The small size of zebrafish present some unique challenges, and at the same time advantages for conducting necropsies and other diagnostic tests. Our knowledge of the diseases of zebrafish is relatively limited compared to those of food fishes reared in aquaculture. Until recently, most diseases of zebrafish that have been documented are those that are common problems in ornamental freshwater fishes (e.g., Piscinoodinium). With other fish species, it was not until they were recognized as commercially important and reared in captivity in large numbers did specific diseases become recognized.

Viral diseases are often devastating in aquaculture enterprises. However, as many viruses are very host specific, they are usually first identified only when a fish species is reared in large numbers in captivity. Identification of "new" viral diseases in fishes often requires establishment of new cell lines from the host under investigation. At present, viruses are not recognized as causes of disease in zebrafish, and thus zebrafish are not routinely examined for viruses. However, zebrafish cell lines are available.

Likewise, with bacterial infections, aside from mycobacteriosis, only common, opportunistic Gram negative bacteria such as Aeromonas hydrophila and surface infections by gliding bacteria are occasionally encountered. Given the above, our primary approach for diagnostics has been histopathology. This allows for interpretation of pathological changes at the tissue and cell level. Moreover, the small size of zebrafish allows for preparation of the entire fish on one slide.

Click for high resolution image

Sagittal section of whole zebrafish with mycobacteriosis G = granulomas in the kidney and viscera due to the bacterial infection

 The mere presence of a pathogen in a sick fish does not necessarily mean that it is the cause of the disease. It is important, therefore, to conduct a thorough investigation, beyond just detecting the presence or absence of known pathogens, to determine the cause of the disease. Histological examinations, in addition to gross necropsies and in vitro culture of microorganisms, are often required to determine the association of the observed pathogens in the disease being investigated.

Diagnosis of a disease begins at the research facility. Some of the most crucial data are collected by the staff maintaining the fish by thorough record keeping and careful observations of affected fish. The following information is important for disease diagnosis and implementing control strategies. Astrofsky et al. (2002) recently published a guide for conducting clinical investigations on laboratory zebrafish.

To assist with obtaining an accurate diagnosis, the following information should be obtained: origin (stock and strain), previous disease problems in the affected population, and husbandry conditions (e.g., water hardness, diet, medication history, system design, and mortality rate).

Fish behavior may be useful for indication of an emerging disease and some behavioral changes are useful for presumptive diagnoses. For example, if fish are flashing (rubbing on the sides of tanks), this may indicate that they are infected with external parasites. Other indications of disease include cessation of feeding, lethargy, and abnormal position in the aquarium (e.g., at surface or at the bottom). Abnormal respiratory pattern may indicate gill damage, and whirling or spiraling swimming often indicates neurological damage.

Examinations should be conducted on diseased fish that are collected while still alive (moribund). It is important to determine the primary cause of mortality and to differentiate this from secondary or opportunistic pathogens that may have taken advantage of already diseased fish. To accomplish this, several affected fish should be examined whenever possible. It is also useful to include apparently normal, asymptomatic fish so that early pathological changes and the underlying cause of morbidity can be determined. Dead fish may be suitable for some parasitological examinations and for observing obvious macroscopic pathological changes. However, bacteriological examinations conducted on dead fish can yield misleading results, and many histological changes are obliterated by post-mortem autolysis.

The small size of zebrafish makes conducting a necropsy somewhat tedious, but with patience and practice this can be efficiently accomplished with the aid of a stereo dissecting microscope. Note surface abnormalities (e.g., frayed fins, cloudy eyes, ulcers, skin discolorations, parasites, and tumors). Prepare wet mounts of the skin mucus and a few scales by scraping the surface of the fish with a coverslip and placing the coverslip on a glass slide. Some water may be added to the preparation so that the area between the slide and the coverslip is completely filled with liquid. Examine the wet mount with a compound microscope, starting with low power. Reducing the light and lowering the condenser will produce higher contrast, which will make microscopic parasites and other pathogens more visible.

Gills

Remove the operculum. Note color of gills (pale gills usually indicate anemia). Check for parasites, cysts, excessive mucus, and hemorrhages with a dissecting microscope. Prepare a wet mount by removing a few filaments with scissors, placing the filaments in a large drop of freshwater on a glass slide, and overlaying with a coverslip. Examine for small parasites, fungi, and bacteria using a compound microscope.

Internal Examination

Open the visceral cavity. Note if ascites, hemorrhages or other abnormalities are present. Expose the kidney by removing the swim bladder and note any kidney abnormalities. Many diseases cause enlargement or discoloration of the kidney, but this may be difficult to see in zebrafish due to their small size. Examine the heart for any abnormalities. Multiple, whitish cysts in the visceral organs is suggestive of Mycobacterium infections. Examine squash preparations of organs with a compound microscope to detect encysted parasites, fungi or granulomas. Squash preparations are made by removing a small piece of tissue, and gently squashing it between a slide and coverslip so that a thin preparation suitable for examination with a compound microscope or dissecting microscope is made.

