Surgery in reptiles: Case studies (Proceedings)

Reptile surgery can be very complicated. Luckily in this day & age of Reptile Medicine, we are afforded several options for anesthetic as well as surgical techniques. This lecture will present a few in order to give an overview of some of the current options available to the practitioner.

Diagnosis and Treatment of Aural Abscesses in Turtles

Here we will discuss the diagnosis, etiologies, and techniques for treating aural abscesses in an Ornate Box Turtle (Terrapene ornata ornata).

Signalment

A female, approximately twelve year old, Ornate Box turtle (Terrapene ornata ornata), originally wild caught, kept in captivity for three years.

Presenting Complaint/History

This turtle presented with slowly progressive swellings on both sides of the turtle's neck, puffiness to both eyes, periodic nasal discharge from both nares, along with decreased appetite and activity.

Physical Examination

On presentation, the turtle appeared depressed, weak, and lethargic with significant bilateral aural swellings, mild bilateral blepharoedema, and mild mucopurulent nasal discharge.

Questions

a. What's your diagnosis ?

b. What husbandry conditions are likely to have induced this disorder?

c. How should it be treated ?

d. How can it be prevented ?

Differentials

Differentials for these clinical signs include the following: bilateral aural abscesses; hypovitaminosis A; upper/lower respiratory tract disease; other infectious; systemic disease; trauma; chemical or parasitic inflammation.

While the exact cause of aural abscesses cannot be definitively stated, most appear to be the result of several predisposing factors. Commonly, affected reptiles are the victims of improper husbandry including chronic suboptimal temperatures and inadequate nutrition. This then results in immunosuppression with development of secondary opportunistic infections. Therefore to arrive at a diagnosis one must begin by taking a thorough history and doing a complete physical examination. Some level of systemic evaluation of the patient is also

indicated, as ancillary diagnostic testing may provide important therapeutic and prognostic data(2,3,20) .

Treatment/Prognosis

At this time, because of the nature of the reptilian inflammatory response, the location of the inflammatory exudate, and the relative ease of surgical manipulation, a surgical approach appears to be the most appropriate method of treatment(1,11). In the case of chelonians with long standing abscesses and/or those patients with evidence of systemic disease, delaying surgical intervention and instead initiating systemic therapy of antibiotics, fluids, and other appropriately deemed supportive care for three to four days may prove beneficial towards improving the overall outcome of treatment. Delaying surgical intervention while providing needed supportive care tends to decrease the local inflammatory response and may decrease local hemorrhage intraoperatively.

Also, a calculated delay for treatment before surgery may enhance the animal's ability to tolerate the anesthesia through correction of fluid and electrolyteimbalances and may allow time to elevate the patients core body temperature to a point within the preferred optimal temperature zone (POTZ) for that species(8,20).

The author uses the physical and laboratory examination findings to determine the overall health status of the animal. From that point it is determined what degree of supportive care is indicated and when surgical intervention should be planned. Typically most presenting turtles are debilitated and respond to two to three of supportive care prior to the surgery. These turtles are often given warmed intracoelomic fluids, vitamin supplements parenterally as deemed appropriate, and preoperative antibiotics all determined on a case by case basis. They are caged in intensive care where all of their species specific physiologic needs can be met. Surgery is planned when it is determined that the animals are more alert, responsive, and hydrated, and would therefore be better able to tolerate the procedure and anesthesia.

Surgical treatment of aural abscesses should occur using appropriate anesthesia. This allows the surgeon the opportunity to perform thorough debridement of the tympanic cavity and alleviates the pain associated with the treatment of the disease process. Typically the surgical procedure does not take very long, and the author has had good success using Propofol at 10 mg/kg IV (Rapinovet, Pitman-Moore)(4,5). In the author's experience, Propofol provides a quick induction, is short acting, and produces a level plane of anesthesia with a relatively rapid rate of recovery and few side effects. Other anesthetics may also be employed.

