Salmonid Whirling Disease by Maria E Markiw U.S. Fish and Wildlife Service National Fisheries Research Center-Leetown National Fish Health Research Laboratory Box 700 Kearneysville, West Virginia 25430

Current Address: U.S. Geologic Survey, Biological Resources Division Leetown Science Center National Fish Health Research Laboratory 1700 Leetown Road Kearneysville, West Virginia 25430 Introduction Whirling disease is a parasitic infection of trout and salmon by the myxosporean protozoan Myxobolus cerebralis (Syn. Myxosoma cerebralis). This parasite has selective tropism for cartilage; infection can cause deformities of the axial skeleton and neural damage that results in "blacktail." The disease is named for the erratic, tail-chasing, "whirling' in young fish that are startled or fed. Heavy infection of young fish can result in high mortalities or unmarketable, deformed individuals. Although the parasite was first reported in 1903 in central Europe (Hofer 1903), its complete life cycle was not described until the early 1980's.   History and Geographic Range Whirling disease occurs in much of Europe (Halliday 1976) where it probably originated. It occurs in the former Soviet Union (Uspenskaya 1955) and was seemingly introduced into the British Isles where it is now common (Elson 1969; O'Brien 1976; Hudson and Holliman 1985). It was accidentally introduced into New Zealand (Hewitt and Little 1972) and into the United States. The detailed history of the disease and its introduction into the United States (into Pennsylvania and Nevada in about 1955) were discussed in a recent review by Hoffman (1990). Although Hoffman provided an extensive list of the present worldwide distribution of the infection, the cited occurrence in several countries is subject to dispute because of the applied methods of spore detection and identification. Myxobolus cerebralis was probably established much earlier than reported because it may require several years for the parasite to become established at sufficiently high intensity for clinical signs to appear in fish. In the United States, whirling disease has been detected in 22 states: Alabama, California, Colorado, Connecticut, Idaho, Maryland, Massachusetts, Michigan, Montana, Nevada, New Hampshire, New Jersey, New York, Ohio, Oregon, Pennsylvania, Utah, Virginia, Washington, West Virginia, and Wyoming.   Diagnosis Clinical Signs Moderate or heavy clinical infection of fish with whirling disease can be presumptively diagnosed on the basis of changes in behavior and appearance. When alarmed or feeding, some infected individuals show an abnormal whirling behavior. The caudal peduncle and tail may become dark or even black, but these characteristics fade in preserved specimens (Fig. 1). The whirling behavior is believed to be the result of impaired coordination caused by neural damage from lesions and disintegration of cartilaginous tissue around the organs of equilibrium. These clinical signs appear, depending on temperature and intensity of the infection, about 35 to 80 days after initial infection and can persist for about a year. Deformities of the axial skeleton or head, shortening of the snout, and cranial depressions persist through the life of the infected fish (Fig. 2). Individually, these signs are not conclusive. Injury or deficiency in dietary tryptophan and ascorbic acid can evoke similar signs (Wolf et at. 1981). For example, contamination with toxaphene may cause spinal defects in fish, described as "broken back," and crippled fish also cannot swim properly. However, the collective appearance of all signs throughout a population indicate clinical infection with whirling disease. Fig. 1. Black tail in 4-month-old rainbow trout (Oncorhynchus mykiss).  Fig. 2. Deformed skeletons of 8-month old rainbow trout (Oncorhynchus mykiss) with whirling disease. In gross pathological examination, internal organs appear normal. Histological sections of cartilage, particularly skull, gill, and vertebrae, show areas of lysis and inflammation. If the infection has existed for 3-4 months, depending on temperature, spores of the myxozoan M. cerebralis had time to form in or around the cartilage lesions (Fig. 3). The presence of M. cerebralis spores in cartilage areas is considered pathognomonic for whirling disease. Fig. 3. Myxobolus cerebralis spores, 8-10 mm in diameter, in histological section of cartilage tissue of rainbow trout(Oncorhynchus mykiss); stained with methylene blue. x 1,000 magnification. To date, no reliable nondestructive serological procedures have been developed for detecting the causal organism of whirling disease in fish. Nonspecific, false positive, and false negative reactions have been found in tested fish (Griffin and Davis 1978; Markiw, unpublished data). The long life cycle of the parasite, about 3 months in fish and 3.5 months in tubificid worms, may result in continual changes of antigenic components. Although hematoxylin and eosin stains are routinely used in histology, they do not enhance the appearance of spores of M. cerebralis. Methylene blue, Giemsa or May-Grünwald Giemsa, or Ziehl-Neelsen stains are recommended because the polar capsules react strongly and make the spores prominent.   Identification Myxobolus cerebralis is the only myxosporean found in the cartilage of salmonids. The mature spore is lenticular in side view and nearly circular in front view (Fig. 3). The spores are 8­10 mm in greatest diameter and have two prominent ovate polar capsules with coiled filaments that may be extruded in certain situations. Aberrant spores (Fig. 4), either in shape or with unequal polar capsules, may also be found (Lom and Hoffman 1971; Markiw and Wolf 1974a). The iodinophilous vacuole is not present. However, this test is not always reliable taxonomically and can be performed on only fresh spores. Sole identification by morphology may be difficult because M. cerebralis-infected fish may have mixed infections with other myxosporeans from the central nervous system, muscle, or skin. Attempts at morphologic identification by inexperienced persons may result in uncertainty; therefore, referral to a knowledgeable parasitologist is recommended. Fig. 4. Myxobolus cerebralis spores and prespore stages isolated and concentrated from infected cartilage by the enzymatic digestion method. Listed are the spores of other Myxobolus species, similar to M. cerebralis, that can occasionally be found in the head, but not in the cartilage or bone, of salmonids. Preserved spores are usually about 10% smaller than fresh spores. Myxobolus kisutchi--in the central nervous system of coho salmon (Oncorhynchus kisutch) and chinook salmon (Oncorhynchus tshawytscha). The preserved spores (formalin) are 7­8 mm in diameter, appear uniform in shape, and contain an iodinophilous vacuole. Myxobolus squamalis--in the scales of rainbow trout (Oncorhynchus mykiss) and salmon from the western United States. The preserved spores (formalin) are 8­9 mm in diameter and appear uniform, with equal polar capsules and with a narrow ridge that parallels either side of the sutural ridge. Myxobolus arcticus--in the central nervous system of coho salmon, sockeye salmon (Oncorhynchus nerka), Dolly Varden char (Salvelinus malma), lake char "Neyva" (Salvelinus neiva), Arctic grayling (Thymallus arcticus), Arctic char (Salvelinus alpinus), and whitefish (Coregonus clupeaformis), the fresh spores are large, 14.3­16.5 x 7.6­7.7 mm, with large, elongated polar capsules (recent description by Pugachev and Khokhlov 1979). Myxobolus neurobius--in the central nervous system of brown trout (Salmo trutta); Arctic grayling; European grayling (Thymallus) from central Europe, Eurasia, and North America; and arctic char and wild young Atlantic salmon (Salmo salar) in Newfoundland (Maloney et al. 1991). The preserved spores (glycerin) are oval and appear in a wide range of sizes, 10­12 x 8 mm (Schuberg and Schroeder 1905); but fresh spores are larger, 13.4­14 x 8.5­9.2 mm, according to a recent description by Pugachev and Khokhlov (1979). Myxobolus insidiosus--in the muscle of cutthroat trout (Oncorhynchus clarki), chinook salmon, and coho salmon from the western United States. The fresh spores are about the same size and shape as M. arcticus, 12.8­17.3 x 9­11.5 mm. Histological location and identification of M. cerebralis spores in lesions of skeletal tissue, particularly of the head, have been recommended for confirmation of diagnosis. However, this approach is not reliable with lightly infected fish that have only a few spores. Hamilton and Canning (1988) used Historesin-embedded sections of infected rainbow trout for detection of M. cerebralis spores and prespore stages by an indirect fluorescent antibody test. They used Percoll-purified spores as antigen for production of mouse anti-M. cerebralis serum. The antiserum reacted with early stages of the parasite, but the fluorescence pattern was more clearly seen in younger (5­month-old) fish, which had been experimentally infected, than in older fish in samples from fish farms. A presumptive diagnosis is based on location, size, and morphology of the spores and epizootiological data, such as geographical location and history of the hatchery, the species of fish, and the clinical signs. Diagnosis is usually confirmed by the identity of the spores, which is determined from a direct fluorescent antibody test with rabbit antiserum against M. cerebralis or Triactinomyxon spores conjugated with fluorescein isothiocyanate (Markiw and Wolf 1978; Markiw 1989a). The response of M. cerebralis spores can be seen with fluorescence microscopy (Fig. 5). Antiserum, prepared at the National Fish Health Research Laboratory (West Virginia), showed cross reactivity (++) only with Myxosoma cartilaginis of bluegills (Lepomis macrochirus). Fig. 5. Serological identification of Myxobolus cerebralis spores by the direct fluorescent antibody test. Note typical distribution of reactive antigen by M. cerebralis spores. The fluorescent antibody test works best with fresh spores or with spores fixed in methanol. The specific fluorescence of older specimens of spores that have been stored in formalin for a week or more is reduced and that of older specimens is insignificant or nonexistent.   Detection of Myxobolus cerebralis Detection of spores in moderate or heavy M. cerebralis infections at 5­6 months is relatively easy because about 25,000 to 2 million spores are in the head cartilage. Quantitatively, about two thirds of all spores are in the head; more than half of those are in the cartilage of the gill arches. When signs of the disease are evident, the simplest procedure for detecting spores is to split the head of a suspect fish sagittally, scrape the areas of the organs of equilibrium, and examine the scraping microscopically at X20 or X40 magnification. When examining asymptomatic suspect fish, the simplest and most rapid first step is to remove and grind the gill arches and suspend the homogenate in several volumes of water. After the particulate matter has been allowed to settle for 2­3 min, several drops of a supernatant are examined microscopically. If no spores are found after a search of 5­10 min, gill arches from another fish should be homogenized and examined. If no spores are found during these simple procedures, one of two methods of spore concentration should be used: the modified plankton centrifuge method of O'Grodnick (1975a) or the pepsin-trypsin­dextrose (PTD) digestion method of Markiw and Wolf (1974a). These methods are the most sensitive yet developed. The modified plankton centrifuge method includes trypsinization of the harvest for clarity to increase sensitivity (Markiw and Wolf 1980). The procedure can be completed in 2­3 h and, therefore, has been more widely used in laboratories where large numbers of M. cerebralis examinations are made. The method works well on young fresh or frozen heads and formalin­fixed material. The PTD digestion method is more sensitive and was developed for detection of spores in very lightly infected carriers (about 100 spores per head) and in 4- to 5­year old fish (Markiw and Wolf 1974b). Fresh or frozen materials are suitable and the procedure takes 6­8 h. The method does not work with materials preserved in formalin or other fixatives. Detection of spores in large fish requires much labor and skillful handling. Lorz et al. (1989) developed a labor saving technique for examinations of large fish. They reduced examined head tissue by using core samples from the area of the otoliths, 110 mm long and 19 mm in diameter, taken from the head with a cork borer. The borer was inserted into the head dorsally and perpendicular to the long axis of the body, about 10 mm behind the eye, and was pushed through the roof of the mouth. Then samples were processed by the enzymatic digestion method (PTD). The authors claimed to have detected more infected fish from core samples than from cranial elements from the entire head. This new technique may be useful for examination of large fish in epizootiological studies and for compliance with international laws. For the inspection of a hatchery for the presence of M. cerebralis, samples for examination should be weighted toward the most susceptible species and ages of available fish. If all were reared under the same conditions, rainbow or brook trout (Salvelinus fontinalis) should be examined before brown trout or coho salmon, and younger fish before older fish. Fish from earthen ponds should be examined before those from concrete raceways. If reared at 12° C or warmer water temperature, 2.5­ to 4­month-old fish yield mature spores. All equipment for diagnostic procedures should be decontaminated before each lot of fish is examined. Before disposal, infected fish tissues or liquids should be autoclaved or boiled in water for 20­30 min or incinerated. A 5-10 min disinfection in half-strength household bleach (Clorox, 5.25% sodium hypochlorite solution) or methanol is recommended to inactivate spores adhering to utensils.   