It is critical to fix fish for histology as soon as possible after fish are killed to avoid post-mortem changes. If possible, do not use dead fish because significant autolytic changes may occur in 15-20 min after death. Fish are preserved in formaldehyde based fixative such as Davidson’s or Dietrich’s solutions for histology. Open the visceral cavity of the fish by cutting and removing a small section of the abdominal wall. Then place fish whole in the fixative at approximately 1:20 (v/v) tissue to fixative. We have found Dietrich’s or Davidson's solution to be the best all around fixative for processing whole zebrafish for histological examination. The entire fish can be processed (usually cut in sagittal sections) and representative organs can be visualized in multiple sections on one or two slides.

Electron Microscopy

Under special circumstances, electron microscopical examinations may be warranted. The general principles for histology applies for collecting specimens for electron microscopy, but freshness of tissues at fixation and proper infiltration of tissues is even more critical. Therefore, small pieces of tissue should be minced in cold glutaraldehyde based fixative into pieces about 2 mm³ maximum. The fixed tissue is stored overnight in this solution, then transferred to the appropriate buffer solution. Samples in EM fixatives and buffers should be refrigerated. There are many EM fixatives and buffers. The Appendix provides a recipes for Millonig’s buffer and fixative. Transmission electron microscopy can be performed with limited success on tissues fixed in neutral buffered formalin, but is very poor with acidic fixatives, such as Davidson's or Dietrich’s solutions.

Some bacterial infections, such as surface gliding bacteria and Mycobacterium spp., can be identified by simple Gram or acid fast stains (e.g., Ziehl Neelsen). However, the diagnosis of other bacterial diseases requires isolation of the bacteria in culture. In this case, live fish should be delivered to the laboratory, but this may not be practical in some situations. For this reason, methods for obtaining initial cultures are provided below. The bacterial cultures can then be sent to a laboratory for complete identification.

Use only freshly sacrificed fish for bacteriological examinations as dead fish from the tanks are essentially worthless. Disinfect the surface of the fish with 70 % ethanol. Flame-sterilize the dissecting instruments. Two methods are used for obtaining inocula from the kidney. For the visceral method, open the visceral cavity by making an incision in abdominal wall. Make sure not to cut into the gastrointestinal tract. Then push aside the swim bladder with flame-sterilized forceps and insert a sterile swab or loop into the kidney. For the dorsal method, enter the kidney by lateral incision dorsal to the kidney (i.e., near the lateral line). Streak the specimen on Tryptic Soy Agar or blood agar bacteriological plates, seal the plates, keep at a room temperature, and send the plates to the microbiology laboratory for further diagnosis.

For gliding bacteria (Cytophaga, Flexibacter and Flavobacterium spp.), we recommend Cytophaga Medium. Although the lesions may exhibit massive numbers of gliding bacteria, other bacteria (e.g., Aeromonas spp.) may overgrow the former. Therefore, the best way to obtain pure cultures of gliding bacteria is to homogenize the infected tissue (e.g., gills, skin and muscle) in sterile water, and inoculate plates in serial log dilutions.

Gram-stained preparations may reveal bacteria when they are numerous in infected tissue. Smear or imprint suspect tissues thinly on a glass slide, air dry and fix the slide by gently heating the slide over an open flame for 3-5 seconds. Gram stain kits are available from scientific supply houses and include instructions for their use.

As with bacterial diseases, isolation of viruses in culture may be required to diagnose a viral disease, and culture is best conducted on tissues collected from freshly killed fish. If this is not practical, the fish should be refrigerated for no longer than 24 h before examination. As a last resort, fish for virus examination can be frozen. The specimens are then transported to a qualified fish virology laboratory. At present, there are no known viral diseases of zebrafish colonies. However, as has been the case with other relatively new forms of fish culture, as fish health researchers thoroughly investigate outbreaks and incorporate tissue culture in their examinations, it is very likely that "new" viral diseases will be recognized in zebrafish.

Molecular Biology

DNA-based diagnostic tests utilizing polymerase chain reaction (PCR) are becoming increasingly common in fish health diagnostics because they are very sensitive and specific. With zebrafish, we have developed a PCR test for the neural microsporidian Pseudoloma neurophilia, and Astrofsky et al. (2000) used PCR tests to identify and differentiate different species of Mycobacterium. Because of the extreme sensitivity with PCR, great care should be taken to avoid cross contamination between samples. Bleach instruments between samples. Samples are either frozen immediately or preserved in ethanol. Mixed results are achieved when using formalin preserved tissues for genotypic analysis. Moore et al. (2001) found that decalcification procedures, which are routinely used when processing zebrafish for histology, are particularly detrimental for molecular analyses.

Transport of Specimens

It is not always possible to obtain live or fresh specimens. If only dead fish can be provided for laboratory examination, they should be refrigerated and examined within 24 h. If fish cannot be delivered to a laboratory within 24 h, then fish should be preserved by fixation in tissue preservatives for histology unless otherwise indicated. Each method of preservation has certain advantages and disadvantages, as indicated in following table.