Proper sterile surgical preparation should be employed prior to surgery. This is especially important if it is anticipated that specimens shall be collected for culture and sensitivity. Each Veterinarian may have a different surgical approach to chelonian aural abscesses. The main objective, however, is to ensure complete removal of the abscess contents from the tympanic cavity, while avoiding possible damage to the columella (stapes) during debridement, and ensuring that fluids or other materials are not aspirated by the patient during or after the procedure(9,13,20). The author's technique involves making a full thickness horizontal incision through the entire tympanum from the three o'clock position across the center of the tympanum to the nine o'clock position. A second vertical incision is then made from the 12 o'clock position down through the center of the tympanum to the six o'clock position. Together these two incisions form a cross.

The inflammatory debris may then be gently removed using small ear loops or forceps. Again, care must be taken not to damage the columella during the debridement and subsequent flushing. If desired, specimens for culture and sensitivity should be collected at this time. Ideally the author tries to remove the caseous material in one piece. After removal of all the grossly visible inflammatory debris, all aspects (especially the caudal extent) of the tympaniccavity need to be evaluated as caseous material may extend quite far, especially in more chronic cases.

Once it has been determined that all visible pieces of abscessed material have been removed, the tympanic cavity and the eustachian tube need to be gently but liberally lavaged using an appropriate antimicrobial agent. The author

prefers to use diluted chlorhexadine solution [1 part chlorhexadine : 50 parts saline] (Nolvasan, Fort Dodge, Iowa)(4,5). Diluted chlorhexadine solution has a wide spectrum of antimicrobial activity, is relatively nontoxic to tissue, and maintains a sustained residual activity. While there are reports of using dilutedpovidone-iodine (P-I) solution, it remains controversial as it has been demonstrated in in-vitro studies that concentrations in excess of 1% P-I solution

killed fibroblasts. Also, the active antimicrobial ingredient in P-I, free iodine, is inactivated in the presence of the proteins of serum – which are often present postoperatively within the tympanic cavity(20,22).

Throughout the entire procedure, the oral cavity must be examined frequently to prevent the possibility of aspiration as debris and liquids may be forced/flushed through the eustachian tube into the oropharynx. While flushing the tympanic cavity and eustachian tube, in an effort to avoid such aspiration, the author typically places the turtle in a head down position. This allows fluid and debris that enter the oropharynx to exit via gravity out of the mouth. After thorough flushing, the surgery site may be filled with an appropriate antibiotic ointment and allowed to heal by secondary intention. The author prefers to use a triple antibiotic ophthalmic ointment (TriOptic-P,Pfizer, PA). The Veterinarian or client should continue to lavage and pack the surgical site on a daily basis until healed. The decision to place the patient on systemic antibiotics is dependent upon the results of physical and laboratory evaluations. Typically if the author has started the animal on presurgical systemic antibiotics, these are continued for a prescribed course of treatment based on the initial physical examination and lab workup.

The potential for concurrent hypovitaminosis A should be addressed as well as any other husbandry issues determined to be a problem during the initial history & physical exam. For a difinitive antemortem diagnosis of Hypovitaminosis A, a vitamin A assay of the liver, or a large volume of blood is required(6,10,18) . In this case, the abscesses were the result of overall poor husbandry combined with a diet deficient in either beta carotene or preformed vitamin A retinol or retinyl ester. Additional treatment, if concurrent Vitamin A deficiency is present, would consist of parenteral injections of 1000 – 2000 IU Vitamin A/kg (Aquasol A, Armour Pharmaceutical, NY) once a week for two (mild cases) to six (severe cases) weeks(2,8,20). Higher doses of vitamin A do not speed recovery and can result in hypervitaminosis A.

