Invasive Species Compendium

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Myxobolus cerebralis
(whirling disease agent)

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Datasheet

Myxobolus cerebralis (whirling disease agent)

Summary

  • Last modified
  • 14 July 2018
  • Datasheet Type(s)
  • Invasive Species
  • Preferred Scientific Name
  • Myxobolus cerebralis
  • Preferred Common Name
  • whirling disease agent
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Metazoa
  •     Phylum: Cnidaria
  •       Subphylum: Myxozoa
  •         Class: Myxosporea
  • Summary of Invasiveness
  • Myxobolus cerebralis, the myxozoan that causes whirling disease in salmon and trout, was first reported in Germany in the late 1890s. The resistance of European brown trout and the fact that whirling disease wa...

  • Principal Source
  • Draft datasheet under review

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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

  • Myxobolus cerebralis Hofer

Preferred Common Name

  • whirling disease agent

Other Scientific Names

  • Lentospora cerebralis Plehn
  • Myxobolus chondrophagus Hofer
  • Myxosoma cerebralis Kudo
  • Triactinomyxon gyrosalmo Wolf and Markiw

Summary of Invasiveness

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Myxobolus cerebralis, the myxozoan that causes whirling disease in salmon and trout, was first reported in Germany in the late 1890s. The resistance of European brown trout and the fact that whirling disease was not detected outside Europe for over 50 years suggest that it originated in that region. M. cerebralis has primarily been spread by transfers of subclinically infected fish but parasite establishment has been facilitated by the fact that its invertebrate host, Tubifex tubifex, occurs worldwide across a broad range of conditions. Impacts on aquaculture have been significant, but fundamental changes in trout culture and improvements in diagnostics have reduced impacts in hatcheries. Disease in wild populations, with significant impacts on these populations, is reported from the US Rocky Mountain region; the effectiveness of efforts to reduce impacts is difficult to assess. M. cerebralis is not regulated by the World Organisation for Animal Health; however, some countries require imports to be certified free of the parasite and most US states require inspection and certification.

Taxonomic Tree

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  • Domain: Eukaryota
  •     Kingdom: Metazoa
  •         Phylum: Cnidaria
  •             Subphylum: Myxozoa
  •                 Class: Myxosporea
  •                     Class: Myxobolus cerebralis
  •                         Order: Bivalvulida
  •                             Suborder: Platysporina
  •                                 Family: Myxobolidae
  •                                     Genus: Myxobolus
  •                                         Species: Myxobolus cerebralis

Notes on Taxonomy and Nomenclature

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Myxozoan taxonomy is based primarily on the morphology and morphometrics of the myxospore stage and species descriptions follow the guidelines of Lom and Arthur (1989), although contemporary descriptions incorporate DNA sequence information (primarily the ssurRNA gene), which helps to distinguish between phenotypically similar species. Before the first life cycle was elucidated in 1984, actinospores and myxospores were assigned independent binomens in separate classes; the class Actinosporea was suppressed in 1994 by Kent et al. (1994). The current taxonomical scheme is outlined in Lom and Dyková (2006).

Myxobolus cerebralis Höfer 1903 is one of more than 2000 species of the phylum Myxozoa Grasse, 1970 (Lom and Dykova, 2006). It is a member of the predominant class Myxosporea Butschli, 1881 and the most speciose genus, Myxobolus Butschli, 1882 (syn. MyxosomaThelohan, 1892). The name is associated with its myxospore stage, known from fishes. The parasite has undergone several name changes over the decades after its discovery, but its binomen has reverted back to the original (Lom and Noble, 1984). Previous names include: Myxobolus chondrophagus Hofer, 1904; Lentospora cerebralis Plehn, 1905; and Myxosoma cerebralis Kudo, 1933. The actinospore stage in its life cycle was originally assigned an independent name, Triactinomyxon gyrosalmo (Wolf and Markiw, 1984), but has since been synonymised (Kent et al., 1994).

The phylum Myxozoa was initially considered part of the Protozoa but its affinities with Metazoa (multicellularity and phylogenetic position) have been recognised for several decades (Siddall et al., 1995), although its exact relationship has altered between the Bilateria and Cnidaria. The most recent study of new genomic sequences firmly places the Myxozoa within the phylum Cnidaria (Nesnidal et al., 2013); however no-one has proposed how to apply this knowledge taxonomically. The subphylum Myxozoa Grassé, 1970 is also currently unranked due to recent changes. 