Chronic, systemic bacterial infections by various Mycobacterium species are frequently diagnosed in aquarium fishes. The species most commonly found in fishes are M. marinum, M. abscessus, M. chelonae, and M. fortuitum. These species, as well as M. peregrinum and M. haemophilum, have been associated with mycobacteriosis in zebrafish (Astrofsky et al. 2000; Kent et al. 2004).

Usually mycobacteriosis is chronic with low level mortality, but in certain circumstances, infections may be very acute and result in severe epizootics in zebrafish colonies. Factors that are most responsible for the differences seen in the severity of outbreaks in fish mycobacteriosis have yet to be determined. It is not know if highly virulent strains or species cause the acute outbreaks, or if the outbreaks occur in fish that are immunocompromised by other factors, such at poor water quality, excessive crowding and handling, etc.

Mycobacterium spp. of fishes can infect humans. In most cases, the infections are confined to the extremities, and though not usually life-threatening may require aggressive antibiotic treatments to resolve (Kern et al. 1989; Shih et al. 1997; Hoyen et al. 1998). However, these infections may be lethal to immune compromised individuals (Lessing et al. 1993). Considering the zoonotic potential of Mycobacterium spp. from fishes, those handling aquarium fishes, including zebrafish, should wash their hands after coming in contact with water containing fish, and avoid exposing open lesions to aquarium water and fishes. Most cases of human mycobacteriosis associated with fishes are caused by M. marinum. This species is considered not to grow, or to grow very poorly, at 37 C. The majority of M. marinum cases are not life threatening and are confined to extremities, where body temperatures are lower. However, some strains of M. marinum are capable of growth in culture at 37 C, and we demonstrated that others not capable of growth at this temperature in media can grow at 37 C in macrophage cell lines and in mice (Kent et al. 2006).

Clinical Signs and Gross Pathology. Clinical and macroscopic changes associated with Mycobacterium may be variable, dependent upon the site and extent of the infection, and on the "chronicity" of the infection. Clinical signs include lethargy, anorexia, and emaciation. Fish may exhibit ulcers, hemorrhage or hyperemia around the head (similar to Gram negative septicemia), raised scales, frayed fins, and pallor of the skin or gills. A hallmark macroscopic change is the presence of multiple, white nodules in various visceral organs. These are visible by close examination of organs with a dissecting microscope, but may be hard to find in small fish, such as zebrafish.

Astrofsky et al. (2000) described various forms of the disease as they occur in zebrafish. Infected fish showed decreased overall reproductive capabilities, as well as characteristic changes such as dropsy (generalized edema with swollen abdomens) and frank ulcers of the skin.

Microscopy: In "typical" fish mycobacteriosis (i.e., the chronic form), histological examination reveals numerous granulomas, often with necrotic centers, throughout the visceral organs. These can be seen in wet mount preparations, but can be confused with encysted parasites. Staining of these tissues with acid-fast stains will often reveal acid-fast (red) bacilli in the lesions. In more aggressive infections, associated with rapid mortality, we have seen massive accumulations of macrophages replete with bacteria throughout the viscera (severe, chronic peritonitis) and occasionally extending into the muscle. Some of these cases are very acute and exhibit few, if any, granulomas. We often visualize mycobacteria in the swim bladder wall and intestinal epithelium, indicating that these are likely sites for initiation of infections.

Diagnosis. Preliminary diagnosis is achieved by observing multiple granulomas in visceral organs in either wet mounts or histological sections. These may also be caused by fungal or parasitic infections. Confirmatory diagnosis is requires visualization of acid fast bacilli in either histological sections or in tissue imprints. Often zebrafish are decalcified during processing for histology, and we recently found that certain aggressive decalcification procedures may inhibit the acid-fast staining of Mycobacterium spp. in tissue sections (Kent et al. 2006).

Some Mycobacterium species are relatively difficult to grow in culture, and thus this procedure is infrequently employed with fish.

PCR tests for screening for the presence of the causative agent have been described (Colorni et al. 1994; Astrofsky et al. 2000). Such tests, while extremely sensitive, usually do not provide information on disease status or the severity of infection, but may be useful for screening for carrier/subclinical infections. We have also used PCR tests to obtain mycobacteria sequence directly from infected fish, which is very useful for species identification of bacteria that are difficult to grow in culture (Kent et al. 2004; Poort et al. 2006).

Control and Treatment. The infection is usually very difficult to eradicate in fish culture systems with antibiotics. However, Conroy and Conroy (1999) reported that kanamycin at 50 ppm with 4 doses 2 days apart apparently controlled the disease in guppies. Boos et al. (1995) treated firemouth cichlids and Congo tetras orally with rifampicin and tetracycline with some success. Nevertheless, depopulation of affected tanks and avoiding cross contamination has been the most effective way for controlling the infection in aquarium fishes (Astrofsky et al. 2000).

Because the disease is rather insidious, the infection is wide spread in aquarium fishes, and treatment with antibiotics is difficult, the best approach is to avoid the infection. This is accomplished by strict quarantine procedures and understanding a thorough history of the stocks introduced into research facilities. In addition, optimizing water quality and husbandry conditions will reduce the likelihood of epizootics occurring when a few carry fish are present in the population.