Symptoms generally resolve gradually within two to six weeks depending on the severity of presenting clinical signs. The cellular debris under the eyelids can be carefully removed with a moistened cotton tip applicator and digital pressure. Ophthalmic antibiotic ointments, such as TriOptic-P, can be used at the clinician's discretion. If nasal discharge or other evidence of respiratory disease (dyspnea,rales) or systemic disease are present, then all indicated diagnostic tests (hematology, radiography, culture, etc.) should be preformed and proper treatment/medications instituted(17,21,24). Prevention involves, along with proper species specific environmental maintenance, providing growing box turtles with either a natural source of beta carotene, or supplementing their diet with an oral source of preformed vitamin A in the form of a commercial product specifically formulated for the species(2,7,20).

If the animal is anorexic, routine force feedings + fluid & electrolyte supplementation should be administered as needed to maintain a positive caloric balance. Most aural abscesses respond well to surgical treatment and heal

completely. Recurrences do occur and are often the result of either inadequate surgical debridement or failure to address the underlying, predisposing causes of the disease(20,21). Finally, at this time, it is uncertain to what degree the turtle's auditory capability is compromised by this disease process. The trauma associated with the disease itself, the surgical repair, and the fibrotic changes that most likely occur during the healing process must certainly affect the ability of the columella to transmit sound to the inner ear. However, this assumed decreased hearing capability does not appear to have any long-term effects on the reptiles commonly presented and treated for aural abscesses(2,7,20).

References

1. Bennett, R.A. 1998. Clinical, Diagnostic and Therapeutic Techniques. Proc. 5th ARAV Conference, Kansas City, MO. p. 35 - 40.

2. Boyer, T.H. 1992. Common Problems of Box Turtles (Terrapene spp.) in Captivity. Bulletin ARAV 2(1):11.

3. Boyer, T.H. Mader, D.R. 1996. Turtles, Tortoises, and Terrapins. In: Reptile Medicine and Surgery. D.R. Mader (ed), 1st edition. Sauders Publishing. Philadelphia, PA. p 332 - 336.

4. Bruederle, J.A. (1998). CVMA Exotics Formulary. Chicago, IL. p. 16 - 29.

5. Carpenter, J.W., Mashima, T.Y. and Rupiper, D.J. 1996. Exotic Animal Formulary, 1st edition.Greystone Publications. Manhatten, KS.

6. Cooper, J.E. & Sainsbury, A.W. 1995. Self-Assessment Picture Tests in Veterinary Medicine, Exotic Species. Mosby-Wolfe, Phil. PA.

7. de Vosjoli, P. and Klingenberg, R. 1995. The Box Turtle Manual. Advanced Vivarium Systems Inc., Lakeside, CA.

8. Donoghue, S., Langenberg, J. Mader, D.R. 1996. Nutrition. In: Reptile Medicine and Surgery. D.R. Mader (ed), 1st edition. Sauders Publishing. Philadelphia, PA. p 163.

9. Evans, E.E. 1963. Comparative Immunology; Antibiody response in Dipsosaurus dorsalis at different temperatures. Proc. Soc. Exp. Biol. Med.; 112: p. 531 - 533.

10. Exotic Animals, A Veterinary Handbook. 1995. Veterinary Learning Systems, Trenton, NJ.

11. Frye, F.L. and Williams, D.L. 1995. Self-Assessment Color Review of Reptiles and Amphibians. Iowa State University Press. Ames, Iowa.

12 . Frye, F.L. 1991. Biomedical and Surgical Aspects of captive Reptile Husbandry, 2nd edition, vol. 1. Krieger Publishing Co., Malabar, FL.

13. Gatten, R.E. 1989. Aspects of the Environmental Physiology of Amphibians and Reptiles. Proceedings of AAZV, Greensboro, N. Carolina.

14. Jacobsen, E.R. 1988. Exotic Animals, Contemporary Issues in Small Animal Practice; Jacobsen ER, Kollias GV (eds). Churchill Livingstone, New York, NY. Chapt. 3, p. 35 - 48.

15. Jacobson, E.R. and Kollias, G.V. 1988. Exotic Animals. Churchill Livingstone, N.York, N.Y.

16. Johnston, D.E. 1991. Exotic Animal Medicine in Practice. Vol. #2. Veterinary Learning Systems Co Inc., Trenton, NJ. USA.