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 onlyIntroduced Not invasive Wyk, 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 presentIntroduced1967Steinbach 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 occurrences1970sIntroducedMeyers, 1969No current information; last detected at a small private hatchery in the 1970s
-WyomingPresentIntroduced1988Mitchum, 1995

Europe

AustriaPresent1972Halliday, 1976Based on FAO report, 1972
BelgiumPresent1972Halliday, 1976Based on FAO report, 1972
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, 1955Reports infections 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, 1990
Yugoslavia (former)PresentTomasec, 1960

Oceania

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

History of Introduction and Spread

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M. cerebralis is believed to be indigenous to Europe and Western Russia and to have been introduced elsewhere through movements of subclinically infected fish. In Europe, whirling disease was first detected in Germany in the late 1890s, when rainbow (Oncorhynchus mykiss) and cutthroat (Salvelinus fontinalis) trout, both non-native species, were imported and reared at hatcheries to supplement the culture of native brown trout (Salmo trutta). During the first half of the twentieth century, the parasite spread within Germany and to two Denmark and Finland, but by the 1970s it was reported throughout Europe and the former Soviet Union. It is unclear whether this perceived rapid spread was as a result of the unrestricted transfers of subclinically infected trout that occurred following WWII or if the parasite was already present in the more resistant native brown trout, which then served as a reservoir of infection for the susceptible introduced species. The presence of fish culture facilities on the same rivers where the parasite is detected in natural populations also makes it difficult to determine whether the parasite was introduced or enzootic.

Outside Europe, dissemination was most likely a result of transport of infected fish. It is unknown precisely when introduction of M. cerebralis to North America occurred, but exchanges of live fish, fish eggs, and frozen fish were common between Europe and the US and the parasite was probably introduced unintentionally through the transfer of infected fish or fish products (Hoffman, 1990). The first confirmations of whirling disease occurred nearly simultaneously in Pennsylvania and Nevada in the late 1950s, but suspected infections were present in New York as early as the 1930s. Subsequent spread in the US was largely by interstate movements of subclinically infected fish and through stocking of natural waters. The recent introduction of the parasite into the US is supported by the low level of intraspecific variation between DNA sequences (small subunit ribosomal DNA (ssu rDNA; <1%) and internal transcribed spacer-1 (ITS1; 1.7%)) of European and North American isolates of M. cerebralis (Whipps et al., 2004; Arsan et al., 2007a). For further discussion see Hoffman (1990), Bartholomew and Reno (2002) and Steinbach et al. (2009).

A second highly visible introduction occurred in New Zealand. Clinical disease signs had been reported as early as 1955, but M. cerebralis was not confirmed until 1971. By the time of its detection there was a complete ban on importations of salmonids except in a heat-treated form.

Introductions

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Introduced toIntroduced fromYearReasonIntroduced byEstablished in wild throughReferencesNotes
Natural reproductionContinuous restocking
Nevada Europe  1950s Aquaculture (pathway cause) Yes No Hoffman (1990) Importation of fish resulted in parasite introduction
New Zealand Europe 1950s Aquaculture (pathway cause) Yes No Hewitt and Little (1972) Importation of fish eggs resulted in parasite introduction, likely in the packing material
Pennsylvania Europe 1950s Aquaculture (pathway cause) Yes No Hoffman (1990) Importation of fish resulted in parasite introduction

Risk of Introduction

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The movement of M. cerebralis-infected fish is thought to be the primary vector by which the parasite has spread (Hoffman, 1970, 1990; Hedrick et al., 1998; Bartholomew and Reno, 2002). Salmonid eggs cannot serve as vectors for M. cerebralis as the parasite is not transmitted vertically (O’Grodnick, 1975a), and eggs do not become infected if exposed to triactinomyxons (Markiw, 1991). However, contaminated water or packing material containing eggs could transmit the parasite.

Movement of M. cerebralis-infected fish can occur naturally or through human activities. Legal transfers of infected fish occur primarily as a result of fish stocking activities. Improvements in diagnostic methods have reduced this risk; however, difficulty in timely detection of M. cerebralis infection continues to result in accidental introductions of the parasite. Illegal transfers of M. cerebralis-infected fish are now perhaps the highest-risk human activity spreading the parasite. The construction of ponds on private property has become extremely common, and individuals may stock their ponds by purchase of fish through the private aquaculture industry (Steinbach et al., 2009).

Other pathways for spread of M. cerebralis include movement of water, or sediments containing the parasite, by anglers, boaters, and other recreational enthusiasts. Piscivorous wildlife, including fish, birds and mammals, which ingest M. cerebralis-infected fish, can spread the parasite between drainages. Passage of viable myxospores through the digestive system of piscivorous birds has been demonstrated (Taylor and Lott, 1978; El-Matbouli and Hoffmann, 1991). Other potential means of spread include the release of infected T. tubifex from the aquarium trade (Lowers and Bartholomew, 2003; Hallett et al., 2005, 2006), improper disposal of infected fish parts, use of infected fish parts as bait, and effluent from commercial fish processing (Arsan and Bartholomew, 2008), although these have not been demonstrated.

