing trichobezoar may include boredom-associated over-grooming or ingestion of carpet/clothing fibers or other linear-type fabric strings, inadequate dietary roughage, anorexia because of off flavor or off odor feed, inability to smell from rhinitis, pain from sore hocks, malocclusion, lack of fresh water, or other stress factors. Once gastrointestinal motility is altered the rabbit may stop eating and drinking, and critical metabolic problems can result if this problem is not corrected.4,7-13
Common presenting complaints include anorexia, lethargy, weight loss, oligodipsia, diarrhea, or conversely, small or scant, dry feces. Other frequent clinical signs are dehydration, depression, hunched posture, tense abdomen, hypothermia, and bloating. Although definitive diagnosis of trichobezoar cannot be made without surgical exploration, a tentative diagnosis can be made based on history, clinical signs, palpation of an abdominal mass in the vicinity of the stomach, and contrast radiography, especially with fluoroscopy. Care should be taken when palpating the upper abdomen because the liver in these animals is often friable.
Rabbits with trichobezoars are frequently dehydrated and cachectic and should be treated as medical emergencies. Initial efforts should be directed at reestablishing normal homeostasis, including aggressive parenteral fluid administration before definitive therapy is pursued. Initial medical therapy involves the administration of intravenous or subcutaneous fluids, oral electrolyte fluids, and assist-feeding of a fiber-rich formula such as Oxbow’s Critical Care. Fluids such as fresh pineapple juice or crushed papain tablets in water are promoted in many publications, but other than the fluid content, and possibly the sugar content of the pineapple juice, these have not been shown to dissolve or break-up a trichobezoar. These fluids should be given in small amounts (10 to 20 mL) four to six times a day for up to 3 to 4 days. Often, this oral fluid administration both “refloats” the hair mass in the stomach and aids in quickly rehydrating the animal. Refloating allows the proteolytic enzymes and stomach acids to penetrate the trichobezoar and to begin digesting the hair. The fiber-rich roughage is necessary to encourage gut motility. A valuable tool in assessing the efficacy of medical treatment is the production of fecal pellets in increased quantities. Radiographs are useful in assessing the size of the trichobezoar and its movement. Barium as a contrast agent must be used cautiously in animals that depend on cecal digestion. If the cecum becomes coated with barium, crucial metabolism and gut flora will be altered. Because of this, the author does not utilize contrast studies in rabbits with gastrointestinal motility disorders.
Other medical treatment strategies for treating trichobezoars, formerly a strictly surgical condition, have been successful in recent years, including the use of metoclopramide.4,8-10 These newer regimens have reduced the number of animals that ultimately require surgical treatment. As a general rule, if no improvement is seen with medical therapy for trichobezoars after 3 days, these animals become surgical candidates for an exploratory gastrotomy. Animals presented for gastric foreign bodies other than trichobezoar are surgical candidates for gastrotomy (Figure 45-14). All animals having a gastrotomy are given prophylactic antibiotics, as previously mentioned, which are generally maintained for 5 to 7 days. Postoperatively, these rabbits resume food and water consumption as soon as possible. I recommend maintaining these rabbits in the hospital for several days until they return to normal eating, drinking, and defecating.
Orchiectomy (castration) is one of the most common surgical procedures performed in companion rabbits. The usual indications for removing testicles are for birth control or to modify or eliminate certain offensive behaviors intact male rabbits (bucks) often develop when they reach sexual maturity. These behaviors include: urine spraying, territory marking with both urine and feces, and aggression toward their owners or other rabbits.
Although not a panacea, castrating bucks generally makes them more docile, reduces fighting, and diminishes urine spraying. Other indications for castration are related to scrotal injury, including trauma from fighting or severe urine scalding.4
Male rabbits have two separate scrotal sacs, rather than one, as found in other placental mammals. These hairless structures lie slightly cranial to the penis. The testes are found in the abdomen at birth and descend into the scrotal sacs at approximately 3 months of age. Bucks reach sexual maturity between 4 and 5 months of age, depending on the breed of rabbit. Dwarf breeds mature more quickly than giant breeds. Castration is usually performed after the testicles descend. In addition to their peculiar scrotal anatomy, rabbits have open inguinal canals that allow the testicles to move easily between the scrotal sacs and the abdomen.8 In intact bucks, epididymal fat, which lies cranial and medial to the inguinal canal on each side is normally inhibited from entering the scrotal sac by each testicle. Because this fat lies on the abdominal side of the inguinal ring, it, in turn, inhibits intestinal herniation through the canal. Castration techniques involving incision of the scrotal sac and removal of the testicle could potentially lead initially to epididymal fat herniation and, subsequently, herniation of the intestine into the scrotum unless the inguinal canal is closed surgically following castration.
