Necropsy and Biopsy Procedures
Written by Robert B. Moeller Jr., DVM
March 2000

Fish, like other vertebrates, have a complex nervous system and when handled, can experience stress. When performing most diagnostic procedures, fish should be anesthetized before handling. If a fish is to be submitted for pathology, euthanasia should be done prior to placing the animal into 10% formalin or other fixative. It would be considered inhumane to place a fish in 10% neural buffered formalin or other fixative without euthanasia.

The 1993 AVMA Report of the AVMA Panel on Euthanasia states that acceptable anesthetics to be used for the euthanasia of fish are tricane methane sulfate, benzocaine and barbiturates. A conditionally acceptable method for euthanasia is a blow to the head followed by decapitation. Other commonly used methods are carbon dioxide (four alka-selzer tablets to 500 mls of water), electrocution and hypothermia. Other anesthetic agents are available for immobilizing fish for euthanasia; an excellent review of these agents and their mechanism of action can be found in Stoskopf's book, Fish Medicine (1993).

The preferred method of euthanasia would be to anesthetize the fish to a deep plane of anesthesia (stage III or stage IV) and then sever the spinal cord just caudal to the brain. The most common and practical way to anesthetize a fish is to place the anesthetic agent in water. The fish will go through all four stages of anesthesia prior to death. The four stages of anesthesia and the clinical presentation of each are as follows:

Stage I; Induction and light sedation: The fish goes through an excitement phase with erratic swimming followed by reduced activity. The respiratory rate increases and there is a loss of some response to tactile stimulation.

STAGE II; Sedation: Fish swim slowly, have decreased gill movement (respiration), and a loss of equilibrium.

STAGE III; Anesthesia: Fish have a complete loss of equilibrium and are unable to swim. Gill movement (respiration) becomes very slow. The fish is unresponsive to external stimuli.

Stage IV; Anesthetic overdose: The fish has a total loss of gill movement and the opercules become distended. The fish goes into cardiac arrest.

Collecting Samples for Bacteria and Fungi

Sampling for bacteria and fungi should be done on fish that are brought in for examination alive or from fresh fish that had died only recently (usually less than 6 hours).

Ideally, the fish is alive when sampling cutaneous lesions. The area to be cultured should not be handled prior to culturing. Like cutaneous smears, large ulcerated areas should be avoided and small developing lesions cultured. The sterile loop or culturette should be rubbed into the lesion.

As with cutaneous lesions, gills from live fish are the best to culture. Gills are cultured by gently rubbing and rolling the sterile culturette through the gill arches. The culturette should pick up abundant mucus on the tip of the culturette. Since the gills and cutaneous lesions are exposed to the aqueous environment, a mixed bacterial culture should be expected. If a pure culture of a potential pathologic bacteria is obtained, this should be considered a possible cause of disease in this fish.

The kidney is one of the most important internal organs to culture. There are two methods of culturing the kidney. The first method is to cut the dorsal fin off, sterilize the open area with heat, cut the vertebra with a sterile scissors or scalpel and snap the fish by bringing the head and tail together. This exposes the kidney for culturing. The second method, can allow for possible contamination of internal organs. Here the fish is dipped into 70% alcohol and the abdominal cavity opened aseptically to allow all organs to be exposed. Sterilize with heat, the desired internal organs to be cultured, cut open the organ with a sterile scalpel and culture.

Another method for culturing, particularly in cases where a bacterial septicemia is suspected, is to examine heart blood. Collection of heart blood from the atrium is most productive since the atrium contains phagocytic cells that assist in clearing bacteria from the blood.

Biopsy Procedures

For biopsy specimens and bacterial cultures, a fish does not need to be killed. The fish should be anesthetized prior to performing most clinical examinations and biopsies. Biopsy procedures on fish usually are cutaneous smears, fin biopsies and gill biopsies.

Cutaneous smears are done primarily for ectoparasites. Large ulcerated lesions should be avoided; Try to find smaller developing lesions for sampling. Prior to performing cutaneous smears, bacterial cultures should be taken. The procedure for cutaneous smears involves passing several clean microscope slides over the area of interest. Only light pressure on the glass slide needs to be used to remove some epidermis and mucus. On one slide, a drop of water is placed on the smear and a cover slip is placed on the slide for examination. The other slides should be air dried or fixed in alcohol. These are stained with either new methyl blue stain or Diff-Quick stain.

A fin biopsy is accomplished by spreading out the fin and a triangular wedge shaped piece of tissue is cut between the rays of the fin. Place the fin biopsy on a slide with water and cover slip for examination.

