Management of Fish

Fish are poikilothermic, and all physiologic processes are greatly influenced by water temperature. In freshwater, the internal tissues of fish are hyperosmotic to the environment, whereas in saltwater they are hypoosmotic. Surface injuries to the skin make osmoregulation more difficult and may be of serious consequence because of the loss of fluid balance and circulatory collapse.

The structure of the fish kidney varies with the species; generally it is divided into an anterior “head” kidney, which is usually located anterior to the gas bladder, and a posterior “caudal” kidney, which is retroperitoneal and ventral to the vertebral column. Hematopoietic, renal, and endocrine tissues are found in the kidney, with hematopoietic tissue located cranially and excretory tissue located caudally. Divalent ions are excreted principally via the kidney, and monovalent ions and nitrogenous excretions via the gills. Accordingly, lesions of the kidney and gills may seriously interfere with respiration, excretion, and fluid balance.

The gas bladder (also known as the swim or air bladder) in bony fish, which originates as an appendage of the foregut, regulates body buoyancy and may also be used for sound production. Physostomous fish have an open connection between the swim bladder and the GI tract, whereas physoclistous fish do not. Although a single chamber in most fish, the gas bladder consists of dual chambers in cyprinids and three chambers in cod and suckers belonging to the genus Moxostoma.

A sensory lateral line system along the sides of the body and head receives stimuli from the aquatic environment and mediates adaptive responses through the CNS.

Fish depend on increases in environmental temperature for efficient antibody production during infections (or after vaccinations), when most pathogens are replicating at a more rapid rate. The optimal temperature for antibody production varies with the species of fish (tropical, temperate, or coldwater). Extremes in environmental temperature (above or below that of the natural habitat) inhibit antibody production. Like T lymphocytes in other vertebrates, those in fish are responsible for cell-mediated immunity. Immunity is not as age dependent in fish as it is in other animals; young fish are usually immunocompetent and can be vaccinated successfully. Antibodies are found in the mucus of the fish skin and GI tract.

Although vaccination of fish against specific diseases has been economically important in preventing losses, there is a need for improved methodology. Advances include increased use of autogenous vaccines; several companies will work with veterinarians and their clients to develop custom vaccines for specific situations. A few vaccines (eg, for Aeromonas salmonicida) are available or in development for pet fish, particularly koi.

As with all species, a good history is critical to establish a diagnosis. Questions of particular interest for fish cases include the number of animals affected, whether one species or multiple, the chronicity of the problem, and a thorough description of animal housing and care, including the volume and design of the system (see Aquatic Systems), number and size of animals stocked, species, new additions, use of quarantine, and previous medications. Owners can be asked to bring fish and water samples into the clinic, or the practitioner may wish to visit the site. Site visits allow the system to be more accurately evaluated and the behavior of fish readily observed. If fish are brought to the clinic, the owner should provide an animal showing the signs of concern. A live animal can be transported in a cooler with a battery-powered aerator. A separate water sample should be provided in a plastic bag and transported on ice. A minimum of 1 quart of tank water should be requested for analysis. If desired for recovery after anesthetizing the animal, a larger volume of tank water may be requested. Recently deceased specimens have diagnostic value and may be submitted to the veterinary clinic, or the owner may be directed to submit these to a laboratory experienced in fish necropsy and diagnostic testing (see below). Water samples should be submitted with necropsy specimens.

Although the same principles are used in necropsy of fish as in other animals, great emphasis is placed on an accurate and thorough history, premortem signs, fresh necropsy material, and direct microscopic examination of fresh tissue smears and squash preparations. Fish decompose quickly, and many saprophytic microorganisms reproduce rapidly in the decaying tissues, which complicates isolation of pathogens unless samples are collected immediately after death. A general fish necropsy may include blood collection (premortem); biopsy of gill, skin, and fin tissues; bacterial or viral culture of internal organs; and histology. A veterinary clinic or diagnostic facility familiar with fish necropsy protocols and aquatic microbiology should be used. Whenever possible, live fish should be submitted. If the fish has just died, the eyes should be clear and the gills normal in coloration and texture. There should be no “dead fish” smell; however, some moribund fishes may have a strong odor. Freshly dead fish should be placed in a sturdy plastic bag and submitted on ice. A water sample should always be submitted with the fish. An animal that has died and been placed in a freezer has limited diagnostic value, but freshly dead frozen fish may be useful for bacteriology, virology, or toxicology testing.