Most of the techniques described for castrating male rabbits are adaptations of techniques used in dogs and cats: scrotal approach open castration with incised tunica albuginea and preservation of the epididymal fat; scrotal approach closed castration without incising the tunica albuginea; and the prescrotal approach over the inguinal rings with inguinal ring closure.4,6,10,12 From experience, each technique is easily learned and has minimal complications. However, the scrotal perineal area can be traumatized or irritated either by the surgical procedure or by surgical preparation using any of these techniques. This trauma or irritation potentiates iatrogenic injury and the opportunity for subsequent bowel herniation or infection. Pre-surgical scrubbing should be gentle and the author prefers the use of chlorhexidine-based surgical scrubs delivered with cotton rather than gauze, and irrigated with sterile saline rather than scrubbed. The rabbit is positioned in dorsal recumbency with the thorax elevated. The incision is made on the midline just cranial to the scrotal sacs and penis. The incision should be as small as feasible (1-2 cm in length) to allow protrusion of the testicle. Ligation of the testicular vasculature is the same as in other species. For most bucks, a single suture applied through the inguinal ring and through the spermatic cord is adequate for hemostasis and closure of the inguinal ring. In this manner, the inguinal ring is closed, and no herniation can occur. The incision in the scrotum can be closed with a drop of tissue adhesive. The procedure is repeated on the opposite side. The use of manual pressure to elevate each testicle may cause bruising of the scrotal tissue, and therefore is not recommended. After both testicles have been removed, and the incisions glued, the author applies a topical anesthetic lidocaine gel to the scrotal tissue. Recently, the author has been applying Penetran ointment to the surgical area. This is an organic ammonia-based ointment that decreases pain and inflammation. It is absorbed into the skin completely so there is no residue for the rabbit to ingest.
Healx Soother (Harrison’s Pet Products, West Plam Beach, Fl). Post-operative analgesics and NSAIDS are continued for 2 to 4 days. With gentle tissue handling, the scrotum is not bruised or irritated, and with systemic pain control, most bucks seem unaware of the surgery.
An abdominal castration technique has also been described, and although it avoids the potential trauma to the scrotal or penile area, it does require post-operative use of a cervical collar and is more invasive. This method must be used in cryptorchid bucks (Figure 45-15).
When the animal is discharged from the clinic the owners must be advised that the desired effects of castration are not instantaneous. Although the animal’s testicles have been surgically removed, male hormone levels have not been eliminated. Urine spraying, territory marking, and aggression may continue for a few weeks. In addition, libido and probably viable sperm (remaining in the vas deferens) are present for a month, and thus the potential for impregnating intact females does exists during that time.
Urinary calculi (urolithiasis) are commonly encountered in clinical practice in pet rabbits, particularly in rabbits on an alfalfa-based pelleted diet. A healthy adult rabbit produces an average of 130 mL/kg of urine each day; this urine is usually turbid and varies in color from white to yellow to brown to orange to bright red.8-11,14-16 The turbidity of the urine is due primarily to mineral precipitates. Because urine is the primary route for calcium and magnesium excretion in rabbits, various crystals, including ammonium magnesium phosphate, calcium carbonate monohydrate, and anhydrous calcium carbonate precipitates, are normally found on urinalysis.8 The wide spectrum of colors and intensity is related to dietary pigments and the animal’s hydration status; higher alkalinity and dehydration are usually associated with brighter, more intense colors.8 The etiology of urolithiasis is still not clear, but several predisposing factors have been proposed, including urine stasis, genetic predisposition, dietary imbalances or diets high in calcium such as alfalfa-based diets and concurrent hypercalcemia), chronic urinary tract infections, and inadequate water intake.8,9,14 Normal urine pH in rabbits is around 8.2, but at 8.5, calcium carbonate and phosphate crystals precipitate. Urine sludge is frequently seen and may or may not exacerbate the formation of calculi. The sludge itself can be irritating to the mucosa of the bladder and urethra, and add to the discomfort of the rabbit, reluctance to urinate, and with urine retention, increase the probability of ascending infection and stone formation. The most common presenting complaint in rabbits with urolithiasis is hematuria. As mentioned previously, rabbit urine may be any of several colors, depending on urine pH and diet. Diets high in calcium, such as alfalfa, can cause the urine to become bright red orange to red. Hematuria should, therefore, be confirmed by urinalysis or urine dipstick. This condition is often diagnosed after the animal has been presented for other problems. Hematuria was reported as the chief complaint in only one of seven rabbits with urinary calculi.10 Other signs may include polyuria, perineal irritation from urine scalding, stranguria, lethargy, anorexia, hunched posture, abdominal distension, and chronic or intermittent cystitis.4,9-11 Diagnosis can be confirmed through physical examination, palpation, and radiography.
The treatment of choice for urinary calculi is cystotomy (Figure 45-16). A urine culture should be taken by cystocentesis during the surgical procedure, before entering the bladder, and submitted for culture and antibiotic sensitivity testing. Any calculi removed should also be analyzed for possible dietary adjustment as part of the postoperative treatment.