Several species of gliding bacteria (i.e., Flavobacterium, Flexibacter or Cytophaga spp.) caused surface infections in freshwater and marine fishes. One of these diseases, bacterial gill disease (BGD), is an infection caused by a variety of these bacteria, precipitated by crowding and poor water quality. Therefore, BGD is also called "environmental gill disease". Two gliding bacteria, Flavobacterium columnare and F. branchiophilum, are species specifically incriminated with the disease in freshwater. Other bacteria may cause or be secondarily involved with the lesions. Key predisposing factors are high organic loads and possibly high ammonia in the water. Therefore, the condition is often found in high production trout farms. With zebrafish, we have seen BGD following shipment in which the fish were in transit for a long time.

Clinical Signs and Gross Pathology. Typical of most gill diseases, fish exhibit labored breathing and may accumulate at the surface with BGD. Fin and tail infections (known as fin and tail rot), is characterized by erosion of the fins. Infected skin usually appears white due to loss of the overlying epidermis and exposure of the dermis.

Microscopy. Wet mount examinations of infected gills reveal fused secondary lamellae due to epithelial hyperplasia. Masses of filamentous bacteria are observed on the surface of the gills. Likewise, masses of bacteria are seen in wet mount preparations of infected skin or fins. Phase contrast microscopy is useful for visualization of the bacteria. Histological sections reveal severe epithelial hyperplasia of the gills and occasionally necrosis. Careful examination usually reveals mats of bacteria on the gill surface.

Control and Treatment. The best way to avoid BGD and fin and tail rot is to avoid overcrowding and maintain proper water quality. When shipping, assure that fish are not feed for a few days before transport, maintain lower densities in shipping bags, and select the fastest route of travel.

The most common chemical treatment for BGD in salmonids has been chloramine T at about 6-15 ppm as a 1h bath (Bowker and Erdahl 1998). Repeated treatments may be required. Lumsden et al. (1998) compared this drug with hydrogen peroxide for treating experimental infections in trout, and ultimately recommended 100 mg/L hydrogen peroxide (1 h baths) as an effective treatment as well. However, we have not tested hydrogen peroxide on zebrafish.

Piscinoodinium pillulare, the cause of velvet disease, is a yellowish, parasitic dinoflagellate in which one stage infects the skin and gills of fish. The parasite can multiply very rapidly in aquaria and will rapidly kill fish. This is a common pathogen of ornamental fishes. The dinospore stage infects fish, and transforms into a trophont stage that attaches to the surface, especially gills, and apparently feeds on the host’s epithelium. After several days, this stage leaves the host and forms a tomont. This off-host stage undergoes several divisions, resulting in 256 infective motile dinospores which go on to infect other fish. The life cycle is completed in about 2 wk under optimal conditions (ca 23-26 ^oC).

Clinical Signs and Gross Pathology. Heavily infected fish exhibit lethargy, may hang near the surface of the water column, and exhibit labored breathing. Fish may exhibit a greyish to rusty color sheen on the surface.

Diagnosis. The infection is diagnosed by observation of the trophonts in either wet mounts or histological sections. For the former, Piscinoodinium is distinguished from many other surface-infecting protists by it’s lack of motility.

Control and Treatment. We do not have personal experience treating Piscinoodinium infections with zebrafish. Noga (1996) recommended prolonged immersion in salt at 1 teaspoon/5 gal. Very heavy, life-threatening infections may be treated with a quick (1-3 min) bath in full strength sea water (35 ppt). As with Ich, the presence of off-host developmental stages should be considered when implementing a treatment regime. Therefore, it is best to remove all the fish, treat them in a new aquarium, and disinfect the original tank.

Neural microsporidiosis in zebrafish was first reported in 1980 in France (de Kinkelin 1980). The parasite has now been identified in zebrafish at many research and commercial facilities, and was assigned to a new genus and species, Pseudoloma neurophilia by Matthews et al. (2001). The infection is linked to severe emaciation (often referred to as ‘skinny disease’), but its precise role in every case (primary cause or secondary opportunist) has yet to be resolved. In a survey of a population with a known high prevalence of infection, 97% of the skinny fish and 30% of the normal fish were infected. This demonstrates that, as with many parasites, the infection may be also found routinely in normal, healthy appearing fish. Conversely, there are several other causes of emaciation in zebrafish. Handling and crowding stress causes increased severity of the infection, along with reduced growth and fecundity (Ramsay et al 2009).

Transmission. Microsporidia are obligate intracellular fungus-like parasites with a complicated life cycle. The life cycle concludes with the production of an infectious and resistant spore, which is the only stage of the parasite that can live outside of the host cell. With most microsporidia, transmission occurs via the ingestion of the infective spore stage, and we have found the infection in exposed adult fish as soon as 4 wk after feeding infected tissues. Fry fish are very susceptible, and may show a more acute form of the disease, with high mortality about 1 wk post exposure (Ferguson et al. 2007). While the parasite is easily transmitted by feeding on infected carcasses, we have found that it is also transmissible when fish are kept physically separate, but in the same water. We presume this is by infected egg material, etc. being released from infected females, as zebrafish frequently spawn under natural aquarium conditions.

