Water Quality and Idiopathic Diseases of Laboratory Zebrafish

C H A P T E R

39 Water Quality and Idiopathic Diseases of Laboratory Zebrafish Katrina N. Murray1, David Lains1, Sean T. Spagnoli2 1

Zebrafish International Resource Center, University of Oregon, Eugene, OR, United States of America; 2Biomedical Sciences, Oregon State University College of Veterinary Medicine, Corvallis, OR, United States of America

Introduction Diagnosis of non-infectious diseases often involves reviewing a history of dietary, environmental, and water parameters, assessing gross and behavioral clinical signs, and evaluating microanatomic changes in tissue sections. A thorough evaluation of these parameters can also be important in assessing whether identified infectious agents are in fact opportunistic and secondary to environmental or physiological changes. Suboptimal water quality can induce a range of signs from subclinical to severe morbidity and mortality with acute to chronic effects. Many laboratory zebrafish facilities monitor water parameters electronically and utilize handheld meters and assays for back-up and periodic confirmation of computerized readouts. The ability to do on-demand testing of water parameters at different locations on and off the water system is important because the parameters to which the fish are exposed can vary depending on multiple factors. On a recirculating water system parameters may be affected by time of day, demand for water on the system, and whether water is flowing through new equipment or areas that have been stagnant for long periods of time. Water parameters on a flow through system may vary with fluctuations at the water source, especially seasonally. Microenvironments like Petri dishes, glass bowls, shipping bags, and spawning tanks should also be considered when investigating pathologies associated with suboptimal environmental parameters. In this chapter, we review the most frequently encountered non-infectious diseases of laboratory zebrafish. Diseases of water quality include ammonia and nitrite toxicity, chlorine and chloramine exposure, heavy metal toxicities, supersaturation, and

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nephrocalcinosis. Idiopathic diseases and those with multiple potential non-infectious etiologies are also discussed. These include egg-associated inflammation, spinal deformities, operculum malformations, hepatic megalocytosis, cardiac pathologies, tissue hyperplasia, and fin lesions.

Diseases of Water Quality Ammonia Toxicity Description. Ammonia is released into the aquatic environment during the breakdown of organic material and as a waste product of fish metabolism. In water, ammonia exists as NHþ 4 and NH3. The unionized form predominates at higher pH and temperature and is considered most toxic to fish, as it more readily crosses the gills. However, once it has crossed the gills, NH3 is converted to NHþ 4 , which is more toxic in vivo, damaging cellular structures, and metabolism (Rosenberg, 2012). Elevated ammonia results in increased tissue consumption of oxygen coupled with decreased transport of oxygen to the tissues (Schwedler, Tucker, & Beleau, 1985). Pathobiology and Clinical Signs. Clinical signs of acute ammonia toxicity in fish include hyperventilating, or piping, at the surface, being in lateral recumbency on the bottom of a tank, rapid and erratic swimming, anorexia, and mortality (Daoust & Ferguson, 1984). Convulsions have also been described in trout (Smart, 1978). Chronic exposure to elevated ammonia can cause immunosuppression and impede growth (Colt & Armstrong, 1979; Walters & Plumb, 1980). Prolonged exposure is often associated with gill epithelial hypertrophy and hyperplasia with the fusion of lamellae in

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FIGURE 39.1 Gill epithelial hypertrophy and hyperplasia with the fusion of secondary lamellae. Scale bar is 200 mm in A and 20 mm in B. Sagittal section with H&E stain.

severe cases (Fig. 39.1). However, this gill lesion has not been reproduced in studies that selectively expose fish to high levels of ammonia (Braun, Steele, & Perry, 2009; Daoust & Ferguson, 1984), suggesting the epithelial change is a nonspecific effect of poor water quality, of which elevated ammonia is often a component. Diagnosis. Ammonia toxicity is diagnosed by measuring unionized ammonia in system, tank, or microenvironment water. Many test kits simply report total ammonia nitrogen (TAN), but the amount of NH3 can be calculated if pH and temperature are known (see Table 29.1 in Water Quality for Zebrafish Culture chapter). Ideally, zero ammonia should be measured in the system water. Even 0.02 mg/L of NH3 can be harmful and greater than 1.0 mg/L is usually lethal (Meade, 1985). Control and Treatment. Treatment should involve immediate efforts to reduce measured ammonia and correcting the source of the problem. Reducing unionized ammonia can be achieved by increasing the frequency of water changes, adding an ammonia adsorbent (e.g., zeolite), and using ammonia-neutralizing agents (e.g., ClorAm-X). The percentage of total ammonia in the toxic unionized form will decrease with reductions in pH and temperature and with higher salinity (Wedemeyer, 1996). However, changes in other parameters should be made cautiously and gradually as rapid changes can result in stress, immunosuppression, and even death. Long-term reduction of ammonia may entail improving biological filtration and evaluation of stocking densities, live food preparations, decomposition of uneaten food, and static water locations.

Nitrite Toxicity Description. During nitrogen cycling, nitrites rise in a water system following an ammonia peak. Therefore, nitrite toxicity may occur alone or with ammonia

toxicity. Chloride cells transport nitrite across the gills (Bath & Eddy, 1980) to the bloodstream where it oxidizes the ferrous iron (Feþ2) in hemoglobin to ferric iron (Feþ3), forming methemoglobin, which does not bind oxygen. Furthermore, the presence of methemoglobin also increases the affinity of the remaining hemoglobin for oxygen, reducing its ability to release and deliver oxygen to tissues. Pathobiology and Clinical Signs. Nitrite toxicity, or methemoglobinemia, is also referred to as brown blood disease because the conversion of red hemoglobin to brown-colored methemoglobin can give blood and gills a brown appearance. Clinical signs associated with nitrite toxicity are typical of hypoxia. Fish may be lethargic, remain near the water inlet, and hyperventilate in well-oxygenated water (Kroupova, Machova, & Svobodova, 2005; Lewis & Morris, 1986). If ammonia toxicity is concurrent, gills are likely to have been damaged, impairing oxygen absorption and exacerbating general hypoxia. Nitrites can also induce hypertrophy and increased turnover of chloride cells, possibly to maintain normal body chloride levels in spite of competition by nitrites for uptake (Gaino, Arillo, & Mensi, 1984). Long-term exposure to nitrites can impair growth rates in zebrafish (Voslarova, Pistekova, Svobodova, & Bedanova, 2008). Mortalities are typically attributed to tissue hypoxia. Arillo, Gaino, Margiocco, Mensi, and Schenone, (1984) proposed that liver hypoxia and dysfunction are central to nitrite-induced mortality. As in other fish, zebrafish larvae are less sensitive to higher nitrite concentrations than older fish, which is likely a function of gill development and nitrite uptake (Voslarova, Pistekova, & Svobodova, 2006). Similar to ammonia toxicity, the poor water quality associated with elevated nitrites may result in hypertrophy and hyperplasia of gill epithelial cells (Wedemeyer & Yasutake, 1978).

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Diagnosis. Diagnosis is made by observation of clinical signs of methemoglobinemia and measuring elevated nitrites in the water. Nitrites should be kept below 0.1 mg/L (Wedemeyer, 1996). Control and Treatment. Nitrite toxicity is primarily a problem of recirculating water systems where biological filtration facilitates nitrogen cycling. Nitrites can be reduced by increasing water changes and adding chloride, which will compete with nitrite for uptake by the gills. Bowser, Falls, Vanzandt, Collier, and Phillips, (1983) recommend 3 mg of chloride to 1 mg of nitrite for treating channel catfish. Correcting the instigating cause is equally important. Improving biological filtration and examining stocking densities and feeding protocols should be considered. Once nitrites are removed, methemoglobin is reduced to nearly normal levels of hemoglobin in 24 h (Huey, Simco, & Criswell, 1980).