Detection of Early Infection Under experimental conditions, the initial infection of whirling disease can be detected microscopically in wet mounts of the skin or fins or in histological sections (Fig. 6) in the form of aggregates of small (1.5­2 mm) intracellular sporozoites (sporoplasms). These can be detected only during a few hours after penetration of the infective Triactinomyxon spore stage because the sporozoites move or are transported rapidly from the external epithelial layers into deeper strata (Markiw 1989b). After initial infection of the fish, mature spores of M. cerebralis can be found in 2.6 months at a water temperature of 12.5° C. Fig. 6. Initial form of whirling disease infection in histological section of dorsal epithelium of rainbow trout fry 30 min after exposure to the Triactinomyxon spores. Intracellular inclusions of numerous small (1.5-2 mm in diameter) sporozoites (sporoplasms) are intensely stained with May-Grünwald Giemsa.   Life Cycle The whirling disease protozoan has a two­host life cycle (Fig. 7) involving a fish and the aquatic oligochaete Tubifex (Markiw and Wolf 1983; Wolf and Markiw 1984; Wolf et al. 1986); two separate stages of sporogony occur, one in each host. Antigenic homology of the two morphologically distinct spore forms was demonstrated serologically (Markiw 1989a). Fig. 7. Diagram of a two-host life cycle of the whirling disease parasite. In brief, spores of M. cerebralis are released into the aquatic environment when infected fish die and decompose or are consumed by predators or scavengers. The myxosporean-type spores are ingested by worms in whose gut epithelium the next phase develops (Fig. 8). Transformation into the actinosporean Triactinomyxon, the infective stage to fish, takes about 3.5 months at 12.5° C, after which infected worms release numerous mature forms into the water for several weeks. The Triactinomyxon spores are much larger and have three polar capsules and three grapple­like appendages, 170­180 mm long (Fig. 9). The Triactinomyxon stage enters susceptible fish through the epithelial cells of the skin, fins, buccal cavity (particularly at the base of the gills), upper esophagus, and lining of the digestive tract. Transformation into M. cerebralis spores then takes about 2.6 months at a water temperature of 12.5° C. This life cycle was confirmed by El­Matbouli and Hoffmann (1989) for M. cerebralis; a similar life cycle was shown for Myxobolus cotti. Fig. 8. Histological section of a tubificid worm infected with actinosporean 130 days after exposure to Myxobolus cerebralis spores. Note (arrows) several undeveloped cysts in a gut wall and mature Triactinomyxon in the lumen. May-Grünwald Giemsa stain. x 450 magnification. Fig. 9. Living unstained mature waterborne Triactinomyxon spore stage. The epispore contains three polar capsules, about 30-60 sporozoites, and three grapple-like appendages, 170-180 mm long. x 450 magnification. Although a two-host life cycle of the whirling disease organism is now widely accepted and the parasite has been recycled at this laboratory in fish or tubificids for nearly a decade without losing its infective potency, Hamilton and Canning (1987), Prihoda (1983), and Uspenskaya (1978) claimed direct transmission of the parasite from fish to fish by way of aged spores.   Transmission Salmonids contract whirling disease in two ways: by ingesting tubificids that harbor the specific actinosporean Triactinomyxon and by brief contact with waterborne Triactinomyxons released from infected tubificids. The experimentally produced actinosporean stage of M. cerebralis is short­lived, persisting 3­4 days at 12.5° C and fewer days at warmer temperatures (Markiw 1992b). Studies of the dynamics of the infective stage for fish (Markiw 1986) demonstrated that, after a single exposure to M. cerebralis spores, a population of infected tubificids can release viable Triactinomyxon spores for as long as a year at a level detectable by only sentinel fish. O'Grodnick (1975b) demonstrated that whirling disease cannot be transmitted vertically from infected brood stock to the egg. Shipments of salmonid eggs from waters contaminated with whirling disease are also unlikely to disseminate the parasite because rainbow trout are refractory to the infection during hatching and for a day afterward (Markiw 1991). Contrary to reports from eastern Europe and Russia (Prihoda 1983; Uspenskaya 1978), attempts to effect fish­to fish transmission of whirling disease or through aged spores of M. cerebralis in absence of tubificids in our laboratory have been unsuccessful.   Development Development time for both stages of the whirling disease organism, myxosporean in fish and actinosporean in tubificids, is directly related to temperature. Trout fry that are fed infected worms or exposed to waterborne Triactinomyxon show blacktail after 35­45 days at a water temperature of 12.5° C. Whirling behavior first appears at about the same time or slightly later. Fully mature spores of detected after 2.6­3.5 months at 12.5° C. Under M. cerebralis are experimental conditions, after a short single exposure (3 h) of 2­month-old rainbow trout to quantified numbers of Triactinomyxoninfected trout head cartilage ranged from less than 100 to , production of spores by M. cerebralis in nearly 2 million at 5 or 6 months and showed limitation of parasitism at the highest levels of infection (Markiw 1992a, 1992b). Development time is shortened or lengthened at temperatures above or below 12.5° C; about 50 days at 17° C and 120 days at 7° C (Halliday 1973). Development time in the worm is defined as the interval between first contact with M. cerebralis spores and the release of the first Triactinomyxon. Under experimental conditions at 12.5° C, after single exposure of one population of tubificid worms to M. cerebralis spores, the Triactinomyxons were released in a consistent pattern that began at 104­113 days, peaked during the next 15­60 days, and continued at trace levels for about 6 months. During the next 3 months the infectivity was detectable by only sentinel fish (Markiw 1986). Whether the same infected worms are releasing Triactinomyxons for 11 months or a new generation of worms must become infected with M. cerebralis spores to produce infectivity is not known. One tubificid worm, at peak of productivity (about 130 days after exposure) can harbor 900­1,000 mature Triactinomyxons.   