Diagnostic Techniques for Fish
Greg Lewbart M.S., V.M.D., Dipl. ACZM
The purpose of this article is to familiarize the veterinarian with some of the basic and easy to perform diagnostic procedures used for tropical fish. All of the following techniques can be performed quickly in the clinic or in the field. Most require little more than a simple water test kit, dissecting kit and compound microscope. Noga has published a comprehensive article on the subject of biopsy and necropsy procedures.1 Some fundamental hematology, microbiology and histopathology procedures will also be reviewed.
The period of morbidity among tropical fishes is commonly very short (i.e. fine one moment, dead the next). It is important to quickly arrive at an accurate diagnosis in order to institute the proper treatment and/or husbandry management correction. This proceedings article has been written with these considerations in mind.
As with any sick animal a good history can be the clinicians best friend. Be sure and obtain a complete one. There are several good references on what questions to ask a client with sick tropical fishes.2,3,4 The clinician will require such important information as: How long has the client owned the fish? How experienced is the client? What and how often are the fish fed? Have any new fish been introduced into the aquarium or pond recently and if so were they quarantined? Have the fish been treated with any medications? Armed with answers to these and other background questions the veterinarian or technician is ready to test the water.
There are numerous good references on the subject of water quality parameters and water testing. While comprehensive understanding of water chemistry is an asset for the clinician, it is not necessary for proper and accurate water evaluation. The ability to measure and understand the relation between about a half dozen parameters is adequate. If at all possible, water quality should be evaluated before any diagnostic procedures are performed on the fish patient.
I recommend obtaining a good and durable test kit which can be purchased for under $200.00 (manufacturers and distributors are listed at the end of the article). The kit which we commonly use performs about ten different tests and contains an easy to follow instruction booklet. There are also no probes or electrodes to maintain as all tests are colorometric comparisons or simple titrations.
Regardless of what test kit you have in your clinic you will want to be able to evaluate water temperature, ammonia nitrite, pH and total alkalinity. Measuring dissolved oxygen is important but in most cases the history or tank examination will tell you if there is an aeration problem. Most good water test kits will also include a dissolved oxygen test.
Biopsy and Other Sample Techniques
In many cases, and especially those dealing with a client's pet fish, the clinician may want to take some tissue samples for examination without killing the fish. Many procedures can be rapidly performed with little risk to the piscine patient. Naturally, larger fish fare better than smaller fish and the overall condition of the animal is also a factor in how it will respond to biopsy techniques.
When I have ruled out a water quality problem, and feel there may be a parasitic or bacterial problem, I begin with the following simple procedures: The skin scraping, fin clip and in some cases the gill snip. An anesthetic agent such as tricaine methanesulfonate (FinquelR, MS-222 ) can be used to restrain the fish and make these procedures easier and safer. This particular agent is purchased in a crystalline form and can be used to produce a working stock solution of 10 grams per liter of clean, dechlorinated water. By making dilutions from this stock solution the clinician can accurately formulate safe and effective anesthetic solutions. Each liter of stock solution should be buffered with about 5 grams of baking soda.5 The stock solution should be stored in a glass bottle away from light at room temperature. When prepared and stored in this manner, the stock solution is good for about 30 days. Final concentrations of between 100 and 150 mg/liter will usually anesthetize most fish within a matter of three to five minutes. Several studies have investigated clove oil as a fish anesthetic.6,7 Clove oil is available on an over-the-counter basis at most pharmacies. Eugenol is not completely soluble in water and should be diluted with ethanol at a ratio of 1:10 (clove oil:ethanol) to yield a working stock solution of 100 mg/ml since each ml of clove oil contains approximately 1 gram of drug. Concentrations of between 40 and 120 mg/liter are effective in freshwater and marine species and results are comparable to MS-222, except that recovery may be prolonged.6
After the fish loses its ability to maintain equilibrium, it is removed from the water, the procedures are performed and the fish is placed in a "recovery" vessel containing clean water and aeration. A coverslip can be used to obtain the skin scraping sample. The coverslip should be firmly drawn across the area to be sampled, making sure that some mucus and epidermal tissue remains on the coverslip. An alternative method is to use a sterile scalpel blade for this procedure. The tissue sample which is now on the tip of the scalpel is then placed on a slide which contains several drops of clean water. After the coverslip is applied, the specimen is ready for microscopic examination. A fin biopsy can be easily obtained using a pair of fine scissors. Slide prepared with drops of water should be close by before samples are taken. Several small pieces of gill tissue can be safely cut away using small suture removing scissors after deflecting the operculum. Care must be taken to only remove a couple millimeters of primary gill lamellae. There is usually some bleeding following this procedure but it should subside quickly.