Chlorine and Chloramine Toxicity Description. In zebrafish facilities, a dilution of household bleach (sodium hypochlorite) is commonly utilized to surface disinfect embryos, and higher concentrations are used to disinfect equipment. Chlorine is also used as a disinfectant in some municipal water supplies. When bleach is added to water, hypochlorous acid (HOCl) and hypochlorite ion (OCl) are formed. HOCl predominates in zebrafish system water, where the pH range favors the unionized form. Pathobiology and Clinical Signs. Both HOCl and OCl are strong oxidizers that cross cell membranes and damage structures, enzymes, and nucleic acids, ultimately destroying the gills (Wedemeyer, 1996). Since HOCl is neutral, it crosses cell membranes more freely than OCl and is, therefore, more toxic. Chlorine also reacts with ammonia and nitrogenous compounds to form chloramines, which are more toxic than chlorine and also induce gill necrosis. Clinical signs of chlorine and chloramine exposure include respiratory distress and mortality. Diagnosis. Diagnosis of chlorine toxicity is made by measuring chlorine in the water. Chlorine is acutely toxic and should be undetectable in zebrafish water systems. Control and Treatment. Chlorine from municipal water can enter a zebrafish water system if carbon filters are damaged, expired, or improperly installed. Fish may also be exposed to chlorine if bleach used for embryo and equipment disinfection is not properly neutralized. If chlorine toxicity is suspected, fish should be immediately moved to chlorine-free water. Sulfur compounds can be utilized to neutralize chlorine. Sodium thiosulfate should be used at a ratio of 7.4 to 1 ppm chlorine (Jensen, 1989). However, this treatment will also release ammonia from chloramines, and therefore, concurrent ammonia treatment may be warranted. Some ammonia

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binders, e.g., ClorAm-X, will also bind chlorine and chloramines. Activated carbon will convert chlorine and chloramines to carbon dioxide and ammonium salts (Wedemeyer, 1996). Chlorine will also dissipate naturally, although this process is slow and 20 h should be allowed per mg/l of chlorine. Vigorous aeration can speed dissipation of chlorine, but not chloramines.

Heavy Metal Toxicity Description. Zebrafish may be exposed to deleterious amounts of heavy metals through water or feed. A common route is by leaching from copper, lead, and zinc-coated (galvanized) plumbing components. Heavy metals are more toxic in low alkalinity, soft water, due to increased solubility. High temperature and low pH also increase metal solubility. Cadmium, zinc, copper, and nickel are likely absorbed via calcium pathways (Alsop & Wood, 2011; Hogstrand, Verbost, Bonga, & Wood, 1996; Niyogi & Wood, 2004). Although metal exposure reduces the uptake of Ca2þ, acute metal toxicity seems to be from loss of total body Naþ (Alsop & Wood, 2011). Chowdhury, Girgis, and Wood, (2016) showed that copper decreases Naþ influx and increases Naþ efflux across the gills in rainbow trout by inhibiting branchial Naþ, Kþ ATPase activity. Pathobiology and Clinical Signs. Numerous nonspecific clinical signs have been associated with metal toxicities. Gill pathologies include increased mucus production and epithelial changes involving separation from pillar cells, hyperplasia, swelling, necrosis, and desquamation (Authman, Zaki, Khallaf, & Abbas, 2015; Farrell, Ackerman, & Iwama, 2010). Copper exposure has been associated with an increase in gill chloride cells and decrease in mucous cells (Baker, 1969), as well as degeneration of chemoreceptors and mechanoreceptors (Gardner & Laroche, 1973). Gill alterations associated with metal toxicities result in decreased surface area for gas-exchange and osmotic dysregulation, leading to respiratory distress. Lead and mercury decrease acetylcholinesterase activity in zebrafish brains after 48 h exposure, but activity returns to normal by 30 days of chronic exposure (Richetti et al., 2011). Unlike other metals, iron toxicity is due to a direct effect of iron oxide precipitation on the gill surface (Abbas, Zaghloul, & Mousa, 2002; Peuranen, Vuorinen, Vuorinen, & Hollender, 1994). Other organs affected by metal toxicity include kidney, liver, and muscle. Reproduction, growth, and immune response may be compromised (Authman et al., 2015). In zebrafish, metal exposure has been shown to decrease embryo survival and hatching (Dave & Xiu, 1991). Zebrafish exposed to harmful levels of chromium through contaminated nonhatching, decapsulated Artemia cysts (decaps) produced orange embryos that

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had reduced survival rates (Tye, Montgomery, Hobbs, Vanpelt and Masino, 2018). Hexavalent chromium (Cr6þ) induces toxicity as a powerful oxidant that crosses cell membranes. Toxic effects are observed in gills, liver, and kidney (reviewed in Authman et al., 2015). Diagnosis. Heavy metal toxicosis is diagnosed by measuring toxic levels of metal in system water, feed, or fish tissue. Toxicity will vary depending on the hardness, alkalinity, and amount of organic material in the water. Submitting fish tissue for metal testing may be especially useful as water and feed levels can vary and could be within normal limits during sampling. Samples should be submitted to specialized laboratories with experience in this kind of testing. Control and Treatment. Treatment involves identifying and removing the source of the metal contaminants. For removing circulating metals, ion exchange filters and metal chelators can be utilized. However, they will also bind Ca2þ and Mg2þ, which are essential to fish survival and may need to be added back during water treatment. Ultraviolet sterilizers may break down metal-chelate complexes if they are recirculated. Reverse osmosis and carbon filters will also remove heavy metals from the water supply. Environmental factors that impact metal toxicity, like water hardness, temperature, and pH, should also be considered. Increasing calcium hardness significantly decreased copper toxicity in juvenile channel catfish (Chowdhury et al., 2016; Perschbacher & Wurts, 1999). In rainbow trout, the effects of calcium were independent of alkalinity, suggesting that the protective effect is from calcium competing for binding sites rather than a decrease in copper bioavailability (Chowdhury et al., 2016). We observed a similar protective effect of calcium at the Zebrafish International Resource Center, where the addition of a water softener in a flow-through quarantine room lowered dissolved calcium, which significantly increased the toxicity of copper leaching from pipes.

exposed to supersaturated water. Fish blood and tissues quickly equilibrate with the partial pressures of dissolved gases in the surrounding water and become supersaturated as well. Once exposed to atmospheric pressure, gases come out of solution, forming bubbles on tank surfaces and in fish tissues. Gas emboli in the vasculature can result in occlusion, tissue necrosis, and mortality (Smith, 1988). In acute stages, bubbles form in well-perfused tissues, like the brain, gills, and vasculature (Fig. 39.2). However, gas can also be lost more easily from these tissues. In chronic stages, bubbles are more likely to be found in poorly perfused tissues like fat, where bubbles are slower to develop and resolve (Machado, Garling, Kevern, Trapp, & Bell, 1987; Strauss, 1979). While gas bubble disease is a population-level problem, not all fish in a supersaturated tank will develop clinical signs. There can be great variability in disease, especially at low levels of supersaturation. The reaction to supersaturation can differ for fish of different size and age. But even within and between tanks of the same age and size, some fish will develop gross lesions

Supersaturation and Gas Bubble Disease Description. Water is supersaturated with gas when the total pressure of dissolved gases exceeds atmospheric pressure. Supersaturation can occur when saturated cold water, which holds more dissolved gas than warm water, is pumped into a sealed system and warmed under pressure. System water can also become supersaturated if there is a leak on the suction side of a centrifugal pump, which will draw air into a pressurized system. Air trapped in distribution plumbing can also dissolve as water sits in the pipes and pressure builds. When the water in an aquaculture system becomes supersaturated, it can be deleterious to fish. Pathobiology and Clinical Signs. Gas bubble disease refers to the tissue changes that can occur in fish

FIGURE 39.2 Gas bubble disease. Bubbles (*) in the heart (A) and the head vasculature (B). Scale bars are 100 mm. Sagittal sections with H&E stain.