Reservoir of Infectivity Trout and salmon can be infected with whirling disease and may harbor M. cerebralis spores. Predators and scavengers, such as birds (Taylor and Lott 1978) that consume infected fish, can release viable M. cerebralis spores into the environment and may disseminate the parasite. The source of the infective agent for fish is usually the water supply or earthen ponds inhabited by aquatic tubificid worms. An outbreak of the disease can occur after stocking with infected fish or transferring fish from facilities where the infection had not yet been detected.     Susceptibility and Host Range Young and adult trout and salmon are susceptible to M. cerebralis infection; but the severity of the infection decreases with age (Markiw 1992a). When fish are infected at an older age, they are usually asymptomatic, healthy­looking, and of normal size, but may carry the spores of M. cerebralis. Severe mortalities (about 90%) may occur among newly hatched fish that were exposed to the infective agent as sac fry; l-day­old rainbow trout are refractory to the infection (Markiw 1991). Not all salmonid species are equally susceptible to the infection. Whereas rainbow trout are most susceptible and brook trout less so, lake trout cannot be infected (O'Grodnick 1979). Other salmonids can be infected, but clinical signs of the disease may or may not develop. Susceptibility varies not only among species but also among strains and may vary tremendously among individual fish within a population (Markiw 1992a). In the following list, species are ranked in descending order of apparent susceptibility (O'Grodnick 1979; Hoffman 1990): rainbow trout, sockeye salmon, golden trout (Oncorhynchus aguabonita), cutthroat trout, brook trout, steelhead (Oncorhynchus mykiss), chinook salmon, Atlantic salmon, brown trout, coho salmon, lake trout (Salvelinus namaycush) and splake (hybrids between brook trout and lake trout). Lake trout and splake are refractory to infection with whirling disease. Testing susceptibility by standard exposure (Markiw, unpublished data) revealed that, as fry, greenback cutthroat trout are 7.5-fold less susceptible to the disease than rainbow trout. Rainbow trout exhibited all clinical signs of the disease, whereas greenback cutthroat trout were asymptomatic. Grayling (Thymallus sp.) and whitefish (Coregonus sp. and Prosopium sp.), which are generally regarded as salmonids, have not yet been tested and their susceptibility or resistance to whirling disease remains undetermined. According to early accounts (Halliday 1976), whirling disease was found in nonsalmonids. However, the author believes that these reports might be erroneous. Critical reexamination and identification of spores by serological methods are necessary. Tubifex is the only tubificid that has been identified as susceptible to M. cerebralis (Wolf et al. 1986). Members of the genera Limnodrilus, Quistadrilus, and Ilyodrilus in mixed populations with Tubifex did not produce Triactinomyxon when exposed to M. cerebralis spores. Other genera of oligochaetes that have been tested (Dero, Stylaria, and Aeolosoma) also did not produce infectivity for whirling disease (Markiw and Wolf 1983).   Prevention and Control At the present time, control of M. cerebralis infections is difficult. However, application of preventive measures can decrease the intensity of the disease in fish culture facilities and perhaps eliminate the spread to nonenzootic areas. Because tubificids are essential intermediate hosts for development of the infective stage in fish, the avoidance of earthen ponds for rearing fish should be considered. Tubificids are normal inhabitants of aquatic environments. They are particularly abundant in rich organic soils and occur in dense red patches (Fig. 10) in settling basins and streams that carry effluent from trout hatcheries. The life span of T. tubifex is about 2.5­3 years depending on environmental conditions (USSR Academy of Sciences 1972). Seasonal variation of oligochaete biomass is commonly observed with the largest biomass in fall and the smallest in spring. The phenomenon might correlate with the intensity of reproduction. The breeding and development of oligochaetes are directly associated with temperature (USSR Academy of Sciences 1972). Fig. 10. Tubificid oligochaetes, intermediate host for development of Triactinomyxon spores that infect trout and cause whirling disease. Worms are small, 2-8 cm long, reddish, and often in patches in streams carrying organically rich hatchery effluents or in earthen trout ponds. Earthen ponds and raceways stocked with fish where cleaning is difficult or neglected are ideal habitats for worms and, once introduced, the whirling disease parasite becomes established. Techniques for prevention are periodical disinfection of the facility and the rearing of small trout indoors in pathogen-free water. Smooth-faced concrete or plastic-lined raceways that are kept clean and free of contaminated water keep the facility free of the disease. Disinfection of waterborne infectivity has also been effective and can be achieved by combining filtration to remove or reduce suspended contaminants with ultraviolet-irradiation (Hoffman 1974, 1975). Some chemotherapeutants reduced losses and infection of young trout, but none prevented or totally eliminated whirling disease. Development of spores decreased when young trout were fed furazolidone (Taylor et al. 1973); furoxone, benomyl, and fumagillin (O'Grodnick and Gustafson 1974, 1975); or proguanil and clamoxyquin (AIderman 1986). El­Matbouli and Hoffmann (1991) reported recently that fumagillin, fed to experimentally infected rainbow trout, defected morphology of M. cerebralis spores and could prevent a clinical outbreak of whirling disease. Chlorine (sodium hypochlorite), administered weekly for 4 months at concentrations of 0.5 ppm for 2 h to control waterborne infectivity (triactinomyxons) and infected tubificids, suppressed the prevalence of infection by 73% in one group of young trout and by 63% in another group of concurrently exposed young trout (Markiw, unpublished). This chlorine treatment regime was not toxic to trout. A rapid vital staining with