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whirling disease

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whirling disease

Summary

  • Last modified
  • 14 July 2018
  • Datasheet Type(s)
  • Animal Disease
  • Preferred Scientific Name
  • whirling disease
  • Overview
  • Myxobolus cerebralis is a freshwater parasite which cycles between salmon/trout and oligochaete worms. The myxozoan parasite was unknown until outbreaks of whirling disease occurred in imported rainbow trout in...

  • Principal Source
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    Compendia
    CAB International
    Wallingford
    Oxfordshire
    OX10 8DE
    UK
    compend@cabi.org
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Pictures

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PictureTitleCaptionCopyright
Clinical signs of whirling disease in rainbow trout. Fish were infected with Myxobolus cerebralis as fry in the laboratory through co-habitation with infected Tubifex tubifex. (a) Live fish have distinct black tails and exhibit whirling behavior when disturbed. (b) Skeletal deformities in fish co-habited with infected worms for 6 weeks post hatch, then held on well water for 5 months and euthanized.
TitleClinical signs of whirling disease
CaptionClinical signs of whirling disease in rainbow trout. Fish were infected with Myxobolus cerebralis as fry in the laboratory through co-habitation with infected Tubifex tubifex. (a) Live fish have distinct black tails and exhibit whirling behavior when disturbed. (b) Skeletal deformities in fish co-habited with infected worms for 6 weeks post hatch, then held on well water for 5 months and euthanized.
Copyright©Stephen Atkinson & Sascha Hallett/Oregon State University, USA.
Clinical signs of whirling disease in rainbow trout. Fish were infected with Myxobolus cerebralis as fry in the laboratory through co-habitation with infected Tubifex tubifex. (a) Live fish have distinct black tails and exhibit whirling behavior when disturbed. (b) Skeletal deformities in fish co-habited with infected worms for 6 weeks post hatch, then held on well water for 5 months and euthanized.
Clinical signs of whirling diseaseClinical signs of whirling disease in rainbow trout. Fish were infected with Myxobolus cerebralis as fry in the laboratory through co-habitation with infected Tubifex tubifex. (a) Live fish have distinct black tails and exhibit whirling behavior when disturbed. (b) Skeletal deformities in fish co-habited with infected worms for 6 weeks post hatch, then held on well water for 5 months and euthanized.©Stephen Atkinson & Sascha Hallett/Oregon State University, USA.
Life cycle of Myxobolus cerebralis. Triactinomyxon actinospores released into freshwater from infected Tubifex tubifex oligochaetes develop into myxobolid myxospores in the cartilage of salmonid fish and can cause whirling disease.
TitleLife cycle
CaptionLife cycle of Myxobolus cerebralis. Triactinomyxon actinospores released into freshwater from infected Tubifex tubifex oligochaetes develop into myxobolid myxospores in the cartilage of salmonid fish and can cause whirling disease.
Copyright©Stephen Atkinson/Oregon State University
Life cycle of Myxobolus cerebralis. Triactinomyxon actinospores released into freshwater from infected Tubifex tubifex oligochaetes develop into myxobolid myxospores in the cartilage of salmonid fish and can cause whirling disease.
Life cycleLife cycle of Myxobolus cerebralis. Triactinomyxon actinospores released into freshwater from infected Tubifex tubifex oligochaetes develop into myxobolid myxospores in the cartilage of salmonid fish and can cause whirling disease.©Stephen Atkinson/Oregon State University
Life cycle counterparts of Myxobolus cerebralis (whirling disease); myxospores are from infected rainbow trout and actinospores from an oligochaete. (Note scale)
TitleMyxospores
CaptionLife cycle counterparts of Myxobolus cerebralis (whirling disease); myxospores are from infected rainbow trout and actinospores from an oligochaete. (Note scale)
Copyright©Stephen Atkinson/Oregon State University
Life cycle counterparts of Myxobolus cerebralis (whirling disease); myxospores are from infected rainbow trout and actinospores from an oligochaete. (Note scale)
MyxosporesLife cycle counterparts of Myxobolus cerebralis (whirling disease); myxospores are from infected rainbow trout and actinospores from an oligochaete. (Note scale)©Stephen Atkinson/Oregon State University
Life cycle counterparts of Myxobolus cerebralis (whirling disease); myxospores are from infected rainbow trout and actinospores from an oligochaete. Actinospore phase contrast; freshly isolated and unstained. (Note scale)
TitleActinospore
CaptionLife cycle counterparts of Myxobolus cerebralis (whirling disease); myxospores are from infected rainbow trout and actinospores from an oligochaete. Actinospore phase contrast; freshly isolated and unstained. (Note scale)
Copyright©Stephen Atkinson/Oregon State University
Life cycle counterparts of Myxobolus cerebralis (whirling disease); myxospores are from infected rainbow trout and actinospores from an oligochaete. Actinospore phase contrast; freshly isolated and unstained. (Note scale)
ActinosporeLife cycle counterparts of Myxobolus cerebralis (whirling disease); myxospores are from infected rainbow trout and actinospores from an oligochaete. Actinospore phase contrast; freshly isolated and unstained. (Note scale)©Stephen Atkinson/Oregon State University