One concept to keep in mind is that many ectoparasites that affect gill tissue are usually also found on the skin and fins. Treating the parasites on the skin may also take care of the parasites on the gills. Gill damage due to environmental problems (high ammonia, bacterial gill hyperplasia) can only be evaluated after a gill biopsy has been performed.
Obtaining blood samples from tropical fish is challenging but not impossible. It is very difficult to do in fish less than three inches long (if you want the fish to survive). A sterile blood sample is a useful way to culture for a suspected bacteremia or septicemia. Fresh whole mount blood smears can be valuable in diagnosing protozoal blood parasites (e.g. trypanosomes) and in an overwhelming septicemia motile gram negative rods can frequently be observed darting across the microscopic field of view. Stained blood smears will reveal numerous nucleated erythrocytes, leukocytes and thrombocytes. Campbell has published an excellent review of piscine hematology.6 Some people insert the needle along the lateral line near the tail of the fish and others take a mid-ventral approach, entering the vein from its ventral aspect. Once the needle touches the vertebral spinal body, the needle can be gently "walked" ventrally until it drops into the caudal venous sinus. Slight constant negative pressure on the plunger of the syringe will facilitate sample collection. If a blood sample is necessary from a very small fish and survival is not important in diagnosis (many other fish may be at risk and a necropsy is the best option) the tail can be removed at its base (caudal peduncle) and the small drop of blood which appears can be collected with a clean capillary tube and then placed on a slide.
Performing a fecal examination on a fish is not usually at the top of a list of diagnostic procedures even though it is a relatively easy and valuable test to perform. If the owner cannot obtain a fecal sample from the bottom of the aquarium, the fish can be placed in water containing tricaine methanesulfonate. Many fish will defecate as they relax in the anesthetic solution. If time is not a factor, the fish can be placed in a plastic bag, clean jar, or small aquarium, and a fecal sample can be collected within a matter of hours in most cases.
One final and very important area of diagnostics involves the field of microbiology. Cultures of skin and gill tissues are not especially helpful due to the ubiquitous nature of aquatic bacterial pathogens. Cultures of clinically healthy fish and clean water will commonly reveal the presence of gram negative bacteria. Clean blood samples are valuable in detecting and identifying septicemia. As with terrestrial animal medicine, every attempt must be made to procure sterile samples for microbiology. de Guzman and Shotts have published a nice review of the subject.9 A popular culture site in fish is the kidney. The kidneys of fish run just ventral to the spinal column and are generally found the length of the body cavity. Culturing kidney tissue is a postmortem technique which is relatively easy to do. Culture samples may be processed in the clinic and antibiotic sensitivities run. Many small animal clinics are not equipped to perform microbiology testing. Fish cultures can be sent out of house since many veterinary schools and state agricultural diagnostic laboratories are familiar with fish bacteriology.10
Radiograph a fish? Why not? Here at NC State it is part of many a pet fish diagnostic evaluation. Radiographic findings have been instrumental in diagnosing and treating a number of fish cases.11,12,13 Fish are relatively easy to radiograph. Most can be handled without anesthesia by simply removing the fish from a tank or bucket, placing it on a radiographic plate (protected by a plastic bag), and making the exposure. Fish will even tolerate contrast studies, CT scans, and bone scans.14
On some occasions, a full necropsy is the only way to arrive at an accurate diagnosis of a disease problem. Necropsies have several obvious advantages over biopsy procedures. Tissues can be looked at thoroughly and completely. Any and all tissues and organs are available for both gross and histopathological inspection. Dead fish autolyze rapidly and as such are usually not good necropsy specimens. If at all possible, the clinician should obtain a moribund fish and quickly kill it for examination. An overdose of tricaine methanesulfonate (over 400 mg/l for 15 minutes) works well. A rapid cut with a scalpel at the base of the cervical spine will quickly dispatch a fish. Once the fish is dead, the operculum can be removed to expose the gills. Samples can be taken for immediate inspection under the microscope and preserved in 10% neutrally buffered formalin for histopathology. Fish tissues can be preserved and handled like any mammalian biopsy sample. Once the abdominal organs are visible, they may be examined and removed. Small pieces of tissue may be excised, squashed on a slide under a coverslip, and examined under the microscope. This quick and easy procedure may be helpful in diagnosing a parasite problem or a condition such as fatty liver disease. Any suspect tissues can be preserved in formalin for histopathology. One nice aspect of tropical fish anatomy is that most of these animals are relatively small and entire organs or organ systems can be fixed in formalin.