fluorescein diacetate and propidium iodide applied to the spores (Markiw 1992a) could be useful for screening the effectiveness of candidate therapeutants that can be used for controlling whirling disease of fish before proceeding to time-consuming in vivo exposure. Immune response of fish to the whirling disease pathogen is critical for vaccination against the disease. Immune response in rainbow trout to M. cerebralis was studied by Halliday (1974), Pauley (1974), and Griffin and Davis (1978). These studies revealed some evidence that rainbow trout produce antibodies against M. cerebralis, but protection against infection has not been demonstrated. The immune response to Triactinomyxon has not been examined. Recent studies demonstrated, however, that host tissue reaction against the pathogen decreased or even eliminated myxosporean infection in lightly infected rainbow trout (Markiw 1992a). This indicates that immunization against whirling disease may work with common specific antigenic components of both stages for producing an immunogen by genetic engineering. In the past, radical methods of controlling the disease in affected trout hatcheries were used. Infected fish stocks were destroyed and buried and the entire facility disinfected. Present methods of control are less drastic. The approaches to managing hatcheries with infected fish are comprehensively discussed by Hoffman (1990). When fish of a hatchery are infected, the intensity of infection determines what can be done with the infected individuals. The infected fish may be slaughtered and smoked for table use (smoking kills the spores; Wolf and Markiw 1982) or placed in enzootic areas. Such arrangements may reduce economic loss to fish culturists. The survey of watersheds for the source of infectivity with susceptible sentinel trout in floating cages (Hnath 1970; Horsch 1987) and the use of more sensitive methods of spore detection help pinpoint contaminated areas. The extent of whirling disease is determined by the infection intensity, not simply by the presence or absence of M. cerebralis. Therefore, control measures do not need to eradicate the parasite completely to be effective. Measures such as culturing resistant species, filtering the water supply, chemotherapy, and periodical disinfection of the facility reduce the potential for establishment of myxosporean infection in fish and actinosporean infection in tubificids and greatly reduce the number of infected individuals and the intensity of the infection. Whirling disease can also be reduced if fish are inspected and certified as disease­free before transfer between facilities. References Alderman, D. J. 1986. Whirling disease chemotherapy. Bulletin of the European Association of Fish Pathologists 6 38­39. El-Matbouli, M., and R. Hoffmann. 1989. Experimental transmission of two Myxobolus spp. developing bisporogeny via tubificid worms. Parasitology Research 75:461­464. El-Matbouli, M., and R. Hoffman. 1991. Prevention of experimentally induced whirling disease in rainbow trout Oncorhynchus mykiss by Fumagillin. Diseases of Aquatic Organisms 10:109-113. Elson, K. G. R. 1969. Whirling disease in trout. 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Salmonid fish farming in England and Wales; the major diseases of fish--trends in incidence and methods of control. State Veterinary Journal 39:91­109. Lom, J., and G. L. Hoffman. 1971. Morphology of the spore of Myxosoma cerebralis (Hofer, 1903) and Myxosoma cartilaginis (Hoffman, Putz, and Dunbar, 1965). Journal of Parasitology 57:1302­1308. Lorz, H. V., A. Amandi, C. R. Banner, and J. S. Rohovec. 1989. Detection of Myxobolus (Myxosoma) cerebralis in salmonid fish in Oregon. Journal of Aquatic Animal Health 1:217­221. Maloney, R., R. J. Cawthorn, M. Markiw, and D. Groman. 1991. The occurrence of Myxobolus neurobius (Myxosporea) in wild young Atlantic salmon and arctic char in Newfoundland. Journal of Aquatic Animal Health 5:146­147. Markiw, M. E. 1986. Salmonid whirling disease: dynamics of experimental production of the infective stage-the triactinomyxon spore. Canadian Journal of Fisheries and Aquatic Sciences 43:521­526. Markiw, M. E. 1989a. Salmonid whirling disease: myxosporean and actinosporean stages cross­react in direct fluorescent antibody test. Journal of Fish Diseases 12:137­141. Markiw, M. E. 1989b. Portals of entry for salmonid whirling disease in rainbow trout. Diseases of Aquatic Organisms 6:7­10. Markiw, M. E. 1991. Whirling disease: earliest susceptible age of rainbow trout to the triactinomyxon of Myxobolus cerebralis. Aquaculture 92:1­6. Markiw, M. E. 1992a. Experimentally induced whirling disease. I. Dose response of fry and adults of rainbow trout to the triactinomyxon stage of Myxobolus cerebralis. Journal of Aquatic Animal Health 4(1):40-43. Markiw, M. E. 1992b. Experimentally induced whirling disease. II. Determination of longevity of the infective triactinomyxon stage of Myxobolus cerebralis by vital staining. Journal of Aquatic Animal Health 4(1):44-47. Markiw, M. E., and K. Wolf. 1974a. Myxosoma cerebralis: isolation and concentration from fish skeletal elements-sequential enzymatic digestions and purification by differential centrifugation. Journal of the Fisheries Research Board of Canada 31:15­20. Markiw, M. E, and K. Wolf. 1974b. Myxosoma cerebralis: comparative sensitivity of spore detection methods. Journal of the Fisheries Research Board of Canada 31:1597­1600. Markiw, M. E, and K. Wolf. 1978. Myxosoma cerebralis: fluorescent antibody techniques for antigen recognition. Journal of the Fisheries Research Board of Canada 35:828­832. Markiw, M. E, and K Wolf. 1980. Myxosoma cerebralis: trypsinization of plankton centrifuge harvests increases optical clarity and spore concentration. Canadian Journal of Fisheries and Aquatic Sciences 37:2225­2227. Markiw, M. E., and K. Wolf. 1983. Myxosoma cerebralis (Myxozoa: Myxosporea) etiologic agent of salmonid whirling disease requires tubificid worm (Annelida: Oligochaeta) in its life cycle. Journal of Protozoology 30:561­564. O'Brien, D. J. 1976. Some aspects of diseases of freshwater fish in Ireland. Irish Veterinary Journal 30:97­100. O'Grodnick, J. J. 1975a. Whirling disease (Myxosoma cerebralis) spore concentration using the continuous plankton centrifuge. Journal of Wildlife Diseases 11:54­57. O'Grodnick, J. J. 1975b. Egg transmission of whirling disease. Progressive Fish-Culturist 37:153­154. O'Grodnick, J. J. 1979. Susceptibility of various salmonids to whirling disease (Myxosoma cerebralis). Transactions of the American Fisheries Society 108:187­190. O'Grodnick, J. J., and C. C. Gustafson. 