Identity

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Preferred Scientific Name

  • whirling disease

Other Scientific Names

  • whirling disease in salmonids
  • whirling disease in trout

Local Common Names

  • Germany: Drehkrankheit

Overview

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Myxobolus cerebralis is a freshwater parasite which cycles between salmon/trout and oligochaete worms. The myxozoan parasite was unknown until outbreaks of whirling disease occurred in imported rainbow trout in fish hatcheries in Europe, where native brown trout are disease-resistant hosts of the parasite. Both the invertebrate and vertebrate host are widely distributed. This has enabled establishment of the parasite outside its native range, following human-mediated movement of infected fish.

Whirling disease was initially only a problem for fish culture operations. Wild trout populations in Europe were not impacted by the disease (Christensen, 1972) but wild rainbow and cutthroat trout populations in the USA, particularly the Intermountain West, were decimated (Vincent, 1996; Nehring et al., 1998). As a result, whirling disease is widely known as a serious fish health issue and remains problematic in the USA, where it has both ecological and financial impacts on juvenile salmonids. However, the impact has varied considerably, with no effects apparent in some regions, whilst losses persist in others or populations are only just recovering (Steinbach et al., 2009).

M. cerebralis is probably the most thoroughly studied and best known myxozoan species. Its life cycle was the first to be solved for this group of organisms and its DNA among the first sequenced; it remains a model for the Myxosporea.

There are reports of whirling disease in cyprinids, caused by other Myxobolus species (Li, 1989; Xu and Wang, 2001), and of similar diseases in other fish caused by unrelated pathogens (Avtalion and Shlapobersky, 1994; Kimura and Endo, 1979); coverage of this datasheet is confined to the disease in salmonids caused by M. cerebralis.

Hosts/Species Affected

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M. cerebralis is a parasite of fishes of the family Salmonidae (which includes salmon and trout) (Hedrick and El-Matbouli, 2002) and one species of annelid worm, Tubifex tubifex (Wolf et al., 1986), alternating between the two host groups. The invertebrate is the definitive host, as meiosis occurs only within this host (El-Matbouli and Hoffmann, 1998), whereas the vertebrate is the intermediate host; either host may be referred to as an alternate host for the parasite.

Susceptibility within each host group varies widely. For example, rainbow trout (Oncorhynchus mykiss) are highly susceptible whereas Lake trout (Salvelinus namaycush) appear to be resistant (no spores develop) (MacConnell and Vincent, 2002). Similarly, mitochondrial 16S rDNA lineage III T. tubifex are susceptible, whereas lineage IV T. tubifex are not (Beauchamp et al., 2002; Arsan et al., 2007b).

