Surgical Management of Cloacal and Urinary Bladder Prolapse in an African Bullfrog (Pyxicephalus adspersus)
American Association of Zoo Veterinarians Conference 2001
Ryan S. De Voe, DVM; Cheryl B. Greenacre, DVM, DABVP (Avian); Branson W. Ritchie, DVM, PhD, DABVP (Avian); G. Heather Wilson, DVM
Veterinary Teaching Hospital, College of Veterinary Medicine, University of Georgia, Athens, GA, USA


Cloacal and urinary bladder prolapses are relatively common in captive anurans. Potential causes include parasitic infestations (e.g., nematodes, protozoa), ingestion of foreign bodies (i.e., substrate), neoplasia and the feeding of excessively bulky food items, especially in larger species. The anatomy of the anuran cloaca is such that cloacal prolapses can easily involve the bladder. The urinary bladder in frogs can be variable in its structure ranging from globoid to almost bilobed in some species. It arises as an outpouching off the ventral surface of the urodeum and is not directly connected to the mesonephric duct system. The urinary bladder serves as a storage reservoir for fluids and is capable of absorbing water in times of dehydration. These anatomic features allow the bladder to prolapse readily, but also allow for more options regarding therapeutic intervention.1-6

Case Study

A 1.0-kg, 11-year-old male African bullfrog (Pyxicephalus adspersus) was presented to the University of Georgia College of Veterinary Medicine Teaching Hospital for evaluation of a persistent tissue exteriorization from the cloaca. The frog was maintained alone in a 55-gallon aquarium with pea gravel substrate at room temperature. It was usually fed 1–2 adult mice or small rats per week. The frog was housed in a classroom during the school year. The exteriorized tissue from the vent had been obvious to the owner for 1 month prior to presentation. During that period the client had been treating the frog by not feeding it, because food restriction resolved a similar problems 6 months previously.

On physical examination the frog was noted to be in adequate body condition with no observable abnormalities beyond the exteriorized tissue, which was visibly diverse. It consisted of a blind-ended, light pink sack with a dense, rubbery consistency ventrally that had a fringe of hyperemic and shredded friable tissue dorsally. Gentle attempts to replace the exteriorized tissue into the cloaca were unsuccessful.

Impression slides were made of the prolapsed tissue and stained with Diff-Quick and Gram stain. Low numbers of gram-positive and gram-negative bacteria were noted. No parasitic organisms or ova were detected.

Whole-body radiographs were taken and showed the presence of multiple irregular radiodense objects (subjectively considered to be gravel) within the gastrointestinal tract. The radiodense foreign bodies were considered small enough to pass naturally.

An exploratory laparotomy was performed on the frog in order to identify the exteriorized tissue and correct the problem. Enrofloxacin (5 mg/kg) was administered intramuscularly. The frog was premedicated with an intramuscular dose of butorphanol (0.4 mg/kg), then anesthesia was induced with a tricaine methanesulfonate bath (1 g/L buffered to a pH of approximately 7 with sodium bicarbonate). Following induction, the frog was placed on clean moist towels, intubated and maintained on isoflurane with intermittent positive-pressure ventilation.

The coelom was approached via a ventral paramedian incision in order to avoid the ventral abdominal vein. The coelom was thoroughly explored. The hyperemic friable tissue was determined to be urinary bladder and was gently retracted into the coelom. The bladder was extremely traumatized and necrotic, and the decision was made to surgically remove it as close to the urodeum as possible. The necrotic tissue was removed, and the resulting defect was closed with 5-0 PDS in a simple continuous pattern. The identity or source of the other tissue protruding from the vent was not readily apparent. A colotomy helped determine that the tissue originated from the caudoventral aspect of the urodeum. The prolapsed tissue was reduced by a non-sterile assistant with the intent of removing it as close to the cloacal wall as possible via the colotomy incision. However, when reduced, the tissue mass completely occluded the mesonephric ducts, and both ducts became extremely dilated within 5–10 minutes. Centesis was used to remove the urine (0.5 ml of urine was removed from both ducts) to prevent rupture of the ducts. The tissue was then pushed back through the vent and the colon was closed with 5-0 PDS in a simple interrupted pattern. The coelom was flushed with warm saline. The coelom was closed with 4-0 PDS in a simple continuous pattern in the body wall, and everting horizontal mattress sutures in the skin. The prolapsed tissue was then approached from the vent and retracted as far as possible from the cloaca. It was then transfixed with 2-0 PDS and removed. The remaining tissue retracted into the cloaca.