17. Johnston, D.E. 1991. Exotic Animal Medicine in Practice. Vol. #1. Veterinary Learning Systems. Trenton, NJ.

18. Lawton, M.P.C. 1993. Ophthalmology of Exotic Species. In Petersen - Jones, S.M. Crispin, S.M. (eds). Manual of Small Animal Ophthalmology. BSAVA, Cheltenham.

19. Long, R.D. 1995. Self-Assessment Picture Tests in Veterinary Medicine, Small Animal Practice. Mosby-Wolfe, Phil. PA.

20. Mader, D.R. 1985. The Interrelationship Between Ambient Temperature & Reptile Health Management. Proc. Second Ann. Symp. Captive Propagation and Husbandry of Reptiles and Amphibians. Northern Calif. Herp. Society. Special Publ. No. 3:39 - 48.

21. Murray, M.J., Mader, D.R. 1996. Reptile Therapeutics. In: Reptile Medicine and Surgery. D.R. Mader (ed), 1st edition. Sauders Publishing. Philadelphia, PA. p 349 - 352.

22. Nesbitt, G.H. and Ackerman, L.J. 1991. Dermatology for the Small Animal Practitioner, Exotics, Felines & Canines. Veterinary Learning Systems Co. Inc., Trenton, NJ USA. P. 228.

23. Swaim, S.F., Lee, A.H. 1987. Topical wound medications: a review. J. Am. Vet. Med. Assoc. 190(12):1588.

24. . Vaughn, L.K., Bernheim, H.A., and Kluger, M. 1974. Fever in the Lizard Dipsosaurus dorsalis. Nature; 252: p. 473 - 474.

25. Walls, G.L. 1942. The Vertebrate Eye and Its Adaptive radiation. In Cranbrook institute of Science Bull no. 19 : 607.

Significance of and Techniques for Delivering Nutritional Support

As mentioned many reptiles presenting are malnourished either because of dietary deficiencies or improper husbandry having an adverse effect on the animals metabolism. It is generally recommended that nutritional support be provided to any patient that has acutely lost 10% of chronically 20% of its' body weight. Nutritional support can be provided by a number of means including syringe/force feeding, orogastric tube feeding, and pharyngostomy feeding tube placement. Though at times the greatest challenge is often opening the mouth - once it is open - the glottis in most reptiles, which is typically closed at rest, is located at the base of the tongue in the rostral oral cavity making it easy to avoid during orogastric intubation.

When continued nutritional support is required for reptiles with mouths that are difficult to open, placement of a pharyngostomy/esophagostomy tube is recommended. To do this, the animal is properly sedated, and a mosquito hemostat is inserted into the pharynx and pointed laterally against the wall of the esophagus. The tip of the hemostat is palpated externally, and, after carefully avoiding the vasculature in the area, a small nick incision is made over the tip. The tip of the hemostat is then gently/bluntly dissected out through the hole in the skin. Careful dissection is used for too large a hole will allow reflux of material out the stoma. A Sovereign (Sherwood Medical, St Louis, MO) red rubber feeding tube (size [Fr. MM] and length {"/cm} dependent on animal size) has been pre-measured and marked - so as to have the tip end up in the stomach. The hemostat is then used to grab the tip/gastric end of the feeding tube and the tube is pulled through the hole into the esophagus and out of the mouth. At this point the blunt/solid tip of the feeding tube is trimmed to open the end, and the tube is then redirected down the esophagus and into the stomach. The tube is then sutured in place using the "chinese finger-trap" technique. The tube is then properly secured to the animal's body. One of the most important steps in maintaining the feeding tube is to flush the tube after feedings and to place some sort of cap to close off the exposed end. In many reptile species it is amazing how well tolerated these feeding tubes may be and many times they can be left in until the animal actually begins to eat on its own with the tube still in place. When time for removal, local or systemic anesthesia may or may not be needed. Once the tube has been removed the site where the tube passed from the esophagus to the skin may or may not need to be closed. I often properly clean & disinfect the area and use topical tissue glue to close the skin only. I have not had any problems with reflux of food material etc.. and/or infection at the site, though often these animals have been on some sort of antibiotic therapy.