A gill biopsy is performed by cutting a few tips of the primary lamella with the blades of the scissors. Place the lamellar tips on the slide with a drop of water, cover slip and examine. Both fin and gill biopsies should not cause undue harm to a fish.

Necropsy Procedure

Ideally, the fish should be submitted alive for the post mortem examination. This gives the pathologist a chance to observe the fish prior to euthanasia and note any important clinical signs. Unfortunately, some situations do not allow the pathologist to evaluate the fish while they are alive. Fish should be dead less than 6 hours. Fish found floating in a tank longer than 6 hours are poor candidates for necropsy due to post mortem autolysis. Dead fish should be wrapped in paper or gauze and refrigerated. Do not freeze the fish.

Prior to performing the necropsy, insure that all necropsy tools, sterile loop or culturettes, glass slides for impression smears, 70% alcohol and 10% neutral buffered formalin or Bouin's solution (I prefer Formalin) are available. A systematic approach should be used when performing the necropsy. Evaluate the external surface and note the general body condition of the fish, identify and note lesions on the skin, fins, eyes, oral cavity and anus. Take cultures of the desired lesions.

After completion of the external examination, place the fish in lateral recumbency on a disposable towel. Remove the eyes and then the operculum with the pseudobranch, place these in formalin. Remove the second and third gill arches being careful not to crush the primary lamella. Take several primary lamella from one of the gill arches and place on the glass slide for parasitic examination. Place the remaining gills into the fixation solution.

Using aseptic techniques, open the abdominal cavity by cutting through the pectoral girdle to the spine and follow the abdominal cavity to the anus, extend this cut along the ventral midline from the gills to the anus and remove the body wall. Remove the body organs (heart, liver, intestines, spleen, gonads and swim bladder) for examination. When submitting the swim bladder for histopathology, insure that the red gas forming organ is present. Sample both the anterior and posterior kidney in fish with both kidneys. In fish with fused kidneys, insure that anterior and posterior sections are submitted for examination.

Remove the brain by opening the skull just dorsal to the eyes and removing the bones over the brain. Sample all cutaneous lesions for histopathology. Insure that normal tissue from the margin of the lesion are submitted with the lesions. Cut into the skeletal muscle and look for parasitic cysts. Finally, open the stomach and intestine and examine food material.

If toxicology is desired, be sure to submit gills, kidney, liver, skeletal muscle, and fat. Toxicologic samples should be immediately frozen and stored in at -70 degrees C. Analysis of the tissue should occur as soon as possible after collection.

DISEASES OF FISH

Robert B. Moeller Jr., DVM
California Animal Health and Food Safety Laboratory System
University of California
18830 Road 112
Tulare, California 93274
559-688-7543

REFERENCES

1. Roberts R.J: Fish Pathology, Bailliere Tindall, London, Second edition, 1989.
2. Ferguson H.W.: Systemic Pathology of Fish, Iowa State Press, Ames, Iowa, 1989.
3. Anderson B.G.: Atlas of Trout Histology, Wyoming Department of Fish and Game, 1974.
4. Fox J.C.: Laboratory Animal Medicine, Academic Press, 1984.
5. Migaki G., Rebelin W.E.: The Pathology of Fishes, The University of Wisconsin Press, 1975.
6. Wolf K.: Fish Viruses and Fish Viral Diseases, Cornell University Press, London 1988.
7. Tucker C.S.: Channel Catfish Culture, Elsevier Science Publishers, Amsterdam, 1985.
8. Principal Diseases of Farm Raised Catfish, Southern Cooperative Series Bulletin No 225, 1985.
9. Wales J.H.: Microscopic Anatomy of Salmonids. An Atlas, United States Department of the Interior, Resource Publication 150, 1983.
10. Grizzle J.M.: Anatomy and Histology of the Channel Catfish, Auburn Printing Co, 1976.
11. Reichenbach-Klinke H. H.: Fish Pathology, T.F.H. Publications, Inc. Neptune City, NJ. 1973.
12. Stoskopf, M.K.: Fish Medicine, W.B. Saunders Co. 1993.
13. DeTolla, L.J., Srinivas, S.: Guidelines for the Care and Use of Fish in Research. Institute of Laboratory Animal Resources Journal. Vol 37:4(1995), pp159-173.
14. Kane, A.J., Gonzalez, J. F., Reimschuessel, R: Fish and Amphibian Models Used in Laboratory Research. Laboratory Animal. Vol 25:6(1996), pp 33-38.
15. Lewbart G.A. Self-Assessment Color Review of Ornamental Fish, Iowa State Press,1998.
16. Bruno D.W., Poppe T.T., A color atlas of Salmonid Diseases. Academic press, 1996.