Susceptibility of the fish host also depends on their age. Fish can become infected from 2 days post-hatch (Markiw, 1991) and are most susceptible to infection and more likely to develop disease when they are younger, before their cartilage ossifies and their central nervous system is fully developed (Ryce et al., 2005). For rainbow trout, this window is generally until 9 weeks post-hatch, whereas for Chinook salmon a resistance threshold can develop as soon as 3 weeks post-hatch (Sollid et al., 2003).

Distribution

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M. cerebralis was first described in Europe and is now exotic on four other continents (Asia, Africa, North America and Oceania). The parasite is not an OIE listed pathogen, and there are few published reports of its current distribution in US States. Information has been compiled by the Whirling Disease Foundation, showing detailed occurrence in hatcheries and watersheds, and most of this data can be found on the Whirling Disease Initiative website (Whirling Disease Initiative, 2014). However, inconsistencies in survey methods and differences in the sensitivity of parasite detection methods have caused confusion about where the parasite occurs. Although there are many reports documenting detection of M. cerebralis, several points should be considered when interpreting this data.

- Detections based on disease signs (whirling behaviour) or characteristic spore morphology are unreliable because disease signs are not exclusive to M. cerebralis and because it is often difficult to distinguish between similar myxobolid spores. Confirmation by histological or molecular methods is essential. Thus some records should be considered unconfirmed or unreliable (eg. Japan, Mexico, South America, Canada).

- Detections of parasite DNA alone, although indicative of parasite presence, should be confirmed with further sampling to determine that parasite establishment has occurred. Establishment in some areas (e.g. Alaska hatcheries, some rivers in Oregon) appears transient.

- Many reports fail to distinguish between parasite introduction and establishment of the life cycle. Thus in many cases detection is based on shipments of infected fish received from Europe, but it is likely that the parasite never became established in the new region and there has been no subsequent work to confirm presence (e.g. South Africa, Lebanon and Morocco).

- Although M. cerebralis may have become established in fish culture facilities, hatchery practices have improved and many of these facilities are no longer positive or have been closed, yet remain on distribution lists.

- Surveys of natural fish populations are rare, so invasiveness and effects on natural populations are often unknown.

- Inclusion of some countries in distribution surveys resulted from misinterpretations of original reports (e.g. Korea, Venezuela) and should be considered invalid or unreliable records.

For further discussion see Bartholomew and Reno (2002) and Bartholomew (2012).

Distribution Table

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The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.

Continent/Country/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes

Asia

LebanonAbsent, intercepted onlyIntroduced Not invasive Halliday, 1976Based on FAO report, 1972

Africa

MoroccoAbsent, intercepted onlyIntroduced Not invasive Preudhomme, 1970
South AfricaAbsent, intercepted onlyIntroducedWyk, 1968