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while others appear unperturbed (Bouck, 1980; Weitkamp & Katz, 1980). The level of supersaturation and exposure time will influence the degree of associated fish morbidity and mortality. In a recirculating aquaculture system this may be influenced by the time of day and demand on the water system. Water that sits under pressure for extended time periods may contain more dissolved gases than when water demand is high and consistently flowing through the system. We have seen gas bubble disease in zebrafish exposed to water that had been sitting, unused, in a pipe for several weeks. Under pressure and over time, the water in the pipe may have become supersaturated by pockets of trapped air. The formation of bubbles in the tissue will alter the buoyancy of fry and may drive them to the water surface. Juvenile and adult fish may exhibit lethargy, altered buoyancy, disequilibrium, and reduced feeding. Adult zebrafish with gas bubble disease generally exhibit rapid respiration and hover at the bottom of the tank, in contrast to incidences of oxygen deprivation, which drive fish to the surface. Unilateral and bilateral exophthalmia and bubbles within the tissues may be grossly visible, particularly in the eyes and fins (Lund & Heggberget, 1985; Machado et al., 1987). Mortalities may occur several days after exposure to supersaturated water and may transpire without accompanying gross lesions. Long-term effects of gas bubble disease include decreased growth rate (Batzios, Fotis, & Gavriilidou, 1998) and predisposition to secondary infections due to stress and tissue damage (Speare, 1991). Diagnosis. The occurrence of gas bubbles in fish tissues is considered pathognomonic. However, supersaturation in zebrafish is frequently not associated with obvious bubbles in the tissues either on gross or microscopic postmortem examination. Therefore, history and clinical signs should be carefully evaluated. Bubble formation on tank surfaces is an important warning sign. Saturometers can be used to measure the total concentration of dissolved gas in the water. Measurements should be taken at several locations on the water system and at multiple times during the day. Fish species vary in their sensitivity to supersaturation. In general, 110% saturation is considered dangerous, but even 102% can be deleterious to salmonids (Krise & Meade, 1988). Control and Treatment. Treatment will involve identifying and correcting the source of excess dissolved gas. Removing excess gas can be accomplished by aeration and allowing water to equilibrate with atmospheric pressure before it is sent to fish tanks. Vacuum and passive degassers can also be used. Water and air should be bled out of pipes that have been out of commission for an extended period before filling fish tanks.

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Nephrocalcinosis Description. Nephrocalcinosis is the result of deposition of calcium precipitate in renal tubules and ducts. High dissolved carbon dioxide and low oxygen, magnesium deficiency (Cowey, Knox, Adron, George, & Pirie, 1977), high selenium (Hilton, Hodson, & Slinger, 1980), and arsenic (Cockell, Hilton, & Bettger, 1991) have all been associated with nephrocalcinosis. High dissolved CO2 seems to be the most important factor in the pathogenesis. Deviations in Mg, Se, and As would likely have additional nonrenal effects (Dabrowska, Meyerburgdorff, & Gunther, 1991; Sorensen, 1991). In zebrafish, nephrocalcinosis is commonly diagnosed on microscopic postmortem examination in the absence of an identifiable inciting cause and many cases may be idiopathic. Pathobiology and Clinical Signs. In zebrafish, the effects of nephrocalcinosis are generally subclinical. Lesions range from a few small deposits to large accumulations of calcareous material resulting in extensive dilation of surrounding structures (Fig. 39.3). In other fish, nephrocalcinosis has been associated with decreased growth, impaired renal function (Lall, 2010), and mortalities when combined with stressful events (Wedemeyer, 1996). Growth rate and renal function have not been assessed in zebrafish with background nephrocalcinosis, although severe cases would likely negatively impact physiology and growth. Many cases of nephrocalcinosis diagnosed on postmortem examination are unlikely to be the primary cause of disease in these fish. As in mammals, mineral deposition within the tubules in these cases is likely the result of altered renal function secondary to systemic disease or dehydration. Diagnosis. Diagnosis is made by histology and the observation of basophilic deposits in the renal tubules and/or collecting ducts. Not uncommonly, granulomas will form around large calcium deposits, and acid-fast staining should be performed in these cases to rule out renal mycobacteriosis. Control and Treatment. Lowering fish stocking density and ensuring good water exchange and aeration are important for maintaining an appropriate level of dissolved CO2. Chen, Wooster, Getchell, Bowser, and Timmons, (2001) saw a reduction in severe nephrocalcinosis in Nile tilapia after switching from CaCO3, in the form of agricultural-grade lime, to NaHCO3 to buffer the water. However, these fish were reared in a situation of high stocking density and feed rates, which typically pushed the pH below 7. On a zebrafish system, CaCO3 in the form of aragonite can be advantageous because it is slow to dissolve and will increase pH and both general and carbonate hardness. Therefore, treatment should first focus on optimizing fish density, water exchange, and aeration.

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FIGURE 39.3

39. Water Quality and Idiopathic Diseases of Laboratory Zebrafish

Nephrocalcinosis. Renal tubules are variably dilated with calcareous debris. Scale bar is 100 mm. Sagittal section with H&E stain.

Idiopathic Diseases Egg-associated Inflammation Description. It is common to find inflamed tissue associated with degenerating egg material in the ovaries of zebrafish. Lesions range from small foci of degenerating material with a mild inflammatory response to extensive, severely inflamed areas with fibroplasia and large rafts of degenerating egg debris (Fig. 39.4). External ulcers have been observed secondary to severe egg-associated inflammation and adhesions to the body wall (Kent et al., 2016). Granulomas may or may not be

present. Lesions are usually sterile, with no evidence of infectious agents. Pathobiology and Clinical Signs. The pathogenesis of egg-associated inflammation is believed to begin with egg retention, degeneration of egg material within the ovary, and a subsequent inflammatory response. Depending on the size of the lesion, zebrafish may present with distended abdomens. Similarly, fecundity may be reduced in accordance with the size of the lesion. Rossteuscher, Schmidt-Posthaus, Schafers, Teigeler, and Segner, (2008) observed spontaneous degenerative and inflammatory lesions in the ovaries of 78% (n ¼ 46) of control zebrafish in a toxicity study. They noted that

FIGURE 39.4 Egg-associated inflammation. (A) Eosinophilic, proteinaceous debris and inflammation in the caudal ovary. (B) Higher magnification of the caudal pole of the ovary. Scale bars are 0.5 mm in A and 100 mm in B. Sagittal section with H&E stain.

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FIGURE 39.5 Spinal deformities. (A) Kyphosis, (B) lordosis, (C) scoliosis, and (D) platyspondyly.

gonad histopathology is often an endpoint in toxicity experiments, and these spontaneous lesions should be considered when determining appropriate sample size and evaluating potential treatment-induced alterations. Diagnosis. Egg-associated inflammation is diagnosed by observation of inflammation in the ovary associated with degenerating egg material in histological sections. Inflammation may be mild to severe and granulomatous. Occasionally acid-fast bacilli (i.e., Mycobacterium spp.) are observed within granulomas, but it may represent secondary infection of the altered tissue. Lesions may be focal or locally extensive with inflammation extending from the ovary to adjacent tissues. Control and Treatment. Housing male and female fish together and regular spawning may prevent egg retention, which is believed to be the underlying cause.