There have also been reports of vertical transmission in other microsporidia of both vertebrates and invertebrates (Bandi et al. 2001; Dunn et al. 2001; Phelps et al. 2008). We detect the spores of Pseudoloma in the ovaries and sometimes within immature and degenerate eggs of zebrafish, and the role of vertical transmission or pseudovertical transmission (transmission to progeny with the pathogen outside of the egg) of Pseudoloma is currently under investigation.

Clinical Disease and Gross Pathology. Emaciation and spinal curvature, such as scoliosis, are common in infected fish. However, emaciation and spinal curvature are nonspecific signs with numerous etiologies. Furthermore, clinically normal zebrafish may also harbor heavy P. neurophilia infections. Therefore, diagnosis of microsporidiosis should not be based on clinical signs alone.

Microscopy. The primary site of infection is the central nervous system (spinal cord and hindbrain) and nerve roots. The infection often extends to the skeletal muscle, where it causes severe, chronic, multifocal inflammation. It also infects the ovaries and eggs, and occasionally other organs, such as the kidney (Kent and Bishop-Stewart 2003).

Click for high resolution image	Clinical signs of microsporidiosis.  A. emaciation ("skinny disease")  and  B. scoliosis.
Click for high resolution image	Pseudoloma from brain. A. Giemsa stain B. Wet mount with Nomarski's phase interference Bar = 10 µm.
Click for high resolution image	Pseudoloma in histological sections.  A. Sagittal section showing spinal cord. X = xenomas in spinal cord. Xenomas also occur in ventral nerve roots (arrows) with associated inflammation (In).  B. High magnification of xenomas in spinal cord.
Click for high resolution image	Pseudoloma in histological sections – other sites of infection and other stains. A. Low magnification showing inflammation in skeletal muscle. Bar = 100 µm.  Inset at upper left shows macrophage with 4 spores B-D. Spores (arrows) in eggs. B) H&E. C) Acid fast, spores fill a degenerate egg, D) Gram, showing blue-staining spores. E. Spores in spinal cord stain blue with the tissue Gram stain.
Click for high resolution image	Fungi-Fluor stained histological sections.  A. Low magnification showing numerous spores (stained blue with DAPI filter) in spinal cord xenomas (arrow).  B. High magnification of spores (S) in ovaries.

Control and Treatment. Ultraviolet light (UV) is used to control pathogens in zebrafish systems. UV sterilization has been successfully employed to kill viruses, bacteria, and protozoa in water before coming in contact with fishes, in both flow through and recirculating systems. Most systems use UV sterilizers following biological filtration. Other microsporidian species have been shown to be very susceptible to UV treatment, and our observations at ZIRC strongly indicate that UV at 30-50,000 µWsec/cm² kills the parasite and thus prevents it transmission in a recirculating system. Fumagillin has been widely used as an oral treatment for fish microsporidiosis, usually with good success (see review by Shaw and Kent 1999). Fumagillin was first developed for treating Nosema apis infections in honey bees. Kano et al. (1982) reported that fumagillin was effective against the microsporidium H. anguillarum in eels (Anguilla japonica). Since this first report on treating microsporidiosis in fish with fumagillin, the drug has been used to treat N. salmonis infections in Chinook salmon (Hedrick et al. 1991) and Loma salmonae infections in Chinook salmon (Kent and Dawe 1994). Our initial investigations with this drug were not successful for controlling or reducing the infection in zebrafish, and we plan to repeat the study using higher doses as we found no side effects with the drug.

Microsporidian spores may be resistant to disinfectants (Shaw et al. 1999), and Ferguson et al. (2007) found that chlorine levels routinely used by zebrafish facilities (i.e., 25 or 50 ppm for 10 min) was not effective for killing spores. Moreover, we have observed eggs filled with spores of Pseudoloma in histologic sections of ovaries (see figures). The spores within intact eggs could be protected from chlorine, even if this concentration is effective for killing spores.

Based on our understanding of the parasite at this time, the following practices would likely reduce the spread and impact of the infection 1) ensure that water in recirculating systems is adequately treated with UV sterilization, 2) with existing infections, remove and euthanize all emaciated and moribund fish as soon as possible to prevent cannibalism and further transmission, 3) consider screening of brood fish, particularly females, for the parasite to reduce the risk of transferring the infection into a facility with their progeny, 4) rear potentially infected fish in optimum conditions and reduce stress to minimize the impact of the infection, and 5) minimize potential for cross-contamination of stocks by assigning dedicated populations of wild-type fish for out crossing to each mutant stock. After breeding, these wild-type fish should not be returned to a common stock tank nor be crossed to any other mutant stock.

Recently we have diagnosed another microsporidium, Pleistophora hyphessobryconis, in a few zebrafish research facilities. This parasite, the cause of neon tetra disease, infects a wide variety of aquarium fishes, including tetras, barbs and goldfish (Lom and Dyková 1992). The infection results in massive numbers of developing parasites and spores throughout the skeletal muscle (see figure below), and spores are found in other organs within macrophages. This parasite certainly should be avoided in zebrafish research facilities, and thus we recommend extreme caution using zebrafish that have come in contact with other aquarium fishes, particularly tetras. Diagnosis, control and treatment are similar as those for Pseudoloma, but a PCR test for Pleistophora hyphessobryconis in not presently available.