1974. A study of the transmission, life history and control of whirling disease of trout. Pennsylvania Fish Commission, Federal Aid in Fish Restoration, Progress Report F­35-R-6. 31 pp. [Mimeograph] O'Grodnick, J. J., and C. C. Gustafson. 1975. A study of the transmission, life history and control of whirling disease of trout. Pennsylvania Fish Commission, Federal Aid in Fish Restoration, Progress Report F-35-R-7. 34 pp. [Mimeograph] Pauley, G. B. 1974. Fish sporozoa extraction of antigens from Myxosoma cerebralis spores which mimic tissue antigens of rainbow trout (Salmo gairdneri). Journal of the Fisheries Research Board of Canada 31:1481­1484. Prihoda, J. 1983. Experimental infections of rainbow trout fry with Myxosoma cerebralis Hoffer, 1903. Page 98 in Parasites and parasitic diseases of fish. Proceedings of the First International Symposium of Ichthyoparasitology, Ceske Budejovice, Czechoslovakia, 8­13 August. [Abstract] Pugachev, O. N., and P. P. Khokhlov. 1979. Myxosporidia of genus Myxobolus--parasites from central nervous system of salmoniformes fish. Pages 137­139 in Systematics and Ecology of Fish from Continental Watersheds of Far East. USSR Academy of Sciences Press, Vladivostok. [In Russian] Schuberg, A., and O. Schroeder. 1905. Myxosporidien aus dem Nervensystem und der Haut der Bachforelle (Myxobolus neurobius n.sp.u. Henneguya nusslini n.sp.). Archiv für Protistenkunde 6:47-60. Taylor, R. E. L., S. J. Coli, and D. R. Junell. 1973. Control of whirling disease by continuous drug feeding. Journal of Wildlife Diseases 9:302­305. Taylor, R. L., and M. Lott. 1978. Transmission of salmonid whirling disease by birds fed trout infected with Myxosoma cerebralis. Journal of Protozoology 25:105­106. Uspenskaya, A. V. 1955. Biology, distribution and economic importance of Myxosoma cerebralis, the causative agent of twist disease in trout. Lectures of the Academy of Science, USSR, 105: 1132­1135. Uspenskaya, A. V. 1978. Biological peculiarities of the spore stage of Myxosoma cerebralis (Myxosporida: Myxosomatidae). Parasitologiya 12:15­20. [Russian; English summary] USSR Academy of Sciences. 1972. Aquatic oligochaeta worms: taxonomy, ecology and faunistic studies in the USSR. Published by Nauka Publishers, Moskow, USSR. 211 pp. [Translated from Russian by the U.S. Department of the Interior and the National Science Foundation, Washington, D.C.] Wolf, K., and M. E. Markiw. 1982. Myxosoma cerebralis inactivation of spores by hot smoking of infected trout. Canadian Journal of Fisheries and Aquatic Science 39:926­928. Wolf, K., and M. E. Markiw. 1984. Biology contravenes taxonomy in the Myxozoa: new discoveries show alternation of invertebrate and vertebrate hosts. Science 225:1449­1452. Wolf, K., M. E. Markiw, and J. K. Hiltunen. 1986. Salmonid whirling disease: Tubifex tubifex (Müller) identified as the essential oligochaete in the protozoan life cycle. Journal of Fish Diseases 9:83­85. Wolf, K., M. E. Markiw, J. Machado Cruz, M. H. Galhano, J. Eiras, and R. L. Herman. 1981. Non-myxosporidan blacktail of salmonids. Journal of Fish Diseases 4:355­357.     Addendum The following are more recent, selected references which are provided for your information: Boustead, N. C. 1993. Detection and New Zealand distribution of Myxobolus cerebralis, the cause of whirling disease of salmonids. New Zealand Journal of Marine and Freshwater Research 27(4):431-436. Whirling disease, caused by Myxobolus cerebralis, was diagnosed in rainbow trout (Oncorhynchus mykiss) at Silverstream Fish Hatchery near Christchurch in 1980. As a consequence, a nationwide survey for M. cerebralis in salmonids was conducted by examination of 5307 wild and hatchery fish. In addition, sentinel rainbow trout were used at six locations to test for whirling disease. This survey and other studies revealed M. cerebralis at locations in the Waimakariri, Rakaia, and Rangitata River catchments, and provide the first New Zealand records of M. cerebralis in brook trout (Salvelinus fontinalis), chinook salmon (Oncorhynchus tshawytscha), and sockeye salmon (O. nerka). This paper demonstrates use of sentinel rainbow trout in detecting low­level asymptomatic infection of M. cerebralis in chinook salmon. Bruno, D. W. 1991. Whirling disease. Aquaculture Information Series, No. 13. Marine Laboratory, Aberdeen, Scotland. 4 pp. A general account of Myxobolus cerebralis infection of fishes is given under the headings: identification and life cycle; transmission; diagnosis; pathology; hosts; control and prevention; distribution within Europe. El-Matbouli, M., and R. W. Hoffmann. 1990. Erste Efahrungen mit Fumagillin bei Drehkrankheit. (Exploratory trial of fumagillin in whirling disease of trout). Pages 140-147 in Tagung der Fachgruppe "Fischkrankheiten" der DVG, Schmiedefeld/Thuringen, Germany. 14-16 November 1990. Deutsche Veterinarmedizinische Gesellschaft, Giessen, Germany. Feed containing 0.1% fumagillin was given to 30 young rainbow trout, commencing 30 days after experimental infection with Myxosoma cerebralis and lasting for 18 weeks. None became ill and only small numbers of M. cerebralis spores were recovered from 6, compared with 18 ill and 22 harbouring large numbers of spores among 30 untreated trout. El-Matbouli, M., and R. W. Hoffmann. 1991. Effects of freezing, aging, and passage through the alimentary canal of predatory animals on the viability of Myxobolus cerebralis spores. Journal of Aquatic Animal Health 3(4):260-262. In transmission experiments with tubificids Tubifex tubifex as primary hosts and fry of rainbow trout Oncorhynchus mykiss as secondary hosts, it was shown that Myxobolus cerebralis spores can tolerate freezing at ­20° C for at least 3 months, aging in mud at 13° C for at least 5 months, and passage through the guts of northern pike Esox lucius or mallards Anas platyrhynchos without loss of infectivity. El-Matbouli, M., and R. W. Hoffmann. 1991. Transmissionsversuche mit Myxobolus cerebralis und Myxobolus pavlovskii und ihre Entwicklung in Tubificiden. (Experimental transmission of Myxobolus cerebralis and Myxobolus pavlovskii and their development in tubificids). Fisherei Forschung 29(3):70-75. Transmission experiments with M. cerebralis and M. pavlovskii show that the development of infectivity for fish is an exogenous process in which a tubificid oligochaete (Tubifex tubifex) play an essential role. M. cerebralis develops in the intestine epithelial cells of T. tubifex to Triactinomyxon, whereas M. pavlovskii develops to Hexactinomyxon, which are only infective for rainbow trout (Oncorhynchus mykiss) and silver carp (Hypophthalmichthys mollitrix), respectively. El-Matbouli, M., T. Fischer-Scherl, and R. W. Hoffman. 