Fresh tissue samples of gill filaments, skin mucus, and fins should be collected, prepared as a wet mount, and examined under a light microscope at 40×, 100×, and 400×. Fresh water should be used to prepare wet mounts of external tissues from freshwater fish, and saltwater should be used to prepare wet mounts from marine fish. If uncertain, water from the tank or from the submitted water sample should be used. Ensuring that the salinity used to prepare mounts is similar to the salinity present in the environment should allow organisms to remain viable long enough for identification. Distilled water should not be used for tissue samples. Tissue should be examined for morphology and for the presence of parasites, bacteria, or fungal elements. Microscopic examination of internal organs is also recommended if the fish has been euthanized. Unstained sections of stomach and intestine should be examined for presence of parasites, and the lower intestine should be examined for flagellates. Unstained sections of spleen, anterior and posterior kidneys, and liver should be examined for presence of parasites, granulomas, or other anomalies. Wet-mount examination of fish tissues is crucial for diagnosis of most parasites.

Blood can be collected from a number of sites, and hematologic parameters measured. In fish >25–100 g, depending on species, the caudal vessels of the caudal peduncle, the duct of Cuvier (common cardinal vein), and the dorsal and ventral aortas are easily accessible. For smaller specimens that are to be euthanized, blood can be collected in a hematocrit tube immediately after euthanasia by severing the caudal peduncle and exposing the caudal vessels. Use of hematology and serum chemistry is limited, because normal values for many fish species are not readily available; however, the information may still be clinically useful. Lithium heparin is the anticoagulant of choice for most fish species, although EDTA is preferred for ictalurid catfishes, and plasma may be used for biochemistry tests. Serology may be diagnostic in certain cases (eg, heavy metal toxicity). Whole blood (1–2 drops) can be incubated in brain heart infusion broth at room temperature on an electric rotator. If cloudiness indicative of bacterial growth develops, a loop of the blood-broth mix can be used to attempt primary isolation of a systemic bacterial pathogen.

Fish should be euthanized and opened under sterile conditions. A sterile swab of posterior kidney, or other organ of interest, may be shipped to a laboratory in transport media, but primary isolation directly onto an enriched media (eg, tryptic soya agar enriched with 5% sheep blood) is preferable. Although blood agar supplemented with salt is helpful for marine fish, it is not necessary if an enriched blood agar is used. Ordal’s or similar cytophaga media should be available for isolation of myxobacteria (slime bacteria, including Flavobacterium columnare). Sabouraud agar is an excellent all-purpose media for isolation of fungal agents. Lowenstein-Jensen or Middlebrook media is recommended for isolation of Mycobacterium spp. Mueller-Hinton is the media of choice for susceptibility testing of most common bacteria isolated from fish. If abscesses or other obvious anomalies are visible, those sites should also be cultured. In general, bacterial or fungal cultures taken from fish tissue should be incubated at room temperature (25°C). Some agents of concern will not grow at all at 37°C, the standard temperature for incubation of cultures taken from mammals. Agents of zoonotic concern, such as Mycobacterium, can be dual incubated at both 25°C and 37°C. An acid-fast stain should be available for bench-top staining of granulomatous material which, when positive, is strongly suggestive of Mycobacterium. If fish are seen spinning or showing other behavioral indications of neurologic disease before death, brain cultures are indicated.

If viral disease is suspected, appropriate tissues may be collected. Specimens should include both fresh tissues placed in reagent ethanol and frozen tissues. Several viral diseases are of regulatory concern to veterinarians practicing fish medicine in the USA (see Table: Fish Diseases of Regulatory Concern in the USA).

Tissue sections no larger than 1 cm³ should be placed in 10% neutral buffered formalin for histopathology. After 24 hr of fixation, they should be removed and placed in reagent alcohol in case molecular diagnostic techniques will be required.

Therapy for pet and ornamental fish is often based on environmental management followed by the use of targeted therapy to control specific pathogens. Use of prophylactic medication in the absence of diagnostic testing is strongly discouraged and may contribute to resistant bacterial infection and other complications.

Drug therapy can be provided via several routes of administration, including exposure by bath (adding medication to water), medicated feed, injection, or topical administration. Generally speaking, bath and topical treatments are most useful for external infections, whereas medicated food and injection are most appropriate for internal infections.