North America

USAPresentPresent based on regional distribution.
-AlaskaAbsent, formerly present2007Introduced2006 Not invasive Arsan et al., 2007a; T. Myers, Alaska Department of Fish and Game, Juneau, Alaska, USA, personal communication, 2013M. cerebralis DNA detected at one rearing facility in fish with no clinical disease. Parasite DNA no longer detected in the 3 years after changes in hatchery management; hatchery subsequently closed.
-ArizonaPresentIntroduced2000Bartholomew and Reno, 2002; Steinbach et al., 2009Initial introduction of parasite in private ponds did not appear to result in establishment. Subsequent detection within the Glen Canyon National Recreation Area in 2007 and 2011
-CaliforniaWidespreadIntroduced1966 Invasive Yasutake and Wolf, 1970
-ColoradoWidespreadIntroduced1987 Invasive Barney et al., 1988; Walker and Nehring, 1995
-ConnecticutPresent1962Introduced1961 Not invasive Hoffman et al., 1962
-IdahoWidespreadIntroduced1987 Invasive Hauck et al., 1988
-MarylandPresent2002Introduced1995Bartholomew and Reno, 2002
-MassachusettsAbsent, intercepted only1966Introduced1966 Not invasive Hoffman, 1990
-MichiganPresentIntroduced1968Hnath, 1970; Yoder, 1972
-MontanaWidespreadIntroduced1994 Invasive Vincent, 1996
-NebraskaPresentIntroduced2001Steinbach et al., 2009
-NevadaWidespread1970Introduced1957Yasutake and Wolf, 1970; Taylor et al., 1973
-New HampshirePresentIntroduced1981 Not invasive Hoffman, 1990
-New JerseyAbsent, formerly presentIntroduced1957Steinbach et al., 2009One hatchery positive in the 1980s is no longer in operation. Parasite does not appear to have become established in the wild
-New MexicoPresentIntroduced1987Hansen et al., 2002
-New YorkWidespreadIntroduced1984Hoffman, 1990
-OhioPresent, few occurrences1970Introduced1968Tidd and Tubb, 1970One private facility reported in 1970; no further information
-OregonLocalisedIntroduced1986Holt et al., 1987
-PennsylvaniaWidespreadIntroduced1956Hoffman et al., 1962First record in USA confirmed in 1958 following an outbreak at a state hatchery in 1956
-UtahWidespreadIntroduced1991 Invasive Wilson, 1991
-VermontPresentIntroduced2002 Not invasive Steinbach et al., 2009
-VirginiaPresentIntroduced1965 Not invasive Hoffman, 1970
-WashingtonLocalisedIntroduced1996Bartholomew and Reno, 2002
-West VirginiaPresent, few occurrencesIntroduced1970sMeyers, 1969No current information; last detected at a small private hatchery in the 1970s
-WyomingPresentIntroduced1988Mitchum, 1995

Europe

AustriaPresent1972Halliday, 1976Based on FAO report, 1972
BelgiumPresent1972Halliday, 1976
BulgariaPresentMargaritov, 1960; Kostova and Chikova, 2011
DenmarkPresentBruhl, 1926
FinlandPresent1932Uspenskaya, 1957Source cites report from 1932 of infections in natural salmon populations
FrancePresentVanco, 1952
GermanyWidespreadNativeHofer, 1903First reported in non-native rainbow trout reared in earthen ponds
HungaryPresentHalliday, 1976Based on FAO report, 1972
IrelandPresentHalliday, 1976Based on FAO report, 1972
ItalyPresent1950 Not invasive Scolari, 1954
LiechtensteinPresentHalliday, 1976Based on FAO report, 1972
LuxembourgPresentHalliday, 1976Based on FAO report, 1972
NetherlandsPresentHalliday, 1976Based on FAO report, 1972
NorwayPresentHastein, 1971
PolandPresentKocylowski, 1953
Russian FederationPresentPresent based on regional distribution.
-Central RussiaPresentBogdanova, 1968
-Northern RussiaPresentUspenskaya, 1955Infections in natural salmon populations
-Russian Far EastPresentBogdanova, 1960Initial report of widespread enzootic focus of infection in the Sakhalin Islands, in cultured and natural populations. However, no subsequent evidence of infection or disease and status unclear
-Southern RussiaPresentUspenskaya, 1957
SlovakiaPresentDyk, 1954
SpainPresentCordero-del-Campillo et al., 1975
SwedenPresentJohansson, 1966
UKPresentElson, 1969; Hoffman, 1970
Yugoslavia (former)PresentTomasec, 1960

Oceania

New ZealandPresentIntroducedHewitt and Little, 1972Although confirmed in 1971, its introduction likely occurred prior to 1952

Pathology

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Whirling disease is characterised by chronic inflammation of the cartilage and other clinical signs, as described in the Disease Course section. Cartilage degradation is generally followed by the formation of lesions. However, in adult fish, parasites are found in isolated pockets in bone and are rarely associated with inflammatory lesions. Parasitic digestion of the cartilage also destroys the structure of the tissue resulting in irregular bone formation and skeletal deformities.