The species infecting zebrafish was first identified as Pseudocapillaria tomentosa by Dr. R. Overstreet, University of Southern Mississippi (pers. comm.), and the specimens that we have evaluated from zebrafish are consistent with this identification. Capillarids infect all classes of vertebrates, and are often pathogenic due to their invasive nature (Moravec 1987; Moravec et al. 1987a ). Many species have been described from fishes, including aquarium species (Williams and Jones 1994).

Pseudocapillaria tomentosa (junior synonym P. brevispicula) has a broad host specificity, infecting some 25 fishes in the family Cyprinidae and members of other orders such as Aguilliformes (eels), Gadiformes (cod fishes), Salmoniformes (salmon) and Siluriformes (catfishes) (Moravec 1987). Pseudocapillaria tomentosa was associated with mortality in captive tiger barbs (Puntius tetrazona) (Moravec et al. 1984) and related parasites cause disease in other aquarium fishes. Capillaria pterophylii has been recognized for many years as a common pathogen of captive angelfish and discus fish (Richenbach-Klinke 1952) and Capillostrogyloides ancistri is highly pathogenic to the bushymouth catfish Ancistrus dolichopterus (Moravec et al. 1987b). Lomankin and Trofimeko (1982) showed that oligochaetes (e.g., Tubifex tubifex) can serve as paratenic hosts for P. tomentosa in laboratory transmission studies. In the same study, they demonstrated that direct transmission in the absence of worms is also a route of infections. Recently, we confirmed that the parasite can be transferred from fish to fish in the absence of oligochaete worms, which demonstrates that the parasite can proliferate in zebrafish research facilities.

Clinical Signs and Gross Pathology. Heavily infected fish are often dark, emaciated, and lethargic. Necropsy may reveal liver enlargement and anemia.

Diagnosis. Identification of capillarid nematodes is most easily accomplished by the presence of unique eggs, which are oval and contain distinctive bipolar plugs. Precise identification to the species level requires careful examination of the male sexual organs, which are rather diminutive in Pseudocapillaria spp. Females of P. tomentosa are about 7-12 mm, males about 4-7 mm. To date, the only nematode infection that we have identified in zebrafish from research facilities is P. tomentosa.

Control and Treatment. Oligochaete worms may be a source of the infection, and thus should be avoided as food, particularly if their source is unknown. As direct transmission occurs between fish, the infection can spread within a population if not controlled. If fish are not highly valuable, the most appropriate choice would be to eliminate the infected population. At present, the infection is not wide spread in research facilities, perhaps due to bleaching of eggs and quarantine procedures.

Ivermectin is a very effective antihelminthic for many nematode and arthropod infections in terrestrial animals, and has been employed for treating nematode infections in fishes (Heckman 1985). Therefore, ivermectin should be considered as a possible treatment for P. tomentosa in zebrafish. However, this drug has not been tested with zebrafish, and there appears to be a great variability is the tolerance of the drug between even closely related fish species – i.e., many fish species are highly susceptible to toxic side effects of ivermectin at doses used in mammals (Johnson et al. 1993: Kilmartin et al. 1997), presumably because of a comparatively reduced blood/brain barrier causing neurotoxicity. This drug has also been associated with respiratory problems in fish (Toovey et al. 1999).

Oral or bath treatments with levamisole and fenbendazole have also been employed to treat nematode infections in fish (Noga 1996), but Hoffman (1982) found that levamisole was not effective for treating capillarid infections in golden shiners. Brood stock zebrafish treated with levamisole have become sterile (D. Weaver, Scientific Hatcheries, Huntington Beach, California, pers. comm.). Pack et al. (1995) reported that a mixture of trichlorofon and mebendazole in the form of Fluke-Tabs (Aquarium Products, Glen Burnie, MD) added to water eliminated the infection. Treated fish gained weight and when examined later the infection was not observed. Dr. Weaver has reported success with treating P. tomentosa infections using pyrantel and garlic powder incorporated in the feed in a production situation.

Click for high resolution image	Wet mount preparations of Pseudocapillaria tomentosa. 1. Posterior of male with diminutive bursa (arrow). Bar = 25 µm. 2. Posterior of male with smooth spicule (arrow). Bar = 25 µm. 3. Female with distinct stichocysts with alternating light and dark bands. Bar = 100 µm. 4. Eggs with distinctive bipolar plugs. Bar = 100 µm.
Click for high resolution image	Histological sections of zebrafish with Pseudocapillaria tomentosa infections. Diffuse, severe chronic inflammation, extending from lamina propria (LP) into to visceral cavity.  Pa = inflammation in pancreas.  Arrow = nematode. Hematoxylin and eosin.  Bar = 100 µm

Renal Myxosporidiosis

Myxozoan parasites (phylum Myxozoa, class Myxosporea) are common parasites of cold-blooded vertebrates, particularly fishes (Kent et al. 2003). Traditionally the Myxozoa have been classified with the Protozoa. However, about 15 years ago analysis of small subunit ribosomal DNA revealed that the Myxozoa rooted within the kingdom Animalia. Two viewpoints have predominated in these discussions. One suggests that myxozoans are most closely related to the Cnidaria (diploblasts), due to the morphological and developmental similarities of their polar capsules and nematocysts respectively. These observations and phylogenetic analysis in support of them were first presented in a key publication by Siddall et al. (1995). Other analyses support phylogenetic proximity to the bilaterian animals (triploblasts) (Zrzavy et al. 2003).