These notes was intended to introduce the veterinarian to some of the basic diagnostic techniques which are commonly utilized in the field of tropical fish medicine. The reader will realize that most of the included procedures are easy to perform and relatively inexpensive As the field of ornamental fish medicine continues to grow and more veterinarians become involved, we will see the appearance of more specialized and informative procedures. Regardless of how sophisticated the diagnostics and modern technology, we can never get away from the reality of good water quality and sound husbandry principles when dealing with the many species of freshwater and marine tropical fishes.
1. Noga, E.J. 1988. Biopsy and Rapid Postmortem Techniques for Diagnosing Diseases of Fish. Vet Clinics of North America Small Animal Pract. 18(2): 401-426.
2. Stoskopf, M. 1988. Taking the History. Vet Clinics of North America Small Animal Pract. 18(2): 283-391.
3. Lewbart, G.A. 1991. Medical Management of Disorders of Freshwater Tropical Fish. Compendium on Continuing Education. 13(6): 969-977.
4. Lewbart, G.A. 1995. Emergency pet fish medicine. In Current Veterinary Therapy XII (Kirk and Bonagura eds.), Saunders, CO, 1369-1374.
5. Stoskopf, M.K. 1995. Anesthesia of pet fishes. In Current Veterinary Therapy XII, (Kirk and Bonagura eds.), Saunders Co., 1365-1369 .
6. Anderson, WG, McKinley, RS, and M Colavecchia. 1997. The use of clove oil as an anesthetic for rainbow trout and its effects on swimming performance. North American Journal of Fisheries Management,17:301-307.
7. Hikasa Y, Takase, K, Ogasawara, T, and Ogasawara S: Anesthesia and recovery with tricaine methanesulfonate, eugenol and thiopental sodium in the carp, Cyprinus carpio. Japanese Journal of Food Microbiology, 4:161-166, 1986.
8. Campbell, T.W. 1988. Fish Cytology and Hematology. Vet Clinics of North America Small Animal Pract. 18(2): 349-364.
9. deGuzman E. and E.B. Shotts. 1988. Bacterial Culture and Evaluation of Diseases of Fish. Vet Clinics of North America Small Animal Pract. 18(2): 365-374.
10. Shotts. E.B. and G.L. Bullock. 1976. Rapid Diagnostic Approach in the Identification of Gram Negative Bacterial Diseases of Fish. Fish Pathology. 102: 187-190.
11. Huml, R.A., Khoo, L.H., Stoskopf, M.K. and L.J. Forrest. 1993. Radiographic diagnosis. Vet. Rad. and Ultrasound, 34:178-180.
12. Lewbart, G.A., Stone, E.A. and N.E. Love. 1995. Pneumocystectomy in a Midas cichlid. JAVMA, 207(3):319-321.
13. Harms, C.A., Bakal, R.S., Khoo, L.H. Spaulding, K.A. and G.A. Lewbart. 1995. Microsurgical excision of an abdominal mass in a gourami. JAVMA, 207(9):1215-1217.
14. Love NE, and GA Lewbart. Pet fish radiography: technique and case history reports. Veterinary Radiology & Ultrasound, 38(1):24-29, 1997.
These notes are a modified and updated version of a paper which appeared in the Journal of Small Exotic Animal Medicine, 1992.
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