Pathogen Characteristics

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M. cerebralis has two microscopic phenotypically distinct spore types - a triactinomyxon-type actinospore and a myxobolus-type myxospore; the spore is the infectious stage for the next host. It is presporogonic development of the myxospore stage of M. cerebralis in fishes that can cause whirling disease. A comprehensive description of development in the fish host is provided by El-Matbouli et al. (1995). Stages prior to sporogony resemble those of other myxozoans. Myxospores are bilaterally symmetrical, broadly oval in frontal view, broadly lenticular in side view with length 8.7 µm, width 8.2 µm and thickness 6.3 µm (Lom and Hoffman, 1971). Two hard valve cells surround a binucleate sporoplasm and two polar capsules, which each contain a coiled (5-6 turns), extrudable polar filament. Actinospores are triradially symmetrical and anchor shaped once waterborne (they are folded within the worm host). The actinospore is composed of three valve cells that form an axis (~150 µm) and three caudal processes (each ~194 µm) (El-Matbouli and Hoffmann, 1998). Within the apical end of the axis there are three polar capsules that each contain a coiled polar filament (5 turns). Below the polar capsules is a sporoplasm that contains 64 germ cells.

Myxospore valve cells contain a protective complex polysaccharide matrix (Lom and Hoffman, 1971); the actinospore stage is relatively fragile. Myxospores become nonviable after: freezing at -20°C for 7 days, holding at 20°C for 2 months, drying, treating with alkyl dimethyl benzyl ammonium chloride at 1500 mg/L for 10 minutes, a dose of ultraviolet (UV) of 40-480 mJ/cm² and chlorine bleach at 500 mg/L for 15 minutes (Hedrick et al., 2008).

Host Animals

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Animal nameContextLife stageSystem
HuchoDomesticated host, Subclinical, Wild hostAquatic: FryOpen water systems/Enhancements and culture-based fisheries (inc. ranching and stock enhacement)|Open water systems/Hard substrate, bottom culture|Enclosed systems/Ponds|Enclosed systems/Raceways / running water ponds|Open water systems/Soft substrate/sediment, bottom culture
OncorhynchusDomesticated host, Subclinical, Wild hostAquatic: FryOpen water systems/Enhancements and culture-based fisheries (inc. ranching and stock enhacement)|Open water systems/Hard substrate, bottom culture|Enclosed systems/Ponds|Enclosed systems/Raceways / running water ponds|Open water systems/Soft substrate/sediment, bottom culture
Oncorhynchus mykiss (rainbow trout)
ProsopiumDomesticated host, Subclinical, Wild hostAquatic: FryOpen water systems/Enhancements and culture-based fisheries (inc. ranching and stock enhacement)|Open water systems/Hard substrate, bottom culture|Enclosed systems/Ponds|Enclosed systems/Raceways / running water ponds|Open water systems/Soft substrate/sediment, bottom culture
SalmoDomesticated host, Subclinical, Wild hostAquatic: FryOpen water systems/Enhancements and culture-based fisheries (inc. ranching and stock enhacement)|Open water systems/Hard substrate, bottom culture|Enclosed systems/Ponds|Enclosed systems/Raceways / running water ponds|Open water systems/Soft substrate/sediment, bottom culture
SalvelinusDomesticated host, Subclinical, Wild hostAquatic: FryOpen water systems/Enhancements and culture-based fisheries (inc. ranching and stock enhacement)|Open water systems/Hard substrate, bottom culture|Enclosed systems/Ponds|Enclosed systems/Raceways / running water ponds|Open water systems/Soft substrate/sediment, bottom culture
ThymallusDomesticated host, Subclinical, Wild hostAquatic: FryOpen water systems/Enhancements and culture-based fisheries (inc. ranching and stock enhacement)|Open water systems/Hard substrate, bottom culture|Enclosed systems/Ponds|Enclosed systems/Raceways / running water ponds|Open water systems/Soft substrate/sediment, bottom culture
Tubifex tubifex

