Abstract

Introduction

Laparoscopy is an integral diagnostic and surgical tool in human medicine, and its use is rapidly expanding in veterinary medicine and surgery. Laparoscopy has been successfully used by avian veterinarians for many years as a method for sex determination, diagnostic sampling, and to a lesser extent, surgical procedures.^1,6 More recently, rigid laparoscopy has gained increased popularity in reptiles and other non-domestic species.^1,2 Although rigid laparoscopy has been performed with fish, its use is still fairly uncommon, and there are few citations in the literature.^3-5 Most of the literature on laparoscopy in fish is centered on its use for the evaluation of reproductive status.^3,5 For the zoo and aquarium veterinarian, laparoscopy can provide additional diagnostic and surgical options.

Fish pose a variety of diagnostic challenges. Many of the diagnostic techniques commonly utilized in domestic animal medicine are less useful in fish. In some cases, the clinician may need to sacrifice individual fish in order to collect diagnostic material. Because laparoscopic surgery allows the clinician to directly image the various visceral organs and collect diagnostic samples, laparoscopy can improve the clinician’s diagnostic capabilities and may reduce the need to cull animals for diagnostic sampling. It has been well documented in human medicine that the use of laparoscopy versus conventional surgical techniques greatly reduces postoperative pain and allows for a much more rapid recovery.²

Materials and Methods

In order to develop rigid laparoscopic techniques in fish, it is recommended that the clinician begin with dead animals (food fish or animals prior to postmortem examination). This will allow the veterinarian to become familiar with both the instrumentation and how to adapt this equipment for use in fish. Similar to other surgical procedures in non-domestic animals, the clinician must critically evaluate the following variables prior to surgery: species-specific anatomy, available instrumentation, and procedure(s) being performed. These variables will directly influence patient positioning and the location(s) of incision site(s) for placement of the camera and instrument ports.

When evaluating coelomic organs, fish are usually placed in dorsal recumbency using a foam wedge to help hold the animal in position. Scales over the incision site are removed, and a scalpel is used to make a stab incision through the skin and underlying muscle tissue. For most species, when a general coelomic approach is being made, a paramedian incision is made just cranial to the vent. A blunt hemostat is then used to enter the coelomic cavity and to enlarge the hole for sheath placement.

Depending upon the procedure and the size of the patient, one to three incisions are made for the laparoscope and/or instrument placement. In small animals (<200 g), typically only a single incision is made and a small (14.5 Fr) laparoscopic examination sheath can then be introduced into the coelom. The insufflation hose can be directly attached to the sheath for coelomic distension and improved visualization. This sheath has two lumens: one for the laparoscope (2.7-mm, 18-cm slender telescope), and the other acts as an instrument portal (5 Fr). Instruments that have been designed for this single sheath system include biopsy forceps, grasping forceps, scissors, cautery wire and a wire retrieval basket. From a single incision, the clinician can commonly visualize and sample the following organs: liver, spleen, intestines, stomach, gonads, pericardium and the serosal surface of the swim bladder.

In larger animals, especially if extensive organ manipulation is required, other instrument ports are commonly required. In fish that are large enough to accommodate multiple instrument ports, a wide variety of surgical procedures can now be accomplished. Most trocars and associated instruments come in either 5- or 10-mm sizes. The 5-mm size will be adequate for most procedures in fish. The 10-mm size does allow slightly improved visualization and a broader range of equipment possibilities.

For procedures that involve either the swim bladder or the kidney, a lateral approach is recommended. The animal is placed in lateral recumbency, and an incision is made on the lateral body wall, in the lower two-thirds of the swim bladder. Due to the tremendous anatomic variation between species, it is recommended that a radiograph be taken prior to the surgery in order to help confirm the exact location of the swim bladder. A sterile hypodermic needle can be used to help correlate the air bladder’s radiographic location with the potential incision site. The needle is placed through the skin at the potential incision site, and its location in regard to the air bladder is confirmed on radiograph. Once the incision site is determined, a #11 scalpel blade is used to penetrate the muscular lateral body wall and enter the swim bladder. In many species, the air bladder is thick and loosely attached to the associated tissues. A sharp instrument allows easier access to the swim bladder and prevents the displacement of the air bladder, which can occur if a blunt trocar or hemostat is used. Since the air bladder is empty, there is little risk of tissue damage when it is penetrated. With the laparoscope inside the air bladder, good visualization of the entire internal lining can be accomplished. Evaluation of the swim bladder often requires very little insufflation to provide excellent visualization. Diagnostic samples can be easily retrieved for cytology, culture, or histopathology.

For renal biopsy, a 4 Fr laparoscopic scissor can be used to make a small incision through the air bladder wall just over the cranial kidney (dorso-cranial aspect of the swim bladder). Care should be taken to avoid large vessels which may travel along the dorsal midline and are adherent to the air bladder. Biopsy forceps (5 Fr) are inserted into the instrument channel and fed into the incision site over the kidney. The renal tissue may not be directly visualized but lies directly underneath the cranial aspect of the air bladder. Once the biopsy sample has been collected, a surgical gelatin sponge (Gelfoam®) can be placed at the renal biopsy site to help minimize hemorrhage.