Spinal Deformities Description. Spinal deformities include abnormal curvature and shortening from compressed vertebrae (platyspondyly). Curvature can occur in ventral (lordosis), dorsal (kyphosis), and lateral (scoliosis) directions. Since bone is metabolically active and is continually being remodeled, factors affecting bone development, growth, and repair can precipitate bone deformities at all ages. Still, ossification of bone during larval stages may make that stage especially sensitive (Nu¨sslein-Volhard & Dahm, 2002). Environmental, dietary, and genetic factors can all influence the development of spinal deformities. In particular, temperature, toxins, and infectious agents have all been linked to spinal deformities (Kent et al., 2016; Pohl, 1990; Sfakianakis, Georgakopoulou, Papadakis, Divanach, Kentouri, & Koumoundouros, 2006; Stickland, White, Mescall, Crook, & Thorpe, 1988). Several nutrients are important for skeletal development and pathology

including calcium, phosphorous, manganese, zinc, selenium, and vitamins A, D, C, E, and K (Cahu, Infante, & Takeuchi, 2003; Lall & Lewis-McCrea, 2007). Vitamin C, ascorbic acid, is a cofactor in the biosynthesis of collagen and often mentioned in the pathogenesis of skeletal deformities. Fish do not synthesize vitamin C so it must be supplied as a dietary nutrient. Traumatic events during fish handling can also result in spinal changes. Pathobiology and Clinical Signs. Spinal deformities are commonly observed in laboratory zebrafish colonies (Fig. 39.5). Curvatures may be noted as individual occurrences in mature fish or affecting large numbers of larval or juvenile fish. In aged zebrafish, spinal deformities have been attributed to degenerative changes including vertebral dislocations, thickening of bone at vertebral joints, fractures, and remodeling of vertebral bone and cartilage (Hayes et al., 2013). The authors have observed vertebral deformities with subsequent impingement of the spinal cord and nerve tracts in zebrafish that exhibited altered swimming behavior. Diagnosis. Diagnosis is made by gross and microscopic examination of spinal curvature, compression, or alteration. Control and Treatment. Multiple factors have been implicated in the pathogenesis of spinal deformities in fish. Environmental, nutritional, infectious, and genetic causes should be considered. An assessment of the onset of clinical signs, affected ages, and distribution within the population may help narrow down possible etiologies.

Operculum Malformations Description. Operculum deformities have been described in multiple intensively reared fish species with changes including outward flaring, inward curling,

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FIGURE 39.6 Operculum malformation. (A) Lateral and (B) ventral views of everted opercula. (C) Sagittal section with H&E stain. The ventral aspect of the operculum is everted, exposing the gill cavity. Scale bar is 20 mm.

and shortening (Beraldo, Pinosa, Tibaldi, & Canavese, 2003; Hilomen-Garcia, 1997; Lindesjoo, Thulin, Bengtsson, & Tjarnlund, 1994). Pathobiology and Clinical Signs. Operculum malformations are common in laboratory zebrafish and almost always involve outward curling or eversion of the opercular flap (Fig. 39.6). The deformity is usually bilateral, but unilateral changes can occur. Gills are often visible as the malformation prevents complete coverage of the gill cavity. Within a zebrafish facility, the prevalence of malformed opercula may be quite high in particular lines and absent in others. During genetic screens, Harris, Henke, Hawkins, and Witten, (2014) recovered zebrafish with opercular defects with Mendelian inheritance. Some of these mutants had general effects on skeletal development and differentiation and were not specific to the opercula. At the Zebrafish International Resource Center, we consistently see outward curling of the opercula without other grossly apparent skeletal changes in

particular wild-type lines. We performed in-crosses and out-crosses to lines with normal opercula and did not observe simple Mendelian inheritance, suggesting multiple genes are involved and/or partial penetrance. Substantial recovery of unilateral inward curling of opercula was observed in gilthead sea bream (Sparus aurata L.) over a 16-month observation period (Beraldo & Canavese, 2010). Reversal of phenotype has not been documented in zebrafish. Histologically, the operculum malformations are not accompanied by degenerative or inflammatory changes. Importantly, deformations of the operculum may decrease growth rate, and the inability to cover the gill cavity may negatively affect the ability of a fish to handle situations of poor water quality and respiratory distress (Koumoundouros, Oran, Divanach, Stefanakis, & Kentouri, 1997). Diagnosis. Diagnosis is made by gross examination and visualization of the gills where they should be covered by the operculum. Mild malformations may only be visible by histology. Control and Treatment. There is likely a strong heritable component to the outward curling of opercula observed in laboratory zebrafish. Particular lines may have a high prevalence of the malformation compared to unaffected lines exposed to the same environmental and husbandry parameters. Breeding strategies to minimize the malformation should be considered, although efforts can be confounded by the fact that the lesions appear to progress as fish mature and mild curling may not be visible until after fish have started spawning. Delaying propagation of a line until phenotypes seem fixed may be advisable. Environmental causes have been linked to operculum deformities in other fish (Hilomen-Garcia, 1997; Lindesjoo et al., 1994), and should be considered along with nutritional and metabolic causes in the case of widespread operculum deformities in laboratory zebrafish.

Hepatic Megalocytosis Description. Hepatic megalocytosis refers to the presence of greatly enlarged hepatocellular nuclei and cytoplasm components (Fig. 39.7).

Pathobiology and Clinical Signs It is not uncommon for hepatic megalocytes to be multi-nucleate, presumably due to mitotic failure. Hepatic megalocytosis and nuclear pleomorphism are considered early lesions in the progression of hepatic neoplasia in English sole exposed to polycyclic aromatic hydrocarbons (Myers, Johnson, & Collier, 2003). In laboratory zebrafish, however, no link between

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Idiopathic Diseases

FIGURE 39.7 Hepatic megalocytosis. In scattered hepatocytes nuclei and cytoplasm are greatly enlarged. Binucleate megalocytes are present. Scale bar is 20 mm. H&E stain.

megalocytosis and neoplasia has been elucidated. Hepatic megalocytosis has been described in chinook salmon from apparently unpolluted water in British Columbia (Stephen, Kent, & Dawe, 1993). Other liver and organ lesions were not present, and the prevalence of hepatic megalocytosis in wild chinook was greater than in chinook farmed in seawater netpens. Hepatic megalocytosis has frequently been associated with exposure to xenobiotics, although these cases also describe morbidity and additional microanatomic lesions. In rainbow trout, pyrrolizidine alkaloids induced hepatic megalocytosis, fibrosis, and necrosis along with venooclusive and renal changes (Hendricks, Sinnhuber, Henderson, & Buhler, 1981). A water-borne or natural food toxin was presumed to be the cause of severe liver disease in Atlantic salmon that included hepatic megalocytosis, hydropic degeneration, necrosis, gross liver changes, and mortality (Kent, Myers, Hinton, Eaton, & Elston, 1988). In netpen-reared Atlantic salmon in coastal British Columbia, microcystin toxicity was the presumed cause of hepatic megalocytosis, severe necrosis, and hydropic degeneration (Andersen et al., 1993). In contrast, hepatic megalocytosis frequently occurs in apparently normal laboratory zebrafish with no accompanying gross, behavioral, or other microanatomic changes. An association has been drawn between the occurrence of hepatic megalocytosis in zebrafish and recirculating water systems with fluidized sand filters (Spitsbergen, Buhler, & Peterson, 2012; Spitsbergen & Kent, 2003). However, these system parameters alone are insufficient for lesion induction as not all facilities with recirculating systems and fluidized sand filters

TABLE 39.1

Percentage of 8-month wild-type fish sampled at ZIRC between January 2016 and November 2017 with hepatic megalocytosis (HM).

Line

Total fish

Fish with HM

% With HM

AB

1281

453

35

WIK

91

1

1

TL

80

0

0

TU

70

0

0

SAT

110

0

0

TAB-14

45

0

0

NHGRI-1

70

0

0

have zebrafish with hepatic megalocytosis. The lesion was observed in moribund zebrafish in a new facility, where it was presumed to be associated with a toxin leaching from new plastics on the system (Kent et al., 2011). Spitsbergen et al. (2012) postulated that tumor promoters and carcinogens in recirculating systems fluctuate over time resulting in cohort variations in phenotypes, including hepatic megalocytosis and liver and gastrointestinal neoplasias. The Zebrafish International Resource Center operates on recirculating water systems with fluidized sand filters. We have noticed a particularly high prevalence of hepatic megalocytosis in the AB line compared to other wild-type lines exposed to the same water and husbandry conditions, suggesting a possible genetic component to susceptibility (Table 39.1). The effects of megalocytosis on liver metabolism and physiology are unknown; however, the possible variations in line susceptibility are worth considering when choosing a line for toxicology studies. Importantly, the observation of this abnormality in such studies should be interpreted very carefully with a close examination of and comparison to control and background livers. Diagnosis. Hepatic megalocytosis is diagnosed by histological examination of the liver and observation of hepatocytes with greatly enlarged nuclei and cytoplasm. Cells may be multi-nucleate. In zebrafish, the megalocytes are usually randomly distributed throughout the hepatic parenchyma but may be focally clustered. Control and Treatment. Toxin exposure should be considered, especially if morbidity, mortality, or other microanatomic changes are noted along with hepatic megalocytosis. When hepatic megalocytosis was associated with leaching plastics in a new zebrafish facility, installation of a carbon filter was preventative (Kent et al., 2011).