Recovery from anesthesia was rapid and complete. The frog was placed in a container of aged water in an incubator maintained at 80°F. The frog listed to the right side and was unable to completely inflate its right lung for the first 2 days following surgery. Presumably the length of the frog’s trachea was overestimated and the endotracheal tube was placed into the left mainstem bronchus. The frog did recover completely prior to its discharge from the hospital.

The frog was discharged with enrofloxacin (5 mg/kg IM q 24 h for 10 days) and instructions to modify the husbandry. The gravel was removed from the bottom of the aquarium and the floor was left bare. The diet was reduced to 1–2 smaller 1/2 skinned mice once per week and was expanded to include insects and fish.

Histologically, the excised tissue was consistent with cloaca with hemorrhage and fibroplasia suggestive of a longstanding lesion. No organisms or neoplastic cells were observed.

Eight months postoperatively, the frog is reported to be apparently healthy with no recurrence of the prolapse.


In this particular case it was thought that the prolapse may have occurred secondary to chronic ingestion of the gravel substrate and overfeeding of bulky prey items. Both of these factors could presumably cause protracted straining resulting in the deformation and eventual fibrosis of the cloacal tissue. It is theorized that once the deformed fibrotic tissue reached a sufficient size it was expelled from the cloaca dragging with it the urinary bladder.

This is a case in point of the importance of determining the origin/identity of prolapsed tissue. If the exteriorized tissue had simply been reduced into the cloaca and retained with stay sutures, this frog would have most certainly ruptured both mesonephric ducts and died.

Most cases of cloacal and bladder prolapse in anurans are not as complicated or chronic as the one in this report. The vast majority are diagnosed acutely and can be effectively treated without exploratory surgery. If the bladder is intact, it can be gently reduced after cleaning and lubrication. Stay sutures can be placed across the vent or the bladder can be pexied via percutaneous placement of sutures facilitated with a lubricated cotton-tipped applicator. If the bladder is severely traumatized and disrupted it is acceptable to simply ligate the tissue and remove the necrotic portion of the urinary bladder. In captivity under controlled conditions, the importance of the urinary bladder for survival is minimal and the animals appear to thrive without it.

An effort should be made to determine the cause of the prolapse in order to administer definitive therapy and avoid recurrence. Impression smears and cytology of all prolapsed tissues should be performed as well as direct mounts plus fecal flotation if a sample is available. It may also be useful to biopsy prolapsed tissue if possible. Radiographs (±contrast) and cloacal and/or coelomic endoscopy may also be performed in pursuit of a diagnosis.

Literature Cited

1.  Carpenter, J.W., T.Y. Mashima, and D.J. Rupiper. 2001. Exotic Animal Formulary. Philadelphia, WB Saunders.

2.  Crawshaw G.J. 1998. Amphibian Emergency and Critical Care. In: Rupley, AE (ed): The Veterinary Clinics of North America, Exotic Animal Practice: Critical Care. Philadelphia; WB Saunders: 207–231.

3.  Noble, G.K. 1954. The Biology of the Amphibia. New York, NY; Dover Publications.

4.  Stetter, M.D. 2001. Fish and amphibian anesthesia. In: Heard, DJ (ed): The Veterinary Clinics of North America Exotic Animal Practice: Analgesia and Anesthesia. Philadelphia, PA; WB Saunders: 69–82.

5.  Wright, K.M. 2000. Surgery of amphibians. In: Bennett, RA (ed): The Veterinary Clinics of North America, Exotic Animal Practice: Soft-Tissue Surgery. Philadelphia, PA; WB Saunders: 753–759.

6.  Wright, K.M. 1996. Amphibian medicine and husbandry. In: Mader, DR (ed): Reptile Medicine and Surgery. Philadelphia, PA: WB Saunders: 436.


Speaker Information
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Ryan S. De Voe, DVM
Veterinary Teaching Hospital
College of Veterinary Medicine
University of Georgia
Athens, GA, USA

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