For herbivorous and omnivorous reptiles , Isocal and Sustacal (Mead Johnson Nutrionals, Evansville, IN), Ensure and Osmolite (Ross Labs, Columbus, OH), are appropriate liquid diets. For carnivorous reptiles Traumacal ead Johnson Nutrionals), Pulmocare (Ross Labs), and Feline and canine Clini care Liquid (Pet-AG, Elgin, IL) are appropriate liquid diets. Care must be exercised in patients suffering chronic starvation. "Refeeding syndrome", a condition referring to hypophosphatemia and hypokalemia, is seen in patients fed large amounts of calorie rich foods and it can be potentially fatal. The phosphorus and potassium move into the cells with glucose and thereby deplete circulating levels - resulting in rapid weakness and often coma - leading ultimately/often to death. It is therefore recommended that reptiles that have been chronically starved be fed 50% of their need calculated based on their real, not ideal weight. This would be continued for several days, until the patient's condition improves. The amount of calories supplemented is then increased in increments of 10 - 20 % until the recommended level of caloric supplementation for that sized animal is reached11.

For an excellent current reference on feeding carnivorous, omnivorous and herbivorous reptiles please refer to Dr. Scott Stahl's Proceedings notes and his references for the 2000 meeting of the Association of Reptilian and Amphibian Veterinarians (ARAV), Reno, NV. [pages: 177 – 182].

Case Report: Foreign Body Recovery In A Spectacled Caiman

(Caiman crocodilus crocodilus)

History/Anamnesis

On June 6th 2006, a five-year-old, male Spectacled Caiman, (Caiman crocodilus crocodilus) was referred with a three-month history of anorexia. The caiman had been seen at two other veterinary hospitals and had been treated symptomatically first with a single vitamin B-complex (Phoenix Scientific Inc., St. Joseph, MO, USA ) at 25mg/kg S.C. and enrofloxacin (Baytril® injectible, Haver/Diamond Scientific Inc., Shawnee Mission, KS, USA) at 10 mg/kg i.m. q24h for 14 days, and the second time again with enrofloxacin injections q24h and vitamin B-complex, and vitamin A & D3 (Injacom 100, Roche Inc Collegeville, PA .) injections at 200 IU/kg/wk and tube feeding. The owner observed no improvement in the caiman's condition during treatment and the client presented the animal for another opinion.

History/Husbandry

The caiman was maintained in a 567.8 Liter (150 gallon) capacity aquarium filled with 283.9 Liters (75 gal) of dechlorinated tap water, lined with 12.7 cm. of red-flint #15 pea-gravel as the substrate and biological medium. The water was cleaned via mechanical and chemical filtration utilizing a "Whisper" suspended power filter (Whisper, Tetra/Second Nature Inc., Blacksburg, VA USA) and every-other-week water change/siphoning off 1/3 the water. For biological filtration there was also an under-the-gravel filter (Lee's, San Marcos, CA USA) with a powerhead (Hagen, Ralph C. Hagen Corp., Canada) attached to the top of the siphon tube. Water temperature was maintained at 22° C (71.6° F) via a suspended aquarium water heater (Aquarius, Tetra/Second Nature, Blacksburg, VA USA). A 75 watt white light bulb was used to create a basking area within the enclosure (temperature approximately 31° C) over a 45.72 cm floating plastic platform on one side of the tank. No full-spectrum or other type of lighting was provided. The animal was offered a variety of foodstuffs, including "feeder goldfish" (Carassius auratus), adult domestic white mice (Mus musculus), and "people food" table scraps fed 2-3 times per week.