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(A)

(B)

(C)

(D)

(E)

FIGURE 39.8 Cardiac pathologies. Sagittal sections with H&E stain of (A) pericardial effusion, (B) dilated cardiomyopathy, and (C) and (D) myxomatous heart valves (*). (D) Higher magnification of the atrioventricular valve. (E) Grossly visible swelling of the cardiac area. Scale bars are 0.2 mm in A, B, and C and 50 mm in D.

Cardiac Pathologies Description. Pericardial effusion, dilated cardiomyopathies, and degenerative cardiac valves are occasionally observed in laboratory zebrafish. Pathobiology and Clinical Signs. Cardiac pathologies may result in grossly visible lesions, including swelling of the cardiac area and raised scales from secondary scale pocket edema. Accumulation of fluid in the pericardial space, dilation of one or more chambers of the heart (sinus venosus, atrium, ventricle, bulbous arteriosis), and myxomatous valvular changes may be apparent in histological sections (Fig. 39.8). Varying degrees of secondary edema may be observed in the coelomic cavity and scale pockets. Diagnosis. Diagnosis is made by examination of cardiac structures in histological sections. Sagittal sections can provide good visualization of multiple chambers, although multiple sections may be needed to see the valves. Mild dilation of one or more cardiac chambers in tissue sections is not necessarily diagnostic for dilated

cardiac disease and may instead reflect changes at euthanasia or sectioning parameters. Additional changes, like gross swelling of the cardiac area, valvular abnormalities, or scale pocket edema, may indicate that the observed dilation is pathological. Control and Treatment. Cardiac pathologies in zebrafish are generally sporadic and infrequent. If they are regularly identified in a facility, an investigation into affected stocks, ages, and distribution should be considered along with possible genetic, environmental, and nutritional factors.

Organ and Tissue Hyperplasia Description, Pathobiology, and Clinical signs. Hyperplasia of certain tissues is not an uncommon finding during routine evaluation of histological sections of zebrafish. Testis, ultimobranchial gland, biliary tree, and thyroid are most frequently affected. It is worth noting that ectopic thyroid tissue outside of the pharynx is occasionally

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Idiopathic Diseases

observed in zebrafish. Therefore, distinguishing between metastatic thyroid neoplasia and hyperplasia of ectopic tissue is a particular diagnostic challenge. Diagnosis. Diagnosis is made by histology and observation of increased size or amount of normal organ or tissue structures. Cellular characteristics should be carefully evaluated to distinguish between hyperplastic, dysplastic, and neoplastic growth. These parameters are discussed in detail in chapter 43, Nonexperimentally Induced Neoplastic and Proliferative Lesions in Laboratory Zebrafish. Control and Treatment. Genetic and environmental factors may influence the development and progression of hyperplastic growth. A high prevalence of bile duct hyperplasia has been noted in TL zebrafish relative to other wild-type lines (Spitsbergen et al., 2012). Calcium and iodine availability and metabolism should be considered when multiple fish exhibit ultimobranchial and thyroid hyperplasia, respectively. Spontaneous occurrence is also possible.

Fin Lesions/Erosion Description. Fin lesions that do not have a primary infectious etiology are often termed fin erosion, to distinguish them from fin rot. Fin erosion encompasses dermal and epidermal damage resulting in fin splitting, fraying, reduced length, nodularity, necrosis, and epithelial hyperplasia (Latremouille, 2003; Turnbull, Richards, & Robertson, 1996). Fin erosion is a common problem in intensive aquaculture. Fortunately, these lesions are not common in laboratory zebrafish. We include a description of them here because the Zebrafish International Resource Center (ZIRC) has experienced two episodes of widespread fin erosion, indicating that zebrafish populations are susceptible and preventive protocols should be considered. Pathobiology and Clinical Signs. Fin erosion is a multifactorial problem (reviewed in Ellis et al., 2008; Latremouille, 2003). Interfish aggression, abrasion from environmental surfaces, and contact from handling and transportation have all been identified as primary causes of fin erosion. Secondary factors, like water quality, opportunistic infections, and diet, can inhibit healing and regeneration. Stress is frequently believed to be a contributing factor as cortisol is a notorious inhibitor of regeneration and repair (Iger, Balm, Jenner, & Bonga, 1995; Roubal & Bullock, 1988). Genetics may also play a role in the propensity for aggressive behavior and immune response. Damage to epidermal club cells results in the release of alarm substance, a pheromone that induces anxiogenic behavior and increased cortisol in conspecific zebrafish (Abreu, Giacomini, Koakoski, Piato, & Barcellos, 2017; Egan et al., 2009; Pfeiffer, 1977; Waldman, 1982). The compound effects of injury, stress,

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and altered behavior should, therefore, be considered when evaluating the prevalence of fin lesions and the roles of primary and secondary factors. Zebrafish fins have a robust regenerative capacity (Azevedo, Grotek, Jacinto, Weidinger, & Saude, 2011). Complete regeneration of the caudal fin takes approximately 2 weeks under optimal circumstances. However, if a fin ray is removed to the base, it may not regrow (Goss and Stagg., 1957). The ZIRC noted a high prevalence of fin lesions, including splitting, fraying, missing bony rays, and severe truncation in the year following a fire in the building (Fig. 39.9). Postfire recovery and reconstruction involved significant and random variations in sound, vibration, and light intensity in the fish room. Tanks were covered in opaque plastic sheeting during ceiling work, which affected tank illumination, feeding schedules, and regular observation of the fish. Six to 8 months after the fire, fin lesions developed in several lines. The most severe lesions occurred in the pelvic fins, which were almost completely truncated in some cases. Although pelvic fin lesions were the most dramatic, they were also the most difficult to visualize and required anesthesia for an accurate assessment. Lesions were also routinely observed in dorsal, anal, and caudal fins. Water and fire-related fine black debris were negative for common toxins. Histological sections of fish with lesions were evaluated, and no infectious agents or lesions besides those on the fins were identified. There was no change in diet source, expiration date, quantity, or content. Lesions occurred in several different wild-type, mutant, and transgenic lines, ruling out a simple genetic cause. Color mutants were the most severely affected stocks. Interestingly, albino zebrafish have previously been described as high-anxiety compared to other zebrafish lines (Egan et al., 2009). Aggressive nipping was observed in some tanks, but aggression did not seem consistently elevated in all tanks. In 20-gallon (75.7-L) tanks, strands of green netting were suspended from floating tubes to give fish a hiding place or distraction. Healing was observed in lesions that did not involve complete removal of a bony ray, and the lesions eventually resolved as operations and room parameters normalized. We concluded that the stress associated with fire recovery and reconstruction played a major role in lesion development. A second incidence of fin lesions occurred 2 years later. The elevated noise and vibration that accompanied installation of a cage washer adjacent to the fish room may have played a role. Diagnosis. Diagnosis is made by gross visualization of fin lesions in multiple fish in a tank or population. Fins may be split, frayed, reduced in length, or nodular. Histologically, necrosis and epithelial hyperplasia may also be observed. Both dermis and epidermis are affected. Histological sections should be evaluated to