Using a bath treatment requires accurate measurement of the volume of water to be treated. Volume of a rectangular tank is calculated by multiplying measurements of the length, width, and depth. Volume of a circular tank is calculated by multiplying 3.14 by the radius squared by the depth. To calculate directly into liters, measurements should be in cm and multiplied by 0.001. If the measurements are made in feet, the result will be in cubic feet; cubic feet can be converted to gallons by multiplying by 7.481. If the container is oddly shaped, volume may be calculated mathematically, but it may be easier to purchase a flow meter to measure the volume required to fill the tank. Alternatively, the volume of inflowing water per minute can be measured by determining how long it takes to fill a 1-L graduated cylinder (or 5-gal. bucket). Using that information, determining how long it takes to fill a tank or ornamental pond can provide a fairly accurate assessment of volume.

Some medicated feeds can be purchased commercially for aquaculture or pet fish. Custom-made medicated feeds can be prepared for use in ornamental fish. Flake, pellet, or gel diets can be used as a base for medicated food for pet fish. Cooking oil spray is an effective binder for use with pelleted or flake foods for ornamental fish. The addition of medication to commercially available gel diets is easily done as the gel cools. In general, medication should not be added when the gelatin is hot, because some medications, particularly oxytetracycline, are heat labile.

Injections can be given either IM or intracoelomic (IC). IM injections are given in the epaxial muscles, lateral to the dorsal fin. Injection of some drugs can cause muscle necrosis, so it is important to alternate injection sites and to limit injections to every 3 days unless required more often. To administer IC injections, fish should be placed head downward in dorsal recumbency to move internal organs away from the injection site, which should be anterior to the anus, just off the ventral midline. Topical treatments, usually in ointment form, should be applied directly to the external lesion(s). The fish can be manually restrained, usually without removing the entire animal from the water. The area to which the treatment is applied should be held out of the water for several seconds (<1 min) to allow some absorption before returning the fish to the water. Repeated applications may be necessary.

Sunburn can occur in surface-swimming fish or can be induced (even in bottom-dwelling species) by feeding photodynamic drugs such as phenothiazine, although ultraviolet light penetrates water poorly. Affected fish will have visible lesions along the dorsal surface. Providing shade solves the problem.

Increasingly, surgery is an option for management of some medical problems in pet or exhibit fish, including neoplastic disease, failure to ovulate (ie, “egg bound” fish), and gas bladder repair for buoyancy problems. Fish skin does not have the subcutaneous tissue that provides flexibility to domesticated animal tissue; therefore, wounds are not usually treated by surgical closure but instead allowed to heal by second intention. The clinical evaluation before surgery is similar to that in other species, although there may be more emphasis on imaging and less on blood work. Radiology and ultrasound work very well in fish, and use of these techniques before an invasive surgical procedure is recommended.

Descriptions in the literature show how a fish may be positioned for a surgical procedure. For smaller animals, such as koi, a foam “bed” is easily constructed and can be covered with something as simple as clear plastic wrap so that the fish does not lose skin, scales, or mucus from direct contact with the foam. The “bed” can be positioned over an aquarium using a plastic tray with holes to allow water drainage. A pump can move water containing an anesthetic solution (usually MS-222) from the aquarium through a small tube or catheter and across the fish’s gills. Ideally, dissolved oxygen, ammonia, and pH should be monitored in the anesthetic solution. The fish can be covered with a clear plastic surgical drape (avian drapes work well), followed by fairly minimal preparation of the incision site. Plucking scales along the incision line and carefully cleaning the area with a sterile swab soaked in sterile saline may be all that is required. Very dilute disinfectants such as povidone-iodine or chlorhexidine can be used, but if the fish is clean this is probably not necessary (or recommended) because the normal mucus has significant antibacterial properties.

Absorbable sutures are generally not recommended in fish, because they may persist for significant periods of time (>1 yr in some cases). Monofilament material and a needle with a cutting edge generally work well. The simple interrupted suture pattern works well to close fish skin, but other techniques can be used successfully. Skin sutures should be removed when the incision site has healed, generally at 3–4 wk. Surgical staples have been used successfully in fish. Results using cyanoacrylate tissue adhesive have been mixed because of a significant inflammatory reaction seen in some species. If postoperative antibiotics are used, enrofloxacin delivered via IC injection at a dosage of 5 mg/kg is a good option but can only be given to non-food fish. Butorphanol given at 0.1–0.4 mg/kg, IM, and meloxicam at 0.15 mg/kg, IM, have been used for postoperative pain control in non-food fish. Increasing the salinity in freshwater systems to 1–3 ppt (g/L) is strongly recommended during the recovery and healing periods. Most freshwater fish should be able to tolerate a salinity of 3 ppt (3 g/L) on an almost indefinite basis.