There are over 2,200 described species of myxozoans, and most are relatively non-pathogenic (Lom and Dyková 2006). Those that that infect lumina of organs are called coelozoic species. These are commonly found in the gall bladder, bile ducts, or urinary tract, and they often cause little tissue damage. Histozooic species (those occurring deep within tissues) usually form small, confined white cysts with little associated tissue damage. However, when these cysts are numerous in vital organs, such as the gills or heart, they can cause disease.

The life cycle of myxozoans is complicated (see figure below). They contain several vegetative stages (trophozoites) and development in the fish culminates in the formation of multicellular spores, called myxospores. Spore morphology is the primary criterion used for identification of myxozoans. Development in an aquatic oligochaete or polychaete is required to complete the life cycle (Kent et al. 2003). The forms found in oligochaetes were originally considered to be different parasites, which were assigned to the class Actinosporea. Because these stages are not separate taxa from myxozoans found in fishes, the taxonomy of the phylum Myxozoa has been revised (Kent et al. 1994).

Whereas serious myxozoan diseases are found in wild or cultured fish with access to open waters, they are less often seen in laboratory systems. This is likely because of the requirement of the parasite to pass through an annelid worm to infect another fish. We have, however, detected a myxozoan, belonging to the genus Myxidium or Zschokkella in zebrafish from several research facilities.

Microscopy. Histological sections of infected fish reveal the parasite in the common mesonephric ducts and occasionally in the lumen of kidney tubules. Infections may be heavy, but are seldom associated with significant histological changes. The organism seen in zebrafish is comprised of a multicelluar trophozoite (plasmodium) in which careful examination reveals myxospores inside. The hallmark identifying character of myxospores is the presence of polar capsules, which can be better visualized with special stains such as methylene blue and Giemsa. The genera Myxidium or Zschokkella have members with polar capsules located on the opposite end of the spores.

Diagnosis. The infection is diagnosed by observation of the parasite in histological sections. Wet mounts of tissues are often used to visualize myxospores.

Control and Treatment. Infections in a zebrafish facility indicate that the oligochaete host is present in the water system (e.g., tanks or filters). Indeed, we have seen reports of small oligochaetes (e.g., Stylaria spp.) in zebrafish system. In addition, oligochaetes used to feed fish (e.g., black worms, red worms) may be sources of myxozoan infections to fishes held in aquaria (Hallett et al. 2005). Fumagillin and a few other drugs show variable efficacy for treating myxozoan infections. Treatment of the zebrafish myxozoan would probably not be warranted as it is not very pathogenic.

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This condition occurs in many species of fish held in captivity, and we occasionally observe it in our diagnostic cases. Nephrocalcinosis is the accumulation of calcium deposits in kidney tubules and collecting ducts. Causes of nephrocalcinosis include high CO₂ (e.g., > 12 mg/L) in water or excessive levels of calcium and magnesium in the diet. The use of calcium carbonate (rather than sodium bicarbonate) to buffer water in recirculating systems has been associated with the condition (Chen et al. 2001). Only when the lesions are extensive is the condition associated with overt clinical disease. In other words, we see many fish with the condition (diagnosed by histology) in which it was probably not a significant cause of the disease.

Clinical Signs and Gross Pathology. We have not observed macroscopic changes due to nephrocalcinosis in zebrafish. However, with larger fish, kidneys with severe nephrocalcinosis may exhibit distinct, white, opaque deposits in the kidney. The ureters may be filled with a chalk-like material, and vermiform deposits may occur in collecting ducts and tubules.

We frequently observe severe, chronic inflammation in the visceral cavity associated with degenerating eggs in diagnostic cases. Rarely, in the most severe cases, aggressive fibroplasia leading to development of fibromas and fibrosarcomas may be observed. We have named the condition "Egg Associated Inflammation" or EAI. Anecdotal information indicates that it is caused by egg retention, and infectious agents are not usually found in the lesions. Occasionally we observe Mycobacterium spp. within granulomas in these lesions, but these infections are probably not the primary cause.

Clinical Signs and Gross Pathology. Female zebrafish typically present with an enlarged or distended abdomen. The ovaries grossly appear as a solid, tumor-like mass in the visceral cavity. In several cases the mass will cause adhesion to the wall of the visceral cavity, and ultimately a free raft of scar tissue will be extruded through the body wall and skin of the fish. This results in a large ulcer with a whitish center on the lateral flank of the fish.