Physical Examination

A 4.25 Kg., 106.68 cm, male Spectacled Caiman presented on referral. The patient wasweak, lethargic, unable to lift its head, and the eyes were open with light responsive pupils. The patient had a respiratory rate of 10 b.pm, and a heart rate of 20 bpm. With the history of anorexia for three months, the caiman's body condition was poor with general muscle wasting apparent in the limbs and epaxial muscles. Skin/scale condition was unremarkable. Deep abdominal palpation was difficult due to the toughness of the skin, however a linear firm object, approximately 15 cm long, was palpated in the caudal coelomic cavity. Otherwise external physical examination was within normal limits.

It was recommended to the owner that a comprehensive reptile blood panel (: i.e hematology and plasma biochamistries) be performed as well as survey radiographs of the coelomic cavity to investigate the palpated firm linear object. Due to financial constraints, the client declined all blood evaluation and allowed two survey radiographs of the coelomic cavity.

Radiographic Examination

The caiman was manually restrained for radiographic evaluation. Ventro-dorsal and lateral radiograph of the abdomen revealed a curvilinear/tubular, air-filled foreign body just caudal to the diaphragm. Small mineral opacities were found adjacent to and within the hollow, tubular gastric foreign body. Based on the "protrusion" of the diaphragm produced by the tube on the lateral view, the most likely location of the foreign body is the gastric lumen. Otherwise the radiograph was unremarkable. After consulting with the owner, the diagnosis was a linear foreign body resulting from the ingestion of the missing air intake/aeration tube from the powerhead. The owner had noted the tube missing, but was unaware of its location.

Options for the case were discussed with the owner. The owner agreed that based on physical and radiographic examination, that either endoscopic retrieval or an exploratory coeliotomy was the best option to remove the tubular gastric foreign body. The animal was admitted and prepared for the procedures the next day.

Pre-, Intra-, And Post-Operative Procedures/Treatments

The caiman was hospitalized and provided a preferred optimal temperature zone of 28 - 32° C (81- 89.6° F). Fluid therapy was initiated using warmed Lactated Ringers Solution (LRS) at 15-ml/kg/day ice. Trimethoprim-sulfamethoxazole (Biocraft Laboratories, Inc., Fairlawn, NJ. USA) at 15 mg/kg p.o. q24h, vitamin B Complex at 1ml/kg q24h S.C., vitamin K (Phoenix Scientific, Inc. St. Joseph, MO, USA) at 0.25 mg/kg i.m. once, vitamin A/D3 (Injacom 100 Roche Inc.,Collegeville, PA) at 200 IU/kg/wk i.m and vitamin E/Selenium (BoSe, Schering-Plough Animal Health, Kenilworth, NJ, USA) at 0.1ml/kg i.m. once were given pre-operatively.

Pre-Operative

1. Intra-coelomic warmed Lactated Ringers Solution (LRS) 15 ml/kg/day

2. Vitamin K 0.25 mg/kg i.m.

3. Vitamin B Complex 1 ml/kg i.m.

4. Vitamin C 15 mg/kg i.m.

5. Vitamin A/D3 0.05ml/kg i.m. one time.

6. Vitamin E/Selenium 0.1ml/kg i.m. one time.

7. Sulfamethoxazole/trimethoprim (5:1) 15 mg/kg p.o q24h

8. Incubation/hospitalization at 28 - 32° C (81- 89.6° F)

Anesthetic Induction (Options):

1: Tiletamine/zolazepam (Telazol, Fort Dodge Fort Dodge, IA, USA) 1-2 mg/kg i.m.

2: Ketamine (Ketaset, Fort Dodge, Fort Dodge, IA) 5-60 mg/kg i.m.

3: Propofol (Rapinovet, Mallinckrodt Vet, Inc., Mundelein, IL USA) 10 mg/kg i.v.

4: Medetomidine (Dormitor, 1 mg/ml, Pfizer Animal Health, New York, NY, USA) 0.15 mg/kg i.m. with reversal by Atipamizole (Antisedan, 5 mg/ml, Pfizer Animal Health, NY, USA) 0.75 mg/kg i.m.