Quarantine is strongly recommended for pet fish and certainly for animals intended for display in public aquaria. A 30-day period is the minimum time for quarantine, but longer periods may be necessary. Quarantined animals should be handled only when contact with all other animals is finished for the day. Quarantined animals should have their own designated equipment (ie, nets, buckets, siphons, etc) and be kept separate from other animals. Disinfectant should be used on equipment and in footbaths at entry points to the quarantine area.

When receiving fish, a thorough history should be obtained regarding previous treatment or disease outbreaks. Fish should be examined early in the quarantine period; a visual examination may suffice, but for valuable specimens a full clinical examination, including recording the weight of the animal and completing gill, skin, and fin biopsies, is recommended. Moribund fish should be examined, and dead fish necropsied. Prophylactic use of antibiotics may be warranted in some shipments, especially recently imported marine specimens. Treatment with praziquantel for monogeneans is often prudent with marine fish as well. Goldfish commonly have significant monogenean infestations, and treatment with formalin or praziquantel may be appropriate. Koi should be quarantined to prevent introduction of koi herpesvirus (KHV), a serious and reportable disease, to established populations. Koi should be quarantined for a minimum of 30 days at a temperature of 75°F (24°C). Fish that become ill during quarantine should be tested for KHV (see Koi Herpesvirus).

Quarantine is not useful for some pathogens, but that does not mean it should not be done. Examples are mycobacteria, which at this time cannot be detected with nonlethal tests, and many viruses, microsporidia, and myxozoans. These diseases are usually detectable using necropsy techniques if quarantined animals that die are thoroughly examined. Quarantine is most useful for detection of external parasites and some internal parasites that can be detected by examination of feces.

New tank syndrome (see Aquatic Systems:Nitrogenous Compounds) usually occurs within the first 6 wk after a new fish hobbyist sets up a tank. Owners may report everything was fine for several weeks, and then the fishes suddenly become lethargic, anorectic, and frequently die. Although parasites may be present in newly purchased fishes, it is critical to perform a complete analysis on a water sample from the affected tank. Often total ammonia nitrogen (TAN) or nitrite, or both, are high enough to result in toxicity. Treatment should include decrease in feeding, water changes, the addition of chloride for nitrite toxicity, and evaluation of adequate biofiltration. It can take up to 8 wk for a biofilter in a tropical fish tank to become established. New tank syndrome can be avoided by several methods. One method, known as fishless cycling, is to completely set up the tank but with no fish addition; ammonia is added to achieve a concentration of at least 1–5 mg/L. This will initiate the “cycling” process, which should be monitored by regular testing for ammonia, nitrite, and nitrate. Once no ammonia or nitrite is present, fish can be added safely. Another method is to slowly add several fish to the tank over several months, although this can result in high ammonia or nitrite levels if not done carefully.

Old tank syndrome (see Aquatic Systems:Nitrogenous Compounds) can occur when water changes are small and infrequent. In some cases, no water is exchanged, and the practice of “topping off” is done by adding water to the tank to replace what has evaporated. Although this can occur in any tank, it is more frequent in tanks that are inhabited by large, carnivorous fishes. These fishes eat large, high-protein meals, frequently in the form of other fish, and consequently excrete large amounts of TAN. Consequently, the biofilter uses up bicarbonate to transform TAN to nitrite and nitrate, and the bicarbonate depletion decreases total alkalinity. This results in either a gradual or dramatic drop in pH. If the former, the fishes will adapt to the slow change often with no outward sign of distress. If the pH decline is rapid, the result could be death for the resident fishes. The water quality parameters of total hardness, nitrate, total alkalinity, and pH are often abnormal. Total hardness (TH) and nitrate are usually significantly higher than normal. TH of the tank water should be compared with TH of the water used for water changes. Total alkalinity and pH are usually significantly lower. Often TAN will be high because of inadequate bicarbonate for the nitrogen cycle. If the fish are still alive, the water quality parameters should be returned to normal by performing daily small water changes to avoid pH shock. From that point on, the tank should receive regular, large water changes to prevent a recurrence of the problem.