Microscopy. Histological examination revels severe, chronic inflammatory changes that originate in the ovaries and may extend throughout the peritoneal cavity. Eggs of varying states (from intact to completely degenerated) are found within the lesion. Prominent fibroplasia occasionally occurs, and in some cases appears to lead to the formation of fibrosarcomas.

The second histological figure (B) shows a mass of degenerating eggs and fibrotic tissue that has extended through the body wall. This same phenomenon occurs in wild striped bass with chronic pertionitis associated with larval tapeworms of Lacistorhynchus tenuis (Moser et al. 1984).

Diagnosis. Identification of the condition is based on observing histological changes described above.

Control and Treatment. The precise cause of EAI is unknown. Because abnormal egg retention is a suspected cause, we recommend timely spawning of females. Holding males and females together may also aid in preventing abnormal egg retention. Striping egg bound females has been tried, but apparently is not reliable for curing the problem.

Clinical Signs and Gross Pathology. We have not related direct clinical disease to hepatic megalocytosis, and it is only observed during histological examinations.

Microscopy. Histological sections of the liver reveal massive hypertrophy of the cell and nucleus of individual hepatocytes, at times 10 to 20X, or greater, than normal size. The abnormal hepatocytes are usually found randomly dispersed within the normal liver parenchyma.

Many research facilities use flow through or partially closed systems in which dechlorinated city water is used as the water supply. This certainly is an advantage for avoiding many pathogens. However, failure of the dechlorination system often results in severe mortality due to either acute or chronic chlorine toxicity. Fish are very susceptible to chlorine. City water usually contains a minimum of 0.2mg/L total chlorine but usually ranges between 0.5 –1.0 mg/L (Noga ). Chlorine is also routinely used to disinfect tanks and equipment, and inadequate rinsing or ventilation is another potential source of chlorine toxicity.

Clinical Signs and Gross Pathology. Chlorine causes severe, acute necrosis of the gills, and thus fish suffer rapid respiratory distress and asphyxiation. Fish may exhibit laboured breathing, and petechial hemorrhages around the head. Cherry-red, swollen gills are characteristics of chlorine toxicity

Microscopy. Microscopic examination shows severe necrosis of the gills.

Diagnosis. Chlorine toxicity is diagnosed by detecting elevated chlorine levels in the water. "After the fact" exposure is suspect in fish showing the signs described above.

Control and Treatment. Chlorine is avoided by removal with charcoal filters (for running water). For static systems, chlorine can be removed by adding sodium thiosulfate at 7.4 ppm/chlorine ppm). Mortalities in exposed fish can be reduced by adding salt to the water and by assuring that the water is adequately aerated. In addition, fish should be immediately transferred to chlorine-free, aerated water.

Microscopic Changes. Chronic ammonia toxicity is often associated with hyperplasia of the gill epithelium. However, some researchers report that this histopathological change is caused by other agents (either abiotic or biological) that are often associated with husbandry conditions (e.g., excessive crowding, inappropriate water exchange) that lead to elevated ammonia levels (Lang et al. 1987).

Diagnosis. As the clinical signs and pathological changes of ammonia toxicity are rather non-specific, identification of the problem relies on detection of elevated ammonia levels.

Control and Treatment. Improving water flow and reduction in fish densities are the immediate solutions to NH₃ toxicity_. Changing water or increasing water flow usually increases pH, and ammonia is more toxic at higher pH. Ammonia toxicity often occurs in new recirculation systems with biological filters as it may take several weeks to establish adequate denitrifying bacteria in the filters. Therefore, fish should be added gradually to new systems, with close monitoring of ammonia levels.

Nitrite toxicity is less common in aquaria situations than ammonia toxicity, but can occur simultaneously. It is more common in ponds – e.g., nitrite toxicity is the cause of brown blood disease in catfish, in which case methemoglobin production by chemical oxidation of heme iron results in haemoglobin incapable of combining with oxygen, thus hypoxia occurs (Speare 1998a). Levels of nitrite should be kept below < 0.10 mg/L.

Clinical Signs and Macroscopic Changes. Severe, acute GBD may cause macroscopically visible bubbles in the skin, fins or gills. The eyes are also a target organ, in which visible bubbles form in the retrobulbar tissues. Fish may exhibit exophthalmia. Unfortunately, many cases of GDB do not present with specific clinical or pathological changes – i.e., fish die without visible bubbles in the tissues. Fish may exhibit exophthalmia. Fish that recover sublethal GDB may be susceptible to infectious diseases.

Microscopy. When present, histological sections may reveal bubbles, particularly in the retrobulbar tissues. Bubbles occur within blood vessels and cause vascular occlusions, leading to formation of thrombi and possibly hemorrhage. Gas bubbles may be visible in gill blood vessels in wet mounts.

Click for high resolution image	Zebrafish with gas bubbles around eye due to GBD
Click for high resolution image	Retrobulbar tissue of the eye from opakapaka (Pristipomoides filamentosus), a deep water snapper, with GBD.  H = hemorrhage I = chronic inflammation B = clear spherical spaces where bubbles occurred