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39. Water Quality and Idiopathic Diseases of Laboratory Zebrafish

(A)

(B)

(C)

(D)

FIGURE 39.9

Fin lesions observed at the Zebrafish International Resource Center. (A) Missing fin rays, (B) split fin, (C) truncated fin, and

(D) healing fin.

rule out a primary infectious cause for the lesions. Bacterial and fungal cultures may be useful, but should be interpreted cautiously and in addition to histology as the culture of external surfaces are easily contaminated by normal environmental flora and opportunistic secondary invaders. Control and Treatment. The etiology of fin lesions at a population level can be multifactorial. Therefore, control efforts should include ruling out infectious, genetic, diet, toxic, and water quality-related causes and minimizing environmental and mechanical stress. Methods to reduce aggression should be considered. Feeding protocols should be evaluated to determine if fish are fed to satiety, if all fish have access to food and whether the location and/or method of food delivery potentiates competition and/or stress. The role of tank dynamics, stocking densities, and environmental enrichment should also be assessed. Spawning tank aggression may be mitigated by limiting the amount of time males and females are mixed, providing hiding places or distracting material (e.g., fake grass), and assuring optimal water quality. Facility alterations in light, sound,

and vibration should also be considered and mitigated to the greatest possible extent.

References Abbas, H. H., Zaghloul K. H., & Mousa, M. A. (2002). Effect of some heavy metal pollutants on some biochemical and histopathological changes in blue tilapia, Oreochromis aureus. Egypt Journal Agricultural Research, 80, 1395e1411. Abreu, M. S., Giacomini, A. C. V. V., Koakoski, G., Piato, A. L. S., & Barcellos, L. J. G. (2017). Divergent effect of fluoxetine on the response to physical or chemical stressors in zebrafish. PeerJ, 5, e3330. https://doi.org/10.7717/peerj.3330. Alsop, D., & Wood, C. M. (2011). Metal uptake and acute toxicity in zebrafish: Common mechanisms across multiple metals. Aquatic Toxicology, 105(3e4), 385e393. https://doi.org/10.1016/ j.aquatox.2011.07.010. Andersen, R. J., Luu, H. A., Chen, D. Z., Holmes, C. F., Kent, M. L., Le Blanc, M., et al. (1993). Chemical and biological evidence links microcystins to salmon ’netpen liver disease’. Toxicon, 31(10), 1315e1323. Arillo, A., Gaino, E., Margiocco, C., Mensi, P., & Schenone, G. (1984). Biochemical and ultrastructural effects of nitrite in rainbow trout: Liver hypoxia as the root of the acute toxicity mechanism. Environmental Research, 34(1), 135e154.

IV. Diseases

References

Authman, M. M. N., Zaki, M. S., Khallaf, E. A., & Abbas, H. H. (2015). Use of fish as bio-indicator of the effects of heavy metals pollution. Journal Aquatic Research Development, 6(4), 1e13. Azevedo, A. S., Grotek, B., Jacinto, A., Weidinger, G., & Saude, L. (2011). The regenerative capacity of the zebrafish caudal fin is not affected by repeated amputations. PLoS One, 6(7). https:// doi.org/10.1371/journal.pone.0022820. ARTN e22820. Baker, J. T. P. (1969). Histological and electron microscopical observations on copper poisoning in winter flounder (Pseudopleuronectes americanus). Journal of the Fisheries Research Board of Canada, 26(11), 2785e&. Bath, R. N., & Eddy, F. B. (1980). Transport of nitrite across fish gills. Journal of Experimental Zoology, 214(1), 119e121. https://doi.org/ 10.1002/jez.1402140115. Batzios, C., Fotis, G., & Gavriilidou, I. (1998). Economic dimension of gas bubble disease effects on rainbow trout culture. Aquaculture International, 6(6), 451e455. https://doi.org/10.1023/A: 1009235518343. Beraldo, P., & Canavese, B. (2010). Recovery of opercular anomalies in gilthead sea bream, Sparus aurata L.: Morphological and morphometric analysis. Journal of Fish Diseases, 34(1), 21e30. Beraldo, P., Pinosa, M., Tibaldi, E., & Canavese, B. (2003). Abnormalities of the operculum in gilthead sea bream (Sparus aurata): Morphological description. Aquaculture, 220(1e4), 89e99. https:// doi.org/10.1016/S0044-8486(02)00416-7. Bouck, G. R. (1980). Etiology of gas bubble disease. Transactions of the American Fisheries Society, 109(6), 703e707. https://doi.org/ 10.1577/1548-8659(1980)109<703:Eogbd>2.0.Co;2. Bowser, P. R., Falls, W. W., Vanzandt, J., Collier, N., & Phillips, J. D. (1983). Methemoglobinemia in channel catfish: methods of prevention. Progressive Fish-Culturist, 45(3), 154e158. https:// doi.org/10.1577/1548-8659(1983)45[154:Micc]2.0.Co;2. Braun, M. H., Steele, S. L., & Perry, S. F. (2009). The responses of zebrafish (Danio rerio) to high external ammonia and urea transporter inhibition: Nitrogen excretion and expression of rhesus glycoproteins and urea transporter proteins. Journal of Experimental Biology, 212(23), 3846e3856. https://doi.org/10.1242/ jeb.034157. Cahu, C., Infante, J. Z., & Takeuchi, T. (2003). Nutritional components affecting skeletal development in fish larvae. Aquaculture, 227(1e4), 245e258. https://doi.org/10.1016/S0044-8486(03)00507-6. Chen, C. Y., Wooster, G. A., Getchell, R. G., Bowser, P. R., & Timmons, M. B. (2001). Nephrocalcinosis in nile tilapia from a recirculation aquaculture system: A case report. Journal of Aquatic Animal Health, 13(4), 368e372. https://doi.org/10.1577/15488667(2001)013<0368:Nintfa>2.0.Co;2. Chowdhury, M. J., Girgis, M., & Wood, C. M. (2016). Revisiting the mechanisms of copper toxicity to rainbow trout: Time course, influence of calcium, unidirectional Naþ fluxes, and branchial Naþ, Kþ ATPase and V-type Hþ ATPase activities. Aquatic Toxicology, 177, 51e62. https://doi.org/10.1016/j.aquatox.2016.05.009. Cockell, K. A., Hilton, J. W., & Bettger, W. J. (1991). Chronic toxicity of dietary disodium arsenate heptahydrate to juvenile rainbow trout (Oncorhynchus mykiss). Archives of Environmental Contamination and Toxicology, 21(4), 518e527. Colt, J., & Armstrong, D. (1979). Nitrogen toxicity to fish, crustaceans, and molluscs. Davis, CA. Cowey, C. B., Knox, D., Adron, J. W., George, S., & Pirie, B. (1977). Production of renal calcinosis by magnesium deficiency in rainbow trout (Salmo gairdneri). British Journal of Nutrition, 38(1), 127e&. https://doi.org/10.1079/Bjn19770068. Dabrowska, H., Meyerburgdorff, K. H., & Gunther, K. D. (1991). Magnesium status in freshwater fish, common carp (Cyprinus carpio, L) and the dietary protein-magnesium interaction. Fish Physiology and Biochemistry, 9(2), 165e172. https://doi.org/10.1007/ Bf02265132.