Climate

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ClimateStatusDescriptionRemark
C - Temperate/Mesothermal climate Preferred Average temp. of coldest month > 0°C and < 18°C, mean warmest month > 10°C
Cf - Warm temperate climate, wet all year Preferred Warm average temp. > 10°C, Cold average temp. > 0°C, wet all year
Cs - Warm temperate climate with dry summer Tolerated Warm average temp. > 10°C, Cold average temp. > 0°C, dry summers
Cw - Warm temperate climate with dry winter Tolerated Warm temperate climate with dry winter (Warm average temp. > 10°C, Cold average temp. > 0°C, dry winters)
D - Continental/Microthermal climate Tolerated Continental/Microthermal climate (Average temp. of coldest month < 0°C, mean warmest month > 10°C)
Df - Continental climate, wet all year Tolerated Continental climate, wet all year (Warm average temp. > 10°C, coldest month < 0°C, wet all year)
Ds - Continental climate with dry summer Tolerated Continental climate with dry summer (Warm average temp. > 10°C, coldest month < 0°C, dry summers)
Dw - Continental climate with dry winter Tolerated Continental climate with dry winter (Warm average temp. > 10°C, coldest month < 0°C, dry winters)

Latitude/Altitude Ranges

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Latitude North (°N)Latitude South (°S)Altitude Lower (m)Altitude Upper (m)
66 35

Notes on Natural Enemies

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No specific natural enemies for M. cerebralis have been identified. However, it is assumed that once the integrity of the spore has been compromised, the organism is vulnerable to degradation by other microbiota. Non-target hosts aid in the reduction of infectious stages. For example, actinospores will attach to non-salmonids (Kallert et al., 2009) and thus be deactivated. Similarly, resistant lineages of T. tubifex will consume and deactivate myxospores, thus preventing infectious stages reaching susceptible lineages (Beauchamp et al., 2006).

Means of Movement and Dispersal

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Natural Dispersal

Both spore stages are disseminated naturally in water (Steinbach et al., 2009).

Vector Transmission

The parasite can be disseminated both locally and over long distances through the movement of infected salmonid fish (Hoffman, 1990). Natural fish migrations may move the parasite locally or stocking of subclinically infected fish may result in long distance and local dissemination of the parasite. Piscivorous birds and wildlife may also disseminate myxospores but the likelihood of this occurrence is highest over short distances (Koel et al., 2010). The release of infected T. tubifex from the aquarium trade (Lowers and Bartholomew, 2003; Hallett et al., 2005, 2006) could also result in release of the parasite.

 

Accidental Introduction

Myxospores in sediment can be accidentally transported on anglers' waders (Gates et al., 2008). Other means of spread could include the improper disposal of infected fish parts as bait and the discharge of effluent from commercial fish processing (Arsan and Bartholomew, 2008), although these have not been demonstrated.

Pathway Causes

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CauseNotesLong DistanceLocalReferences
Aquaculture Yes Yes Bartholomew and Reno, 2002
Fisheries Yes Yes Bartholomew and Reno, 2002
HitchhikerPotential movement on anglers' gear Yes Gates et al., 2008
Interbasin transfersStocking of subclincally infected fish Yes Yes Bartholomew and Reno, 2002
Interconnected waterwaysNatural fish migration Yes Bartholomew and Reno, 2002
StockingStocking of subclinically infected fish Yes Yes Bartholomew and Reno, 2002

Pathway Vectors

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VectorNotesLong DistanceLocalReferences
Aquaculture stockMost frequent means of dissemination. Parasite not easily detected Yes Yes Hoffman, 1990
Clothing, footwear and possessionsMyxospores in sediment can be moved on anglers' waders Yes Gates et al., 2008
Host and vector organismsDissemination mainly through movement of salmonoid fish; possibly by wildlife over shorter distances Yes Yes Koel et al., 2010
WaterBoth spore stages are disseminated naturally in water Yes Steinbach et al., 2009

Economic Impact

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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).

Environmental Impact

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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).

Threatened Species

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Threatened SpeciesConservation StatusWhere ThreatenedMechanismReferencesNotes
Oncorhynchus clarki clarki (cutthroat trout)National list(s) National list(s)PathogenicNehring et al., 1998; Vincent, 1996

Risk and Impact Factors

Top of page Invasiveness
  • Proved invasive outside its native range
  • Abundant in its native range
  • Has propagules that can remain viable for more than one year
  • Reproduces asexually
Impact outcomes
  • Changed gene pool/ selective loss of genotypes
  • Host damage
  • Negatively impacts aquaculture/fisheries
  • Reduced native biodiversity
  • Threat to/ loss of endangered species
  • Threat to/ loss of native species
  • Negatively impacts trade/international relations
Impact mechanisms
  • Parasitism (incl. parasitoid)
  • Pathogenic
Likelihood of entry/control
  • Difficult to identify/detect as a commodity contaminant
  • Difficult to identify/detect in the field
  • Difficult/costly to control

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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