1992. Present knowledge on the life cycle, taxonomy, pathology, and therapy of some Myxosporea spp. important for freshwater fish. Annual Review of Fish Diseases 2:367-402. Some important myxosporean species of freshwater fishes are described and illustrated in this review, with special reference to Myxobolus cerebralis, Sphaerospora renicola and Hoferellus carassii. Detailed information on the life cycles of M. cerebralis, M. cotti and M. pavlovskii is presented proving that these myxosporeans develop in the intermediate host, Tubifex tubifex, to actinosporeans which represent the only infective stage for susceptible fish. These findings imply that the present taxonomy for the Phylum Myxozoa needs revision. Trends for treatment of myxosporean associated diseases are summarized with special emphasis on the antibiotic Fumagillin DCH. El-Matbouli, M., and R. W. Hoffmann. 1995. Recent developments of Myxosporeans in fish. Second European Congress of Protistology and Eighth European Conference on Ciliate Biology, Clermon-Ferrand, France. 21-26 July 1995. European Journal of Protistology 31(4):424. [Abstract]. El-Matbouli, M., R. W. Hoffmann, and C. Mandok. 1995. Light and electron microscopic observations on the route of the triactinomyxon-sporoplasm of Myxobolus cerebralis from epidermis into rainbow trout cartilage. Journal of Fish Biology 46(6):919-935. Penetration of triactinomyxon­sporoplasma of Myxobolus cerebralis through skin, fins, gills and buccal cavity have been demonstrated experimentally in rainbow trout. Furthermore the multiplication­stages of penetrated triactinomyxon­sporoplasms reach the cartilage via peripheral nerves and the central nervous system (CNS). This is in contrast to the assumption that the agent reaches the cartilage via blood, lymph, and/or coelomic fluid. During the first hour following penetration, the sporoplasm migrates between the epidermal cells. Then, it enters the epithelia and multiplies intracellularly. These stages migrate deeper into the subcutis, then through the peripheral nerves and CNS. After about 21 days the parasites reach the head cartilages. During their migration they also multiply to increase parasite numbers. The ultrastructure of the proliferative phase (presporogonic development) and sporogonic phase of the life cycle are demonstrated and discussed. Garden, O. 1992. The Myxosporea of fish: a review. British Veterinary Journal 148(3):223-239. This review describes the current views on the biology of the Myxosporea, and considers the more important myxosporean diseases of fishes in some detail (whirling disease, caused by Myxobolus cerebralis; renal sphaerosporosis, caused by Sphaerospora renicola; proliferative kidney disease, caused by Myxidium lieberkuhni; kidney enlargement disease, caused by Hoferellus cyprini; bi­ and multi-valvulid myxosporeans, including Kudoa and Henneguya; ceratomyxosis, caused by Ceratomyxa shasta; sphaerosporosis, caused by S. molnari). Hoffman, G. L. 1990. Myxobolus cerebralis, a worldwide cause of salmonid whirling disease. Journal of Aquatic Animal Health 2(1):30-37. Salmonid whirling disease was discovered in Europe in 1893 and has since been spread around the world with shipments of cultured and wild fish. The causative agent is the protozoan parasite Myxobolus cerebralis, whose taxonomy and life cycle remain controversial. It cause lesions of salmonid skeletal tissues, particularly in the head. Diagnosis usually is straightforward, but can be complicated by the presence of other Myxobolus species and by other parasites that cause fish to whirl. The pathogen is difficult to eradicate, but its seriousness can be managed if fish are cultured in spore­free source water, in concrete raceways with strong water flows, or in ponds that are regularly disinfected, and if they are constantly monitored for the presence of spores. Fish can carry M. cerebralis spores and still be healthy. Such fish may be suitable for stocking into waters already containing M. cerebralis, and even for human consumption. Hoffmann, R. W., M. El-Matbouli, and G. Hoffmann-Fezer. 1991. Nachweis von Antikorpern gegen Myxobolus cerebralis bei Regenbogenforellen (Oncorhynchus mykiss). (Demonstration of antibodies to Myxobolus cerebralis in rainbow trout (Oncorhynchus mykiss). Pages 136-139 in Tagung der Fachgruppe "Fischkrankheiten" der DVG, Schmiedefeld/Thuringen, Germany. 14-16 November 1990. Deutsche Veterinarmedizinische Gesellschaft, Giessen, Germany. In order to detect serum antibodies to spores of M. cerebralis, the spores were incubated with trout serum, and any bound IgG was detected by peroxidase­labelled rabbit immune serum against trout IgG. Twenty-nine of 49 serum samples from infected trout were positive, and all samples from uninfected trout were negative. Hoffmann, R. W., M. El-Matbouli, and G. Hoffmann-Fezer. 1994. Immunoreactions against the agent of whirling disease in rainbow trout (Oncorhynchus mykiss). Page W-10.4 in Program and Abstracts: International Symposium on Aquatic Animal Health, Seattle, Wash. 4-8 September 1994. [Abstract] Penetration of triactinomyxon­sporoplasm, causing whirling disease of salmonids, through skin, fins, gills and buccal cavity have been experimentally demonstrated in rainbow trout (Oncorhynchus mykiss). Within two weeks the multiplication­stages of the penetrated triactinomyxon­sporoplasm reach the cartilage via peripheral nerves and the central nervous system (CNS). This is in contrast to the assumption that the agent reaches the cartilage via the blood, lymph, and/or coelomic fluid. This short time is sufficient to induce an immune reaction which could be shown by antibodies against spores of Myxobolis cerebralis in infected rainbow trout. Rainbow trout fingerlings experimentally infected with whirling disease were exposed to triactinomyxon spores of M. cerebralis a second time about four weeks after begin of clinical whirling disease. In contrast to controls, no spores penetrating via skin, gills or otherwise could be detected in any of the reinfected fish. Since all triactinomyxon were intact-­in contrast to those exposed to non infected fish­-it seems that a whirling disease infected fish cannot be recognized by the triactinomyxon. Whereas the function of these antibodies has not yet been eluciated, it was demonstrated that rainbow trout once infected with whirling disease are protected. This proves an acquired immunity against M. cerebralis with respect to its triactinomyxon stage. Practical consequences are discussed. Kent, M. L., L. Margolis, and J. O. Corliss. 