475

Daoust, P. Y., & Ferguson, H. W. (1984). The pathology of chronic ammonia toxicity in rainbow trout, Salmo gairdneri Richardson. Journal of Fish Diseases, 7(3), 199e205. https://doi.org/10.1111/ j.1365-2761.1984.tb00924.x. Dave, G., & Xiu, R. (1991). Toxicity of mercury, copper, nickel, lead, and cobalt to embryos and larvae of zebrafish, Brachydanio rerio. Archives of Environmental Contamination and Toxicology, 21, 126e134. Egan, R. J., Bergner, C. L., Hart, P. C., Cachat, J. M., Canavello, P. R., Elegante, M. F., et al. (2009). Understanding behavioral and physiological phenotypes of stress and anxiety in zebrafish. Behavioural Brain Research, 205(1), 38e44. https://doi.org/10.1016/ j.bbr.2009.06.022. Ellis, T., Oidtmann, B., St-Hilaire, S., Turnbull, J. F., North, B. P., MacIntyre, C. M., et al. (2008). Fin erosion in farmed fish. In E. Branson (Ed.), Fish welfare (pp. 121e149). Blackwell Publishing Ltd. Farrell, A. P., Ackerman, P. A., & Iwama, G. K. (2010). Disorders of the cardiovascular and respiratory systems. In J. F. W. Leatherland (Ed.), Fish diseases and disorders (2nd ed., Vol. 2, pp. 287e322). Preston, UK: CAB International. Non-infectious Disorders. Gaino, E., Arillo, A., & Mensi, P. (1984). Involvement of the gill chloride cells of trout under acute nitrite intoxication. Comparative Biochemistry and Physiology a-Physiology, 77(4), 611e617. https://doi.org/ 10.1016/0300-9629(84)90171-3. Gardner, G. R., & Laroche, G. (1973). Copper induced lesions in estuarine teleosts. Journal of the Fisheries Research Board of Canada, 30(3), 363e&. Goss, R. J., & Stagg, M. W. (1957). The regeneration of fins and fin rays in Fundulus heteroclitus. Journal of Experimental Zoology, 136, 487e508. Harris, M. P., Henke, K., Hawkins, M. B., & Witten, P. E. (2014). Fish is fish: The use of experimental model species to reveal causes of skeletal diversity in evolution and disease. Journal of Applied Ichthyology, 30(4), 616e629. https://doi.org/10.1111/jai.12533. Hayes, A. J., Reynolds, S., Nowell, M. A., Meakin, L. B., Habicher, J., Ledin, J., et al. (2013). Spinal deformity in aged zebrafish is accompanied by degenerative changes to their vertebrae that resemble osteoarthritis. PLoS One, 8(9). https://doi.org/10.1371/journal.pone.0075787. ARTN e75787. Hendricks, J. D., Sinnhuber, R. O., Henderson, M. C., & Buhler, D. R. (1981). Liver and kidney pathology in rainbow trout (Salmo gairdneri) exposed to dietary pyrrolizidine (Senecio) alkaloids. Experimental and Molecular Pathology, 35(2), 170e183. https:// doi.org/10.1016/0014-4800(81)90057-5. Hilomen-Garcia, G. V. (1997). Morphological abnormalities in hatchery-bred milkfish (Chanos chanos Forsskal) fry and juveniles. Aquaculture, 152(1e4), 155e166. https://doi.org/10.1016/S00448486(96)01518-9. Hilton, J. W., Hodson, P. V., & Slinger, S. J. (1980). The requirement and toxicity of selenium in rainbow trout (Salmo gairdneri). Journal of Nutrition, 110(12), 2527e2535. Hogstrand, C., Verbost, P. M., Bonga, S. E. W., & Wood, C. M. (1996). Mechanisms of zinc uptake in gills of freshwater rainbow trout: Interplay with calcium transport. American Journal of Physiologye Regulatory, Integrative and Comparative Physiology, 270(5), R1141eR1147. Huey, D. W., Simco, B. A., & Criswell, D. W. (1980). Nitrite-induced methemoglobin formation in channel catfish. Transactions of the American Fisheries Society, 109(5), 558e562. https://doi.org/ 10.1577/1548-8659(1980)109<558:Nmficc>2.0.Co;2. Iger, Y., Balm, P. H. M., Jenner, H. A., & Bonga, S. E. W. (1995). Cortisol induces stress-related changes in the skin of rainbow-trout (Oncorhynchus mykiss). General and Comparative Endocrinology, 97(2), 188e198. https://doi.org/10.1006/gcen.1995.1018.

IV. Diseases

476

39. Water Quality and Idiopathic Diseases of Laboratory Zebrafish

Jensen, G. (1989). Handbook for common calculations in finfish aquaculture. Baton Rouge, Louisiana: Louisiana State University, Agricultural Center. Vol. Publication 8903. Kent, M. L., Buchner, C., Watral, V. G., Sanders, J. L., Ladu, J., Peterson, T. S., et al. (2011). Development and maintenance of a specific pathogen-free (SPF) zebrafish research facility for Pseudoloma neurophilia. Diseases of Aquatic Organisms, 95(1), 73e79. https:// doi.org/10.3354/dao02333. Kent, M. L., Myers, M. S., Hinton, D. E., Eaton, W. D., & Elston, R. A. (1988). Suspected toxicopathic hepatic-necrosis and megalocytosis in pen-reared atlantic salmon Salmo salar in Puget Sound, Washington, USA. Diseases of Aquatic Organisms, 4(2), 91e100. https:// doi.org/10.3354/dao004091. Kent, M. L., Spitsbergen, J. M., Matthews, J. L., Fournie, J. W., Murray, K. N., & Westerfield, M. (2016). Diseases of zebrafish in research facilities. https://zebrafish.org/wiki/health/disease_ manual/start. Koumoundouros, G., Oran, G., Divanach, P., Stefanakis, S., & Kentouri, M. (1997). The opercular complex deformity in intensive gilthead sea bream (Sparus aurata L.) larviculture. Moment of apparition and description. Aquaculture, 156(1e2), 165e177. https:// doi.org/10.1016/S0044-8486(97)89294-0. Krise, W. F., & Meade, J. W. (1988). Effects of low-level gas supersaturation on lake trout (Salvelinus namaycush). Canadian Journal of Fisheries and Aquatic Sciences, 45(4), 666e674. Kroupova, H., Machova, J., & Svobodova, Z. (2005). Nitrite influence on fish: A review. Veterinarni Medicina, 50(11), 461e471. Lall, S. P. (2010). Disorders of nutrition and metabolism. Non-infectious Disorders. In J. F. W. Leatherland (Ed.), Fish diseases and disorders (2nd ed., Vol. 2, pp. 202e237). Oxfordshire, UK: CAB International. Vol. 2. Lall, S. P., & Lewis-McCrea, L. M. (2007). Role of nutrients in skeletal metabolism and pathology in fishean overview. Aquaculture, 267(1e4), 3e19. https://doi.org/10.1016/j.aquaculture.2007.02.053. Latremouille, D. N. (2003). Fin erosion in aquaculture and natural environments. Reviews in Fisheries Science, 11(4), 315e335. https:// doi.org/10.1080/10641260390255745. Lewis, W. M., & Morris, D. P. (1986). Toxicity of nitrite to fish: a review, Transactions of the American Fisheries Society, 115(2), 183e195. https://doi.org/10.1577/1548-8659(1986)115<183:Tontf>2.0.Co;2. Lindesjoo, E., Thulin, J., Bengtsson, B. E., & Tjarnlund, U. (1994). Abnormalities of a gill cover bone, the operculum, in perch Perca fluviatilis from a pulp mill effluent area. Aquatic Toxicology, 28(3e4), 189e207. https://doi.org/10.1016/0166-445x(94)90033-7. Lund, M., & Heggberget, T. G. (1985). Avoidance-response of 2-yearold rainbow-trout, Salmo-gairdneri Richardson, to air-supersaturated water: hydrostatic compensation. Journal of Fish Biology, 26(2), 193e200. https://doi.org/10.1111/j.1095-8649.1985.tb0 4256.x. Machado, J. P., Garling, D. L., Kevern, N. R., Trapp, A. L., & Bell, T. G. (1987). Histopathology and the pathogenesis of embolism (gas bubble disease) in rainbow trout (Salmo gairdneri). Canadian Journal of Fisheries and Aquatic Sciences, 44(11), 1985e1994. https://doi.org/ 10.1139/f87-243. Meade, J. W. (1985). Allowable ammonia for fish culture. Progressive Fish-Culturist, 47(3), 135e145. https://doi.org/10.1577/15488640(1985)47<135:Aaffc>2.0.Co;2. Myers, M. S., Johnson, L. L., & Collier, T. K. (2003). Establishing the causal relationship between polycyclic aromatic hydrocarbon (PAH) exposure and hepatic neoplasms and neoplasia-related liver lesions in English sole (Pleuronectes vetulus). Human and Ecological Risk Assessment, 9(1), 67e94. https://doi.org/10.1080/713609853. Niyogi, S., & Wood, C. M. (2004). Kinetic analyses of waterborne Ca and Cd transport and their interactions in the gills of rainbow trout (Oncorhynchus mykiss) and yellow perch (Perca flavescens), two species differing greatly in acute waterborne Cd sensitivity. Journal of