The animals should undergo diuresis for 2 to 3 days postoperatively with either intravenous or subcutaneous fluids, and appropriate antibiotic therapy should be instituted, if indicated. An opiate analgesic along with an NSAID should be used for 3 to 5 days post surgery. The opiate is usually used for 24 to 48 hours, but the NSAID may be continued for 5 to 7 days, depending on the degree of inflammation noted in the bladder wall during the surgery. Acidifying the urine as is done with carnivorous animals is not indicated. Changing the rabbit’s diet to one based on timothy or grass hays, along with increased fluid intake, and appropriate NSAIDS and antibiotics if indicated will usually prevent recurrence of calculi. However, some rabbits may have a genetic predisposition to calculi formation, and despite corrections of dietary calcium levels, correcting any hypercalcemic conditions, and adequate fluid intake and exercise, some rabbits repeatedly form calculi. Affected animals should be periodically monitored radiographically for recurrence.10
Vasectomy is generally performed on male rabbits for birth control purposes only.12 However, unlike castration, the adverse side effects of an intact buck remain, including libido, urine spraying, aggressiveness, and hormonal urge to mark territory with urine and feces. The technique involves resection of a portion of each vas deferens just cranial to the bladder after a midline laparotomy (Figure 45-17). This surgical technique is currently gaining more use in biomedical research because of interest in producing transgenic animals. Bucks that have undergone vasectomy are used to induce ovulation in embryo recipient does at the same time as the embryo donor female is mated to an intact male.17 As previously suggested in the discussion of orchiectomy, bucks that have undergone vasectomy should be separated from intact does for at least 30 days postoperatively to prevent possible pregnancy resulting from viable sperm remaining in the vas deferens.
Ovariohysterectomy (OVH) is a commonly performed procedure in small animal practice and involves the surgical removal of the ovaries, fallopian tubes and the uterus. Performing an OVH on female rabbits (does) is similar to the procedure performed on dogs and cats and only requires a knowledge of the anatomic differences of rabbits for the procedure to be adapted. One major difference is that rather than having two uterine horns, a uterine body and one cervix (uterus bicornis bicollis) as in dogs and cats, rabbits have two uteri, each opening into the vagina through a separate cervix (duplex uterus) and no uterine body. These anatomic peculiarities, at first glance, appear to complicate the traditional OVH surgery techniques taught for cats and dogs where excision of the uterus is completed at the level of the uterine body. Carefully placing a transfixion ligature just anterior to the cervix (analogous to placement in the uterine body of a dog or cat), however, enables the complete removal of the doe’s reproductive tract (Figure 45-18).
Like cats, rabbits are induced ovulators with ovulation occurring 10 to 13 hours following copulation or after orgasm induced by another doe.8,10 The gestation period is from 30 to 32 days. Female rabbits normally reach sexual maturity at 4 to 5 months, but it is best to wait until they reach at least 6 months of age before performing an OVH. Indications for performing OVH in rabbits are: (1) to prevent or treat uterine adenocarcinoma (a very common neoplasia found in 50 to 80% of does over the age of 3); (2) to correct repeated false pregnancies; (3) to prevent pregnancy; (4) to treat pyometra or uterine hyperplasia; (5) to modify aggressive behavior and biting; and (6) to decrease urine spraying.4,8,10-12 Timing of the OVH may vary as puberty and seasonality varies with breed of rabbit and whether or not it is kept indoors or outside. Owners must be aware that cessation of some of the behaviors associated with estrus will not subside instantaneously, but may take place over several weeks. Owners are advised to launder any bedding, clean the cage well before returning the rabbit to its environment as urine scents and pheromones may still be present in the environment. Environmental pheromones may trigger undesirable behaviors. As mentioned for other abdominal surgeries, adequate pain control post surgery is critical to keeping the rabbit from opening the surgical incision. I (CJD) prefer to schedule a recheck incisional appointment 7 to 10 days postoperatively to ensure the incision has healed appropriately and to remove skin sutures if used.
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- Wixson SK. Anesthesia arid analgesia. In: Manning PJ, Ringler DH, Newcomer CE, eds. Biology of the laboratory rabbit 2nd ed. San Diego: Academic Press, 1994:87 109.
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- Hoyt RF Jr, DeLeonardis J, Clements S. et al. Post operative use of adjustable cervical collars in rabbits. Contemp Top Lab Anim Sci 1994;33:822.
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- Hillyer EV. Pet rabbits. Vet Clin North Am Small Anim Pract 1994;24:25 65.
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- Garibaldi BA, Fox JG, Otto G, et al. Hematuria in rabbits. Lab Anim Sci 1987;37:769
- Kozma C, Macklin W, Cummins LM, et al Anatomy, physiology, and biochemistry of the rabbit. In: Weisbroth SH, Flatt RE, Kraus AL, eds. The biology of the laboratory rabbit. New York: Academic Press, 1974:62-63.
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