Diagnosis

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Clinical Diagnosis

There is a range of methods available for diagnosing M. cerebralis; most require lethal sampling and that the infection has sufficiently progressed. The suitability of the option depends upon the intended purpose, i.e. research or fish health inspection, and the sample type, e.g. fish, worm, sediment or water. Diagnosis in fish requires the detection and identification of prespore or sporogonic stages. Presumptive diagnosis is based on the presence of clinical disease signs and/or the observation of myxospores isolated from cartilage or bone using pepsin-trypsin digest. Myxospores are broadly oval in frontal view and broadly lenticular in side view, with length 8.7 µm, width 8.2 µm and thickness 6.3 µm (Lom and Hoffman, 1971). Confirmatory diagnosis is based on either histology or molecular-based methods. For histology, parasite stages characteristic of M. cerebralis must be observed in cartilage tissue. Note that mature spores do not stain with hematoxylin and eosin but polar capsules stain darkly with Giemsa or methylene blue. Although several PCR assays that amplify parasite DNA have been developed for the detection of M. cerebralis (Andree et al., 1998; Schisler et al., 2001; Baldwin and Myklebust, 2002; Kelley et al., 2004; Cavendar et al., 2004), only one has been approved for diagnostic confirmation: the nested PCR technique of Andree et al. (1998).

For a more thorough review of the various methods available, see Steinbach et al., (2009) and Hallett and Bartholomew (2012). Detailed procedures for screening and confirmatory tests can be found in the American Fisheries Society - Fish Health Section’s (2012) ‘Suggested procedures for the detection and identification of certain finfish and shellfish pathogens’, under ‘Whirling Disease of Salmonids’ by MacConnell and Bartholomew.

Lesions

Lesions only develop in fish exposed to a sufficiently high parasite dose (Hedrick et al., 1999a). Although they can form in the peripheral nerves and epineurium during migration to the cartilage, they are most prevalent in the cartilage itself (Baldwin et al., 2000). Any cartilage can become infected but the location of lesions varies among salmonid species (e.g. primarily cranial regions in rainbow trout (Baldwin et al., 2000) and lower jaw in Yellowstone cutthroat trout (Murcia et al., 2011)), although progression is similar. Lesions begin as small foci of developmental stages and cartilage degeneration with little associated tissue damage and inflammation (Baldwin et al., 2000; MacConnell and Vincent, 2002), and progress to extensive necrosis of the cartilage. At this time, numerous parasite stages are present - older stages centrally and younger stages at the leading edges (Baldwin et al., 2000). Advanced lesions are associated with mono- and multi-nuclear leukocytes (Baldwin et al., 2000).

Differential Diagnosis

Whirling disease shares clinical signs with other maladies and thus diagnosis cannot be based on their presence alone. Also, myxozoan species cannot readily be distinguished based on developmental stages and the formation of mature myxospores takes several months. Furthermore, spore extraction from fish heads may result in the isolation of multiple myxozoan species. The spore stages of M. cerebralis morphologically and morphometrically resemble other species of the genus Myxobolus Butschli, 1882 which contains over 700 described species (Eiras et al., 2005; Lom and Dyková, 2006). Six other Myxobolus species inhabit the cranial tissues of salmonids (Markiw, 1992; Hogge et al., 2008). M. cerebralis is found in cartilage or bone, while Myxobolus neurobius (Schuberg and Schröder, 1905), Myxobolus kisutchi (Yasutake and Wood, 1957), Myxobolus arcticus (Pugachev and Khokhlov, 1979) and Myxobolus farionis (Gonzalez-Lanza and Alvarez- Pellitero, 1984) have been described in nerve tissue and Myxobolus neurotropus from brain and spinal cord (Hogge et al., 2008). Another commonly encountered myxobolid of salmonids, Myxobolus squamalis, is similar in size to M. cerebralis but has two distinctive ridges on either side of the suture and is found in scale pockets (Hoffman, 1999). Histopathology can resolve fine tissue tropic differences and discriminate between co-occurring cranial myxobolids whose close proximity would lead to co-purification using other methods. DNA-based methods provide unambiguous identification of M. cerebralis (Hogge et al., 2008).