5. Induction via face mask with 5% Isoflurane (IsoFlo, Abbott Laboratories, North Chicago, IL, USA) at 2 L/min oxygen flow

Anesthetic Maintenance

Isoflurane 1-3% maintenance at 1-2 L/min oxygen flow

Anesthetic Monitoring & Support

Pulse Oximeter (Vet/Ox, 4402, Sensor Devices, Inc., Waukesha, WI.): to assess Pulse Rate and calculated % Oxygen Saturation

In-Line C0-2 Monitoring Gaymar T/Pump (Gaymar Corp. Paramus, NJ) water circulating heating pad for thermal support.

Vetronix Ventillator for respiratory Support/ventilation

Procedure(s)

Because the Spectacled Caiman was deemed to be debilitated, it was induced via face mask with 5% Isoflurane at 2 L/min oxygen flow, intubated and maintained at 2.0 % Isoflurane at 1 L/min flow with intermittent positive pressure ventilation (IPPV) at 10 breaths/min. With the anesthetized caiman in sternal recumbency, a 100 cm, 3.7mm flexible endoscope with a 2mm instrument channel (American Optical Corp., Southbridge, MA, USA) was introduced into the mouth, down the esophagus, and into the stomach. Using initially air insufflation and then sterile saline insufflation, the stomach was carefully distended and part of the 12 cm air-intake tube visualized. However because of the size, rigidity, and the fact that the stomach had begun to envelope the exposed ends of the tube forming adhesions, using either flexible grasping forceps (FB - 15/36; Olympus Optical C. Ltd., Melville, NY, USA) or flexible biopsy forceps (FB - 37/38; Olympus) the tube could not be recovered. At that point it was determined that further surgical intervention was needed. Momentarily turning off all the lights in the surgery suite, the precise location of the endoscope tip and the foreign body was determined and marked for coeliotomy. The patient was then placed in dorsal recumbency with all four limbs & tail secured to the table via straps. The surgical site was prepared using alcohol and chlorhexidine diacetate (Nolvasan, Fort Dodge, Ft. Dodge, IA, USA).

An exploratory celiotomy was performed using a paramedian incision between scales through the skin from approximately the pubic bone to the caudal extent of the sternal plate. Following the initial skin incision, the muscles of the body wall were sharply incised using Mayo scissors. The incision into the coelomic cavity was initiated caudally and extended craniad. Once the incision through the body wall was made, the ventral abdominal vein was identified coursing just inside the ventral midline. Exploration of the abdomen at this point led to the discovery of the foreign body being located in the stomach on the left cranial aspect of the abdomen. The pyloric and fundic regions of the stomach were exteriorized & isolated using 4x4 gauze sponges. A 2 cm incision was made into a relatively avascular region of the fundus. The missing air intake hose was isolated and manipulated out through the incision. The gastric mucosa was hyperemic as a result of the chronic irritation from the tube. The gastrotomy incision was closed using 5-0 Polydioxanone Suture [PDS] (PDS, Ethicon, Sommerville, NJ, USA) in a simple continuous pattern. This was followed by a second oversew using an inverting Cushing's pattern to allow for improved healing. After the completion of the second inverting layer, the surgical site and coelom were irrigated with 50 ml of warmed 0.9% sterile saline. Using 5-0 PDS, the muscles of the body wall were closed using a simple continuous pattern, followed by the subcutaneous tissues being closed via a subcuticular pattern. The skin was closed using 4-0 Green Monofilament Polyglyconate [Maxon] (Maxon, American Cyanamid Co., Wayne, NJ, USA) in an everting horizontal mattress pattern. Recovery from anesthesia was uneventful.