Distension of the coelomic cavity and air bladder is facilitated by using carbon dioxide insufflation. Pressures of 4–8 mm Hg provide adequate dilation and no apparent interoperative or postoperative problems. In species that have a poorly distendable coelomic cavity, pressures in the 8–12 mm Hg range may be required. Unlike mammals, which have lungs and a diaphragm, there are no concerns in regard to pulmonary compromise at these higher pressures. High coelomic pressure may have some direct effect on cardiac function since the coelomic cavity and pericardium are in direct contact, but this is yet to be determined. In elasmobranchs, high coelomic pressure can cause gastric or colonic prolapse. When working with sharks and rays, lower pressures should be considered, and packing off the proximal esophagus and vent may be advantageous. A Veress needle is commonly used in human and domestic animal laparoscopic procedures to provide insufflation prior to trocar placement.^2,4 The Veress needle is a conventional needle covered by a spring-loaded blunt stylet.² The sharp end of the needle cuts through the abdominal wall tissue until a free space is reached, and then the blunt stylet springs over the needle to prevent organ laceration. This technique has been described in fish⁴ and works best in those animals that have a thin body wall. Due to concerns of possible iatrogenic trauma, in general the author does not use a Veress needle and instead attaches the insufflation hose directly to the port in the instrument sheath. As with a conventional laparotomy, it is recommended that as much gas as possible be removed from the coelomic space prior to closing. Buoyancy problems have not been commonly seen after laparoscopic procedures.

Isotonic, sterile saline has been used by some investigators to dilate the coelomic cavity and to provide supplemental hydration and electrolytes. In the author’s experience, when isotonic fluids are used to distend the coelomic cavity, visualization is more commonly obscured by hemorrhage or fat droplets. Fat droplets can be of particular concern when working with elasmobranchs. Elasmobranchs have a large amount of fat in their relatively large liver. These fat droplets easily become disrupted during manipulations and often obscure vision through the laparoscope.

Rigid laparoscopy can also be very useful in imaging and sampling the gills and the upper GI tract of fish.⁴ In those fish species which have gill filaments that are difficult to see (sharks, rays, eels, etc.), the laparoscope can be used for sample collection by passing the instrument through the opercular slits.⁴ Since the esophagus and stomach of most fish species are relatively short and straight, the rigid laparoscope is also useful in evaluating the oral cavity, esophagus and stomach. Saline can be used to help rinse out the stomach prior to evaluation. Biopsy forceps can be used to acquire gastric mucosa for direct examination, cultures, or histopathology.

Conclusions

Rigid laparoscopic surgery can be performed on fish, including both bony fish and elasmobranchs. It is expected that laparoscopy will become a standard technique in veterinary medicine and that it will provide the zoo and aquarium clinician with a greater variety of diagnostic and therapeutic options. Laparoscopy was found to be a very effective technique to directly visualize visceral organs and collect tissue samples. Although fish have significantly different anatomy as compared to terrestrial animals, the same laparoscopic principles can be successfully applied to this large and varied group of animals.

Acknowledgments

I would like to acknowledge Dr. Andy Stamper, Jane Davis and Jane Capobianco for their support of this work. I would also like to thank the dedicated staff at The Living Seas and the veterinarians at Disney’s Animal Programs who helped and supported me with this project.

Literature Cited

1.  Burrows, C.F., and D.J. Heard. 1999. Endoscopy in nondomestic species. In: Tams, T. (ed.). Small Animal Endoscopy. Mosby: Baltimore, MD. Pp. 433–446.

2.  Cook R.A. and D.R. Stoloff. 1999. The application of minimally invasive surgery for the diagnosis and treatment of captive wildlife. In: Fowler M.E., and R.E. Miller (eds.). Zoo and Wild Animal Medicine: Current Therapy 4. Saunders Publishing: Philadelphia, PA. Pp. 30–40.

3.  Moccia R.D., Wilkie E.J., Munkittrick K.R., and Thompson W.D. 1984. The use of fine needle fibre endoscopy in fish for in vivo examination of visceral organs, with special reference to ovarian evaluation. Aquaculture. 40: 255–259.

4.  Murray M.J. 1998. Endoscopy in fish. In: Murray M.J., Schildger B., and Taylor M. (eds.). Endoscopy in Birds, Reptiles, Amphibians and Fish. Endo-Press: Tuttlingen, Germany. Pp. 59–75.

5.  Ortenberger A.L., Jansen M.E., and Whyte S.K. 1996. Nonsurgical video laparoscopy for determination of reproductive status of the Arctic charr. Can. Vet. J. 37: 96–100.

6.  Taylor, M. 1999. Endoscopy in birds and reptiles. In: Tams, T. (ed.). Small Animal Endoscopy. Mosby: Baltimore, MD. Pp. 433–446.