Laboratory Diagnosis

Typically, fish are processed in a laboratory with access to specific pathogen-free water, a range of chemicals and electrical equipment. The handling of some chemicals (e.g. in pepsin trypsin digest and histology) requires special equipment (e.g. fume hood) and disposal. Some steps in the protocols span several hours. Application of most of the diagnostic methods to non-laboratory situations is generally unsafe and impractical. It is more pragmatic to transport samples to the lab than to transport the lab materials and equipment to the sample.

One method that shows promise for on-site detection of the parasite in fish hatcheries and other non-laboratory situations is a simple, rapid DNA detection assay called loop-mediated isothermal amplification (LAMP), but it has not been validated for this use (LAMP; El-Matbouli and Soliman, 2005). Highest sensitivity and specificity perhaps are obtained through molecular methods but it is important to note that detection of a parasite does not indicate disease.

Disease Immunology

Within the fish host, the parasite migrates from the epidermis via the nervous system to the cartilage. Host cellular and/or humoral responses eliminate parasites that do not reach the nerves (Hedrick et al., 1998), but within the nerves, the parasite is protected. Parasites that reach the cartilage trigger an inflammatory response (Hedrick et al. 1998; MacConnell and Vincent, 2002). Whether the immune response occurs within the epithelium, cartilage or nerve ganglia differs among species (Adkison et al., 2002). Only fish that develop lesions have active acquired immunity (MacConnell and Vincent, 2002).

List of Symptoms/Signs

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SignLife StagesType
Finfish / Bursts of abnormal activity - Behavioural Signs Aquatic:Fry Sign
Finfish / Cessation of feeding - Behavioural Signs Aquatic:Fry Sign
Finfish / Corkscrewing - Behavioural Signs Aquatic:Fry Sign
Finfish / Darkened coloration - Skin and Fins Aquatic:Fry Sign
Finfish / Dorso-ventral bends in spine (lordosis) - Body Aquatic:Adult,Aquatic:Fry Sign
Finfish / Generalised lethargy - Behavioural Signs Aquatic:Fry Sign
Finfish / Lateral bends in spine (scoliosis) - Body Aquatic:Adult,Aquatic:Fry Sign
Finfish / Loss of balance - Behavioural Signs Aquatic:Fry Sign
Finfish / Mortalities -Miscellaneous Aquatic:All Stages Sign

Disease Course

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Disease progression in the fish host is temperature dependent and varies somewhat among species; it is best characterised for rainbow trout. In this fish, the parasite starts dividing in the skin 2 hours post-infection and by day 4 and up to day 24 proliferative stages are found in the central nervous system (at 13-15°C; El-Matbouli et al., 1995). The parasite can be found in the cartilage after 20 days (at 16-17°C, or 35 days at 12-13°C) (Halliday, 1973). In their target tissue, parasite trophozoites (developmental stages) consume chondrocytes, which results in lesions and inflammation and other clinical disease signs.

Clinical signs in the fish host may appear 3-8 weeks post-exposure (MacConnell and Vincent, 2002). Signs characteristic of infection with M. cerebralis include:

- whirling behaviour, caused by granulomatous inflammation that constricts the spinal cord and compresses the brain stem (Rose et al., 2000),

- blacktail, the result of infection of the posterior spinal cartilage; associated inflammation presses on root ganglia that control skin melanocytes in the tail (El-Matbouli et al., 1995),

- skeletal deformities, the result of disrupted osteogenesis caused by cartilage damage and inflammation and

- stunted growth and death, directly through physical damage or indirectly through inability to eat or avoid predators (Hedrick et al., 1998; Steinbach et al., 2009).

Development and severity of disease signs depends upon host age, host species/strain, parasite dose and duration of exposure and temperature. Fish may be infected and not display any clinical signs; these hosts are referred to as carriers. Furthermore, these disease signs may be shared with other pathogens (Steinbach et al., 2009).