Comparative Physiology B: Biochemical, Systemic, and Environmental Physiology, 174(3), 243e253. https://doi.org/10.1007/s00360-0030409-x. Nu¨sslein-Volhard, C., & Dahm, R. (2002). Zebrafish: A practical approach (1st ed.). Oxford: Oxford University Press. Perschbacher, P. W., & Wurts, W. A. (1999). Effects of calcium and magnesium hardness on acute copper toxicity to juvenile channel catfish, Ictalurus punctatus. Aquaculture, 172(3e4), 275e280. https:// doi.org/10.1016/S0044-8486(98)00499-2. Peuranen, S., Vuorinen, P. J., Vuorinen, M., & Hollender, A. (1994). The effects of iron, humic acids and low pH on the gills and phsiology of brown trout (Salmo Trutta). Annales Zoologici Fennici, 31, 389e396. Pfeiffer, W. (1977). Distribution of fright reaction and alarm substance cells in fishes. Copeia, (4), 653e665. https://doi.org/10.2307/ 1443164. Pohl, C. (1990). Skeletal deformities and trace-metal contents of European smelt, Osmerus-eperlanus, in the elbe estuary. Meeresforschung-Reports on Marine Research, 33(1), 76e89. Richetti, S. K., Rosemberg, D. B., Ventura-Lima, J., Monserrat, J. M., Bogo, M. R., & Bonan, C. D. (2011). Acetylcholinesterase activity and antioxidant capacity of zebrafish brain is altered by heavy metal exposure. Neurotoxicology, 32(1), 116e122. https://doi.org/ 10.1016/j.neuro.2010.11.001. Rosenberg, G. A. (2012). Molecular physiology and metabolism of the nervous system : A clinical perspective. New York: Oxford University Press. Rossteuscher, S., Schmidt-Posthaus, H., Schafers, C., Teigeler, M., & Segner, H. (2008). Background pathology of the ovary in a laboratory population of zebrafish Danio rerio. Diseases of Aquatic Organisms, 79(2), 169e172. https://doi.org/10.3354/dao01893. Roubal, F. R., & Bullock, A. M. (1988). The mechanism of wound repair in the skin of juvenile atlantic salmon, Salmo salar L., following hydrocortisone implantation. Journal of Fish Biology, 32(4), 545e555. https://doi.org/10.1111/j.1095-8649.1988.tb05394.x. Schwedler, T. E., Tucker, C. S., & Beleau, M. H. (1985). Non-infectious diseases. In C. S. Tucker (Ed.), channel catfish culture (pp. 497e541). Amsterdam: Elsevier. Sfakianakis, D. G., Georgakopoulou, E., Papadakis, I. E., Divanach, P., Kentouri, M., & Koumoundouros, G. (2006). Environmental determinants of haemal lordosis in European sea bass, Dicentrarchus labrax (Linnaeus, 1758). Aquaculture, 254, 54e64. Smart, G. R. (1978). Investigations of the toxic mechanisms of ammonia to fishegas exchange in rainbow trout (Salmo gairdneri) exposed to acutely lethal concentrations. Journal of Fish Biology, 12, 93e104. Smith, C. E. (1988). Histopathology of gas bubble disease in juvenile rainbow-trout. Progressive Fish-Culturist, 50(2), 98e103. https:// doi.org/10.1577/1548-8640(1988)050<0098:Chogbd>2.3.Co;2. Sorensen, E. M. B. (1991). Metal poisoning in fish. Boca Raton, Fl: CRC Press. Speare, D. J. (1991). Endothelial lesions associated with gas bubble disease in fish. Journal of Comparative Pathology, 104(3), 327e335. https://doi.org/10.1016/S0021-9975(08)80044-8. Spitsbergen, J. M., Buhler, D. R., & Peterson, T. S. (2012). Neoplasia and neoplasm-associated lesions in laboratory colonies of zebrafish emphasizing key influences of diet and aquaculture system design. ILAR Journal, 53(2), 114e125. https://doi.org/10.1093/ ilar.53.2.114. Spitsbergen, J. M., & Kent, M. L. (2003). The state of the art of the zebrafish model for toxicology and toxicologic pathology researchd advantages and current limitations. Toxicologic Pathology, 31(Suppl. l), 62e87. https://doi.org/10.1080/01926230390174959. Stephen, C., Kent, M. L., & Dawe, S. C. (1993). Hepatic megalocytosis in wild and farmed chinook salmon Oncorhynchus-tshawytscha in British-Columbia, Canada. Diseases of Aquatic Organisms, 16(1), 35e39. https://doi.org/10.3354/dao016035.

IV. Diseases

References

Stickland, N. C., White, R. N., Mescall, P. E., Crook, A. R., & Thorpe, J. E. (1988). The effect of temperature on myogenesis in embryonic-development of the atlantic salmon (Salmo salar L.). Anatomy and Embryology, 178(3), 253e257. https://doi.org/ 10.1007/Bf00318228. Strauss, R. H. (1979). Diving medicine. American Review of Respiratory Disease, 119(6), 1001e1023. Turnbull, J. F., Richards, R. H., & Robertson, D. A. (1996). Gross, histological and scanning electron microscopic appearance of dorsal fin rot in farmed Atlantic salmon, Salmo salar L., parr. Journal of Fish Diseases, 19(6), 415e427. https://doi.org/10.1111/j.13652761.1996.tb00381.x. Tye, M. T., Montgomery, J. E., Hobbs, M. R., Vanpelt, K. T., & Masino, M. A. (2018). An adult zebrafish diet contaminated with chromium reduces the viability of progeny. Zebrafish, 15(2), 179e187. Voslarova, E., Pistekova, V., & Svobodova, Z. (2006). Nitrite toxicity to Danio rerio: Effects of fish age and chloride concentrations. Acta Veterinaria Brno, 75(1), 107e113. https://doi.org/10.2754/ avb200675010107.

477

Voslarova, E., Pistekova, V., Svobodova, Z., & Bedanova, I. (2008). Nitrite toxicity to Danio rerio: Effects of subchronic exposure of fish growth. Acta Veterinaria Brno, 77, 455e460. Waldman, B. (1982). Quantitative and developmental analyses of the alarm reaction in the Zebra danio, Brachydanio-rerio. Copeia, (1), 1e9. Walters, G. R., & Plumb, J. A. (1980). Environmental-stress and bacterial-infection in channel catfish, ictalurus punctatus Rafinesque. Journal of Fish Biology, 17(2), 177e185. https:// doi.org/10.1111/j.1095-8649.1980.tb02751.x. Wedemeyer, G. A. (1996). Physiology of fish in intensive culture systems. New York: Chapman & Hall. Wedemeyer, G. A., & Yasutake, W. T. (1978). Prevention and treatment of nitrite toxicity in juvenile steelhead trout (Salmo-gairdneri). Journal of the Fisheries Research Board of Canada, 35(6), 822e827. Weitkamp, D. E., & Katz, M. (1980). A review of dissolved-gas supersaturation literature. Transactions of the American Fisheries Society, 109(6), 659e702. https://doi.org/10.1577/1548-8659(1980) 109<659:Arodgs>2.0.Co;2.

IV. Diseases