A review of what is known about Myxobolus cerebralis (Whirling Disease)
Introduction:
Myxobolus cerebralis is a parasite that can be linked to the mortality of 90% of certain age classes of rainbow trout (Hedrick et al, 1998), both in the wild and in aquaculture. M. cerebralis has been shown to infect 17 species of salmonid fish to varying degrees (Halliday, 1976). Its life cycle has been studied extensively, yet little is known about host-parasite interactions, or immune responses (Hedrick et al, 2001). It seems that susceptible fish are unable to fight off infections, and many develop severe clinical signs, which decrease the marketability of such fish (Hedrick et al, 1998). Because of its effect on important fisheries and game fish, many researchers are working on ways to control the effects of this disease (El-Matbouli and Hoffmann, 1991).
Life cycle:
Myxobolus cerebralis exhibits a two-host life cycle, spending part of its life cycle in a salmoniod fish and the other in Tubifex tubifex an oligocheate worm. Infective spores are ingested by T. tubifex and attach to the gut of the worm. The sporoplasm migrates to the gut epithelium where it undergoes schizogony intercellularly. The single cells then undergo gametogony. Sporogony occurs when the zygote has eight somatic cells, which then form the pansporocyst. The cells within the pansporocyst differentiate and the triactinomyxon spore is shed into the water column from the digestive tract. This occurs approximately three months after infection (El-Matbouli and Hoffmann, 1998). The release of triactinomyxon spores can occur for up to twelve months after infection (El-Matbouli et al, 1995).
The triactinomyxon spores attach to the fish’s epidermis with its polar capsules. By the first hour, the sporoplasm is located intracellularly in the epidermis (El-Matbouli et al, 1995). There they multiply intracellularly, in a presporogenic stage and move into the central nervous system. By using the host’s nervous system as a “highway” to reach the cartilaginous areas of the spine has been hypothesized as a method to avoid the host’s internal defense system (Hedrick et al, 1998). Once located in the spinal area the parasite produces a plasmodium that can lyse the cartilage (El-Matbouli et al, 1995). Over 2 million spores can be developed with in the cartilage (Hedrick et al, 1998).
History:
M. cerebralis was first in the diagnosed in the US in 1958 (Hoffman, 1990). At that time it was believed to have introduce to the US from imported European rainbow trout. This hypothesis is supported by the European brown trout’s apparent ability to survive infection from M. cerebralis and the fact that the first ever reported case was in Germany, 1903, when non-native rainbow trout were introduced to European waters (Halliday, 1976).
Extensive studies were preformed to discover the method of transmission, but very little headway had been made until 1983, when Markiw and Wolf discovered an alternate host for M. cerebralis (Brinkhurst, 1997). The triactinomyxon spore produce in T. tubifex was determined to be the infective agent of the trout. Prior to that the spore had been considered a different species belonging to a separate class, Actinosporea (Siddall et al, 1995). Later it was discovered that most of the members of “Actinosporea” were the alternate stages of Myxosporea. In 1994 it was proposed that Actinosporea no longer be considered a class, but be used in reference to a stage of the life cycle (Kent et al, 2001). There is still controversy over the naming of new Myxosporeans.
In addition to the controversy over the classes of Myxosporea, there is now controversy over whether or not Myxosporea is its own phylum in the Protozoa, or if it belongs in the Metazoa grouped with the parasitic cnidarians. The idea itself is not new, it was first proposed in 1899, citing the similarities of the polar capsules to nematocysts (Kent et al, 2001). However, it was not until the advent of molecular analysis that the idea became more main stream. Hombox and 18s rDNA sequences have both placed the Myxozoans within the Metazoa (Hedrick et al, 1998). A detailed examination of the development of nematocysts and polar capsules has shown considerable parallels (Siddall et al, 1995).
Identification:
Most identification includes the showing of clinical signs in an infected fish. These include a whirling behavior, blackened caudal fin, and spinal deformities. The whirling behavior and the blackened tail are results of pressure on certain nerves by the growth of M. cerebralis in the central nervous system (Halliday, 1976). However there has been cases of misidentification of the disease. A blackened tail and spinal deformities may be the result of vitamin C deficiency (Hoffman, 1990).
More reliable methods of testing fish for the presence of M. cerebralis have been identified. Light microscopy of stained tissues was one such method, but could only be used 2-3 months after infection (Antonio et al, 1999). The use of PCR to amplify parasite DNA has been shown effective for very low spore count, which would not be detected visually (Hedrick et al, 1998)
Antonio, El-Matbouli, and Hedrick devised a method which allow detection of the parasite immediately upon infection in T. tubifex and within 15 days of infection for rainbow trout (1999). This method, in situ hybridization, used DNA probes to find and tag areas within the fish, or worm, that housed the parasite allowing an opportunity to follow the path of the parasite as well.
Host specificity:
Several studies have been preformed on host specificity and susceptibility. By far the most susceptible fish is the rainbow trout, Oncorhychus mykiss (El-Matbouli et al, 1999, Hedrick et al, 1999). However, the presence of M. cerebralis has been recorded in 17 different fish species (Halliday, 1976). One of the studies, (Hedrick et al, 1999) found that Westslope cutthroat trout were as susceptible as rainbow trout based on spore count and number of fish showing clinical signs. One the opposing side, there have been two studies showing that Coho salmon are resistant to M. cerebralis (Hedrick et al, 1999 and Hedrick et al, 2001). Coho salmon were found to have the spores present on the body and entering the epidermis, but no spores were found in the dissected fish (Hedrick et al, 2001). It is unknown how the salmon resists infection.