Epidemiology

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M. cerebralis is a freshwater parasite that alternates between two obligate hosts, metamorphosing into a different spore stage in each (Markiw and Wolf, 1983; Wolf and Markiw, 1984). Triactinomyxon actinospores develop asynchronously in packets of eight in the intestinal epithelium of Tubifex tubifex oligochaetes. When the actinospores are released from the intestine and make contact with the surrounding water, they inflate into anchor-shapes and become waterborne. In proximity to a fish, the actinospore’s polar filaments, each coiled within a polar capsule, fire and, like a harpoon, attach to the fish’s skin. The infectious unit of the actinospore, called the sporoplasm, enters the fish epidermis and migrates through the nervous system to the cartilage, where it continues to multiply and then develops into myxobolid myxospores (development is detailed in El-Matbouli et al., 1995). Although there is some evidence that the myxospores can be released while the fish host is living (Taylor and Haber, 1974; Nehring et al., 2002), it is likely that most myxospores leave the fish once it dies and the cartilage degrades (Hedrick et al., 1998; Hallett and Bartholomew, 2008). When the freed myxospores are ingested by sediment-dwelling oligochaete worms, their polar filaments attach to the worm’s intestine and their sporoplasm penetrates between the epithelium cells where they proliferate and continue the cycle. 

Development in each host takes several months and is temperature dependent, proceeding more quickly at higher temperatures. For instance, development of myxospores can take 11 months at 0-7°C, 90 days at 12-13°C and 52 days at 16-17°C (Halliday, 1973; Hedrick and El-Matbouli, 2002). Only the actinospore stage is infective to fish and the myxospore stage to worms; horizontal or vertical transmission within a fish or worm population does not occur. The myxospore is relatively hardy whereas the actinospore is more fragile (Wagner et al., 2003; Hedrick et al., 2008).

Because M. cerebralis has two waterborne stages, it is readily disseminated with the river current. There must be spatial and temporal overlap of both the invertebrate and vertebrate hosts and spores for the life cycle to continue. Infected fish are the primary means of dispersal. This can occur naturally when fish stray during migration (Engleking, 2002; Zielinski et al., 2010) or through human activities such as commercial stocking or recreational fishing. The parasite may also be spread by piscivorous fish and birds since myxospores can remain viable after passage through the alimentary tract (Taylor and Lott, 1978; Koel et al., 2010).

Impact: Economic

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Historically, the economic impacts of whirling disease have been in relation to the loss of cultured trout. In Europe and the US, both private and publicly owned fish culture operations have sustained large financial losses costs because of M. cerebralis. The parasite has impacted fish culture by causing fish mortalities and reducing fitness, necessitating the destruction of infected fish, requiring disinfection and renovation of facilities, causing the quarantine and closure of facilities and reducing the number of fish available for sale and stocking. In the US, facilities in Utah, California and Colorado were quarantined while millions of dollars were spent to disinfect and renovate them, or they were forced to close when the costs of parasite removal were too great. In 2005, the total value of Utah trout sales dropped almost 30% or approximately $220,000 from the previous year for reasons that included the closure of six privately owned facilities as a result of M. cerebralis detection (House, 2006). In Colorado, the state spent more than $11 million to modernize hatcheries for whirling disease prevention and management between 1987 and 2006 and the federal government completed a multi-million dollar renovation of a National Fish Hatchery to eliminate M. cerebralis (Steinbach et al., 2009). In Europe, although epizootics of whirling disease were widespread in the past century, changes in hatchery practices have greatly minimized losses and the disease is no longer considered a major problem in private fish culture. This transition involved considerable costs but there are no financial reports to support this.

Economic impacts due to the loss of wild fish are often associated with recreational trout fishing. When wild trout population declines were first linked to whirling disease, financial losses due to declines in recreational fishing and tourism were expected. Despite these concerns, no large impact has been documented. In an evaluation of recreational fisheries in Montana and Colorado, no negative effects upon angler satisfaction and local fishing economics could be detected five years after whirling disease caused severe declines in wild trout population (Duffield et al., 1999).