Post-Operative Care

The caiman was placed in post-op recovery/intensive care at 31-32° C (87.6-89.6° F) and made an unremarkable recovery. Hospitalization continued for 96 hours during which time the caiman received sulfamethoxazole/trimethoprim 15 mg/kg P.O. q24h, warmed LRS at 15 ml/kg/day i.ce., and 24 hours post-operatively the caiman was tube fed using a 50:50 mixture of a commercial tube feeding formula (Emeraid I & II, Lafeber Co., Odell, IL, USA) beginning with 5 ml/kg q8h one day after surgery, then 10 ml/kg q8h two days after surgery, then 15 ml/kg q8h on the last day of hospitalization. Forty-eight hours after first tube feeding; fecal material began to be passed along with some gravel. Approximately ninety-six hours after the surgical procedure, the caiman was discharged from the hospital continuing the sulfamethoxazole/trimethoprim 15 mg/kg P.O. q24h for 10 days.

Phone progress reports were taken 24, 48, and 72 hours after release. The activity and appetite of the caiman had returned to approximately normal levels per the owner. Subsequent phone conversations 5, 10, 15, and 20 days after release also indicated no apparent problems. A recheck at the hospital was done 35 days after release. The caiman was bright, alert, and responsive/aggressive. Appetite and activity were "back to normal" and the feces appeared normal to the owner. The weight was 6.75 kg (a gain of 2.5 kg) and the length was 107.24 cm. The surgical site was clean, dry, and intact with sufficient healing that the sutures were removed.

Almost two years have passed since the procedure, and the caiman has been seen for annual examinations. There appear to have been no follow-up complications and the skin incision has healed completely and the scar is fading.

Discussion

Anorexia is a commonly reported symptom of captive reptiles presented to veterinarians.4 Causes of anorexia in reptiles include a variety of infectious and metabolic diseases, inappropriate husbandry, and behavioral problems. Failure to stimulate the appetite of a reptile may be correlated to a clinician's inability to determine the underlying cause of disease. In this case, multiple attempts to correct anorexia were made using empirical therapeutic regimens. In some cases, financial constraints can limit a clinician's ability to pursue diagnostics; however, education of the client regarding the importance of pursuing the underlying cause of the anorexia must be made. Radiography proved diagnostic in this case.

There are several techniques described for gastric foreign body removal. Use of the endoscope has become an invaluable diagnostic tool in veterinary medicine. In the field of zoological medicine, the application of diagnostic endoscopy has shown to be of great value in a variety of species but has probably been most extensively used by avian veterinarians.2 At the present time, use of the endoscope in reptiles has not been used as extensively although there are numerous reports to indicate its use since the 1960s. The majority of previous papers describe the use of the endoscope to examine or retrieve foreign objects from the gastrointestinal tract.1 Advancements in endoscopy in reptile medicine and surgery provide clinicians the ability to perform minimally invasive techniques with shorter surgical recovery time.3 In the case of aquatic animals, the absence of an incision line reduces the likelihood of infection and the time the animal must spend out of the water. In this case, due to the size and stiffness of the foreign body combined with the significant swelling and inflammation of the stomach, endoscopic recovery was not successful. Although more invasive, the surgical approach was elected and curative.

Reports of foreign body ingestion in crocodilians are not common. However, with the voracious appetite and non-selective feeding habits of these animals, one might expect these occurrences to be more common. Increased popularity of crocodilians in zoological institutions and the pet trade are likely to increase the contact veterinarians have with these animals and the number of management related problems reported.

References

1. Ackermann J and Carpenter JW. 1995. Using endoscopy to remove a gastric foreign body in a python. Veterinary Medicine 90(8): 761-763.

2. Brearley MJ, Cooper JE, Sullivan M. 1991. Colour Atlas of Small Animal Endoscropy Endoscopy in Exotic Species. St. Louis, Mosby-Year Book, 111-122.

3. Divers S and Stahl S. 1998. An Introduction to Reptile Endoscopy. Association of Reptilian and Amphibian Veterinarians (ARAV) 1998 Annual Meeting, Practical laboratory. Kansas City, Kansas, USA.

4. Funk RS. 1996. Anorexia. In Mader D; Reptile Medicine and Surgery. WB Saunders, Philadelphia, PA., 346-348.