When presented with the “wrong” hosts, such as goldfish, carp, or guppies, it has been found that the triactinomyxon spores fail to release there polar capsules or germ cells in large amounts (El-Matbouli, 1999). In addition to the chemical cues needed, El-Matbouli postulated that there must be some mechanical cues as well. When place with mucus and fin parts from correct and incorrect hosts, the spores failed to release the germ cells in large amounts. He also observed that infected fish tended to jerk and swim rapidly when first infected.
While numerous studies have been preformed on various salmonid and non-salmonid fishes with respect to host specificity, few have addressed the issue of the host specificity from the oligochaete side. Brinkhurst (1997) points this possibility out in a paper that reviews the oligochaete’s role in transmission of the mxyozoan parasites. He states that most cultures of T. tubifex obtained commercially can be a mixture of tubifecid worms, or Lumbriculus variegatus which is sold under the tubifex name.
Transmission conditions:
When first described, much thought was given to how the M. cerebralis spores could be transmitted. Halliday, 1976, summed up some of the ways that were commonly thought to be ways of transmitting the spore. These ways were through transportation of infected eggs and live or dead infected fish. Markiw (1991) found that eggs could not be infected with the triactinomyxid spore, and were thus ruled out as a transmission pathway.
Live or dead fish could still be a viable pathway for M. cerebralis, as well as predation. A study which showed that spores remained viable after passing through several predatory avians and fish digestive tracts, allowing the spores the possibility of moving to a M. cerebralis free site (El-Matbouli et al, 1991). This same study showed that spores remained viable after 5 months, and freezing allowing the possibility of a spore over-wintering.
However, temperature does play a very important role in the development of the triactinomyxid spores. When tubificid worms were held at 5 degrees Celsius, the development of these spores were slowed, when the worms were held at 25 and 30 degrees, the worms stopped producing spores entirely (El-Matbouli et al, 1999). While temperature “cured” worms no longer had any stages of M. cerebralis, within them, they could still be re-infected and were found to produce spores longer that the single time infected worm. The optimum temperature for triactinomyxid spores to develop was found to be approximately 15 degrees Celsius (El-Matbouli et al, 1999).
Treatment:
Most early treatments were limited to the prevention of the disease only. Hoffman suggested raising trout in concrete raceways rather than earthen pools in order to reduce the possibility of the ponds harboring worms or triactinomyxon spores (1991). In addition, he suggested that trout should be raised in well water until they were at least six inches long, if not for their entire life, also as a measure to reduce the chance of exposure to triactinomyxon spores. The younger fish are more susceptible to developing clinical signs of M. cerebralis as their skeleton has not ossified. The earliest date of infection has been recorded at two days post hatching (Markiw, 1991).
Other ways of destroying the triactinomyxon spores have included UV lighting and ozone filtering of the water, which are considered to be effective ways to clean the water of these spores (Hedrick et al, 1998). More traditional modes of curing the fish are feeding the trout with chemicals and antibiotics. Fumagillin has been shown to be effective in reducing the clinical signs of infected fish, and to reduce spore count (El-Matbouli and Hoffmann, 1991). This treatment must be administered within 14 days of infection to reduce spore count. In order to prevent infection the fish must be feed fumagillin for over a year, till ossification has occurred.
The possibility of the existence of a resistant strain of rainbow trout has been sought after by aquaculturists and scientists alike, but only one possible strain has been identified and is being tested (Kent et al, 2001). It is very uncertain if this strain is truly resistant to M. cerebralis. Other studies have shown that despite the supposedly high selection pressures to develop a resistance to M. cerebralis, there is no difference between the spore count or mortality of a fish that came from parents from uninfected streams, and those that were from infected streams (Ryce et al, 2001).
Conclusions:
Knowledge of M. cerebralis life cycle and transmissions has increased considerably within the past 10 years. There is still much to be learned about controlling M. cerebralis, and about Myxozoans in general. The taxonomy of the Myxozoa is still debatable, new life cycles, and species are constantly being described. There are many areas of research open within this group of organisms.
Works Cited:
Antonio, D B, M El-Matbouli, R P Hedrick. 1999. Detection of early developmental
stages of Myxobolus cerebralis in fish and tubificid oligocheate hosts by in situ hybridization. Parasitology Research. 85: 942-944
Brinkhust, Ralph O. 1996. On the role of tubificid Oligochaetes in relation to fish disease
with a special reference to the Myxozoa. Annual review of fish diseases. 6: 29-40
a El-Matbouli, R W Hoffmann. 1991. Prevention of experimentally induced whirling
disease in rainbow trout Oncorhynchus mykiss by Fumagillin. Diseases of Aquatic Organisms. 10: 109-113
b El-Matbouli, M, 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 Heath. 3: 260-262
El-Matbouli, M, R W Hoffmann, C Mandok. 1995. Light and electron microscope
observations on the route of the triactinomyxon-sporoplasm of Myxobolus
cerebralis from the epidermis into rainbow trout cartilage. Journal of Fish Biology. 46: 919-935
El-Matbouli, M, R W Hoffmann. 1997. Light and electron microscope studies on the
chronological development of Myxobolus cerebralis to the actinosporean stage in Tubifex tubifex. International Journal for Parasitology. 28: 195-217
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disease: Host specificity and interaction between the actinosporean stage of Myxobolus cerebralis and the rainbow trout Oncorhynchus mykiss. Diseases of Aquatic Organisms. 35: 1-12
El-Matbouli, M, T S McDowell, D B Antonio, K B Andree, R P Hedrick. 1999. Effects
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disease. Journal of Aquatic Animal Health. 2: 30-37
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phylum of protists: Phylogeny of the Myxozoa and other parasitic Cnidaria. Journal of Parasitology. 81:961-967
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