Impact: Environmental

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Impact on Biodiversity

There is evidence of change in the composition of local trout populations in the Madison River, Montana USA, where rainbow trout numbers declined but overall trout populations remained constant because of the increase in brown trout numbers. However, both of these species are introduced in that system. Two native fish in the Rocky Mountain region of the USA, Mountain whitefish (Prosopium williamsoni) and cutthroat trout (Oncorhynchus clarki), may undergo local population declines as a result of infection (Pierce et al. 2011; Koel et al. 2006).

Zoonoses and Food Safety

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M. cerebralis is known only to infect salmonid fishes (salmon and trout) and annelid worms (oligochaetes). Myxospores of Henneguya (H.salminicola and H. zschokkei) and Myxobolus (M.plectroplites and Myxobolus sp.) have been observed in human stool samples but have not been directly linked with abdominal illness (McClelland et al., 1997; Boreham et al., 1998; Lebbad and Willcox, 1998; Moncada et al., 2001). However, humans can develop an allergic reaction following the ingestion of Kudoa-infected fish (Martínez de Velasco et al., 2008) and food poisoning has been associated with the consumption of raw olive flounder (Paralichthys olivaceus) muscle infected with Kudoa septempunctata (Kawai et al., 2012). Although M. cerebralis inhabits fish that may be eaten by humans, the parasite is unlikely to be ingested since it is primarily found in the cartilage and not in the muscle, as is the case for the other myxozoans mentioned.

Disease Treatment

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There is no approved drug or therapeutic treatment for M. cerebralis infection. At least ten candidate drugs have been tested (acetarsone, amprolium, clamoxyquin, fumagillin and its analog TNP-470, furazolidone/furoxone, nicarbazine, oxytetracycle, proguanil and sulfamerazine) (Wagner, 2002). Several of these (furazolidone, proguanil) reduced infection and/or inhibited spore formation; however, none prevented or eliminated infection and some resulted in toxicity (TNP-470) or reduced growth (furazolidone) (Hoffman et al., 1962; Taylor et al., 1973; O’Grodnick and Gustafson, 1974; Alderman, 1986; El-Matbouli and Hoffmann, 1991; Staton et al., 2002). Further development of treatments is hindered by regulatory hurdles, and issues associated with application of treatments to wild fish (Wagner, 2002; Steinbach et al., 2009).

Prevention and Control

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Farm-level Control (movement, housing, sanitation, vaccines)

Improved fish culture practices were developed in the mid 1900’s as a result of the severe effects the parasite had on cultured rainbow trout. Replacement of earthen ponds with concrete raceways and the practice of rearing young fish on parasite-free water sources led to a reduction of clinical disease. Disinfection of ponds and raceways with calcium cyanimide or sodium hypochlorite further reduced the likelihood of the parasite life cycle establishing.

Treatment methods to remove the triactinomyxon stage of the parasite from incoming water include ozone, chlorine, UV light and filtration (Hoffman, 1974; Hedrick et al., 2000; Wagner et al., 2003; Arndt and Wagner, 2003).

Local Control (vaccination, restriction of movement, regulation)

Designing effective control strategies requires understanding of how the parasite is dispersed. Certainly human movements of infected fish are a primary cause of dissemination, and changes in stocking practices and improved diagnostic capabilities have reduced the further spread of the parasite.

National and International Control Policy (vaccination programmes, quarantine regulation)

M. cerebralis is not listed as an injurious pathogen by the World Organisation for Animal Health and there is no international requirement for notification of infection status prior to fish movements. However, some countries (e.g. Australia, Canada, USA, New Zealand) require imports of susceptible fish species to be certified free of the parasite, or limit importation of live fish to life stages that pose a low risk for parasite introduction (i.e. eggs). In the USA, most states require fish health examinations and certification that they are M. cerebralis-free before transport into the state or stocking in state waters.

References

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18/10/2013 Original text by:

Jerri Bartholomew, Dept. of Microbiology, Nash Hall 220, Oregon State University, Corvallis, Oregon 97331, USA

Sascha Hallet, Dept. of Microbiology, Nash Hall 220, Oregon State University, Corvallis, Oregon 97331, USA

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