1994. The demise of a class of protists: taxonomic and nomenclatural revisions proposed for the protist phylum Myxozoa Grasse, 1970. Canadian Journal of Zoology 72(5):932-937. The phylum Myxozoa has been considered to comprise two classes, Myxosporea (primarily infecting fishes) and Actinosporea (primarily infecting aquatic oligochaetes). However, a 2­host life cycle involving both myxosporean and actinosporean stages has now been demonstrated for 14 myxosporean species belonging to 6 genera in 4 families. In light of these findings, the taxonomy of the Myxozoa is revised. It is proposed that the class Actinosporea, the order Actinomyxidia and all families in the Actinosporea except Tetractinomyxidae be suppressed. This family and its one genus, Tetractinomyxon, are transferred to the order Multivalvulida (Myxosporea). It is also proposed that actinosporean generic names be treated as collective­group names so they do not compete in priority with myxosporean generic names. Triactinomyxon dubium and Triactinomyxon gyrosalmo are suppressed as junior synonyms of Myxobolus cerebralis. Other actinosporean species are retained as species inquirendae until their myxosporean stages are identified. A revised description of the phylum Myxozoa is provided that includes these proposed taxonomic and nomenclatural changes. Lom, J., and I Dykova. 1992. Fine structure of Triactinomyxon early stages and sporogony: Myxosporean and Actinosporean features compared. Journal of Protozoology 39(1):16-27. The first ultrastructural study of the actinosporean genus Triactinomyxon was carried out on Triactinomyxon legeri from the intestinal epithelium of Tubifex tubifex. The developmental cycle starts with bi­ and uninucleate cells. We propose that these cells may be an early proliferative phase of the cycle and may unite to give rise to the four­cell stage, initiating pansporoblast formation. Valvogenic cells transform in the long stylus and anchor­like projections of the spore. In the capsulogenic cells, the primordium of the polar capsules originates as a simple, dense, club­shaped structure not observed in other actinosporeans. In all other respects, actinosporean ultrastructure follows more or less similar patterns. Comparison of actinosporean and myxosporean species gives evidence of considerable structural similarity, exemplified in both classes by the occurrence of cell junctions in their multicellular spores, identical polar capsules and their morphogenesis, cell­in­cell condition, pansporoblast formation, and presence of dense bodies (sporoplasmosomes) primarily in the sporoplasm. This unity of patterns speaks in favor of the postulated actinosporean­myxosporean transformation, which warrants further study. Molnar, K. 1993. Recent achievements in the chemotherapy of myxosporean infections of fish. Acta Veterinaria Hungarica 41(1-2):51-58. Furazolidone, the only effective drug which can be successfully applied against several myxosporeoses in the form of prolonged feeding has been used with good results against sphaerosporosis of common carp, hoferellosis of goldfish, myxidiosis of eel, and proliferative kidney disease, whirling disease and ceratomyxosis of salmonids. A 4­ to 6­day feeding of medicated diets containing 0.05­0.1% furazolidone was effective against early blood stages of Sphaerospora, while toltrazuril failed to kill these parasites (data are summarized in 2 tables). Siddall, M. E., D. S. Martin, D. Bridge, S. S. Desser, and D. K. Cone. 1995. The demise of a phylum of protists - phylogeny of Myxozoa and other parasitic Cnidaria. Journal of Parasitology 81(6):961-967. The notion that members of the phylum Myxozoa Grasse, 1970 do not properly belong in classifications of protists has frequently been suggested because the infective spores of these parasites are not unicellular. Systematists have failed to be decisive about myxozoan phylogenetic affinities, either finding the suggestion of a cnidarian connection to be preposterous or considering the recent suggestion of a relationship with nematodes to be an obvious failure of molecular phylogenetics. Thus, the group has remained in classifications as a protistan phylum in its own right. The ultrastructure of the development of myxozoans was critically re­examined in order to more fully explore the possibility of morphological synapomorphies with metazoan taxa. These morphological characters, in combination with small ribosomal subunit gene sequences, were used in a phylogenetic analysis in order to assess myxozoan origins. The results unequivocally support the inclusion of myxozoans as a class of highly derived parasitic cnidarians, and as sister taxon to the narcomedusan Polypodium hydriforme. Reassessment of myxozoans as metazoans reveals terminal differentiation, typical metazoan cellular junctions, and collagen production. Their "polar capsules" are redescribed as typical nematocysts bearing atrichous isorhiza. Insofar as taxa cannot be contained within other taxa of equal rank, the phylum Myxozoa is abandoned and it is recommended that the group as a whole be removed from all protistan classifications and placed in a more comprehensive cnidarian system. Uspenskaya, A. V. 1994. New problems in the study of Myxozoa. Parazitologiya 27(5):369-374. The problems arising in the study of Myxosporea, since the discovery that the life­cycle of Myxosoma cerebralis includes an alternation of invertebrate (oligochaete) and vertebrate (fish) hosts and the presence of a developmental stage corresponding to that of Actinosporea, previously considered as a separate class of the phylum Myxozoa are considered. Similar life cycles have also been described for some species of Myxobolus, while other studies demonstrate a direct development cycle, without intermediate host and actinosporidian stage. The implication of these studies for the taxonomy and evolution of Myxozoa are discussed. Future research needs to include the study of the biological cycle of every species of Myxozoa in order to link known species of actinosporidia to species of myxosporidia, using laboratory­bred uninfected fish and oligochaetes and modern technology including molecular biology. Studies of the marine and freshwater actinosporidian faunas are also needed. Walker, P. G., and R. B. Nehring. 1995. An investigation to determine the cause(s) of the disappearance of young wild rainbow trout in the upper Colorado River, in Middle Park, Colorado. Colorado Division of Wildlife, Denver, Colorado. 134 pp. U.S. Department of the Interior || U.S. Geological Survey 11649 Leetown Road, Kearneysville, WV 25430, USA URL: http://www.lsc.usgs.gov Maintainer: lsc_webmaster@usgs.gov Last Modified: February 26, 2004 dwn Privacy Policy and Disclaimers || FOIA || Accessibility