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Bothriocephalus acheilognathi infection

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Datasheet

Bothriocephalus acheilognathi infection

Summary

  • Last modified
  • 22 November 2019
  • Datasheet Type(s)
  • Animal Disease
  • Preferred Scientific Name
  • Bothriocephalus acheilognathi infection
  • Overview
  • Bothriocephalus acheilognathi is a tapeworm that can infect a wide variety of freshwater fish but is primarily reported from cultured and wild carp. It is native to East Asia and over the last few decades has b...

  • Principal Source
  • Draft datasheet under review

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Pictures

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PictureTitleCaptionCopyright
Bothriocephalus acheilognathi (Asian fish tapeworm); SEM - lateral view of the scolex. From humpback chub (Gila cypha), Little Colorado River, Grand Canyon, Colorado, USA.
TitleScolex
CaptionBothriocephalus acheilognathi (Asian fish tapeworm); SEM - lateral view of the scolex. From humpback chub (Gila cypha), Little Colorado River, Grand Canyon, Colorado, USA.
Copyright©Anindo Choudhury
Bothriocephalus acheilognathi (Asian fish tapeworm); SEM - lateral view of the scolex. From humpback chub (Gila cypha), Little Colorado River, Grand Canyon, Colorado, USA.
ScolexBothriocephalus acheilognathi (Asian fish tapeworm); SEM - lateral view of the scolex. From humpback chub (Gila cypha), Little Colorado River, Grand Canyon, Colorado, USA.©Anindo Choudhury
Bothriocephalus acheilognathi (Asian fish tapeworm); SEM - dorso-ventral view of the scolex. From non-native common carp (Cyprinus carpio). Litte Colorado River, Grand Canyon, Colorado, USA.
TitleScolex
CaptionBothriocephalus acheilognathi (Asian fish tapeworm); SEM - dorso-ventral view of the scolex. From non-native common carp (Cyprinus carpio). Litte Colorado River, Grand Canyon, Colorado, USA.
Copyright©Anindo Choudhury
Bothriocephalus acheilognathi (Asian fish tapeworm); SEM - dorso-ventral view of the scolex. From non-native common carp (Cyprinus carpio). Litte Colorado River, Grand Canyon, Colorado, USA.
ScolexBothriocephalus acheilognathi (Asian fish tapeworm); SEM - dorso-ventral view of the scolex. From non-native common carp (Cyprinus carpio). Litte Colorado River, Grand Canyon, Colorado, USA.©Anindo Choudhury

Identity

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Preferred Scientific Name

  • Bothriocephalus acheilognathi infection

Other Scientific Names

  • Schyzocotyle acheilognathi infection

Overview

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Bothriocephalus acheilognathi is a tapeworm that can infect a wide variety of freshwater fish but is primarily reported from cultured and wild carp. It is native to East Asia and over the last few decades has been spread widely throughout the world (to all continents except Antarctica) via human activities including the movement of (mostly cyprinid) fish for aquaculture, the pet fish trade, aquatic weed control and mosquito control, and more recently in movement of bait fish. In addition, birds which eat infected fish may transport its eggs and spread them through defecation. Moderate to heavy infections can be fatal for fish fingerlings, or adults of small species, and infection with B. acheilognathi can be a problem for aquaculture (both through loss of fish and through disruption to operations and the costs of control); it is also suspected of adversely affecting a number of endangered wild fish species.

Hosts/Species Affected

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B. acheilognathi has been reported in over 300 species of fish (Dove et al., 1997; Dove and Fletcher, 2000; Salgado-Maldonado and Pineda-López, 2003; Rojas-Sánchez and García-Prieto, 2008; Bean and Bonner, 2009; Scholz et al., 2012; Kuchta et al., 2018), of which only a selection are recorded in the Hosts table. All freshwater fishes must be considered susceptible, unless shown otherwise. It prefers cyprinid fishes (barbs, carps, minnows, shiners, chub etc.); judging from records, grass carp Ctenopharyngodon idella and common carp Cyprinus carpio are two of its major hosts, although most of the records from common carp appear to be in fish farms (fingerlings of these species are the most severely affected stage, although adults can also be infected). However, it will infect atheriniform species (silversides) and cyprinodontiform species (topminnows, mollies, guppies, and allies). Some cichlids and eleotrids appear to be susceptible in the tropics. Planktivorous fish are particularly susceptible. Perches (Percidae), bass (Centrarchidae), pikes (Esocidae), temperate and tropical catfishes (Siluriformes) and suckers (Catostomidae) seem to be infected only rarely (Hoffman, 1999; Dove and Fletcher, 2000; Rojas-Sanchez and Garcia-Prieto, 2008; Choudhury and Cole, 2012; Scholz et al., 2012).

Distribution

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B. acheilognathi is native to eastern Asia but has been spread widely throughout the world (including all continents except Antarctica) by human activities (Bauer and Hoffman, 1976). It seems to be widely distributed in China (Nie et al., 2000) but its status in Japan appears uncertain (Choudhury and Cole, 2012). The records of Bothriocephalus spp. from native cyprinids in Africa and India are considered to be B. acheilognathi (Pool, 1987; Kuchta and Scholz, 1997; Kuchta et al., 2012) but this needs to be confirmed by molecular analyses that include tapeworms from native barbs in the interior of Africa and from native cyprinids in streams of the Himalayan foothills (Malhotra, 1984). This is particularly important since there is some indication that the Asian fish tapeworm may be a species complex (Liao, 2007; Luo et al., 2003; Choudhury and Cole, 2012). Reports from clariid catfishes in Africa need to be verified because Kuchta et al. (2012) stated that the species could be confused with another similar tapeworm, Tetracampos ciliotheca. Molecular data indicate that isolates from continental North America, Hawaii, and Central America closely match B. acheilognathi isolates from Eurasia (Bean et al., 2007; Choudhury et al., 2013; Salgado-Maldonado et al. 2015; A. Choudhury, St Norbert College, De Pere, Wisconsin, USA, unpublished data).  

In the U.S., the parasite seems to be particularly established in the western and southwestern parts of the country (Heckmann, 2000; Kuperman et al., 2002; Warburton et al., 2002; Choudhury et al., 2006; Archdeacon et al., 2009; Kline et al., 2009). In Mexico, it appears to be widely distributed (Salgado-Maldonado and Pineda-López, 2003; Rojas-Sánchez and García-Prieto, 2008). In Australia, it is established in the eastern part of the continent (Dove and Fletcher, 2000). In Europe, it is absent from northern European (Scandinavian) countries; the current status in many central and eastern European countries seems unclear. In South Africa, it is common in carp in certain areas and remains well established.

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.

Last updated: 10 Jan 2020
Continent/Country/Region Distribution Last Reported Origin First Reported Invasive Reference Notes

Africa

AlgeriaPresent, LocalizedIntroducedMeddour (1988)Lake Oubeira
Congo, Democratic Republic of theAbsent, Unconfirmed presence record(s)Baer and Fain (1958)Lake Kivu; record is of B. kivuensis, synonymized with B. acheilognathi, but synonymy is questionable (see Choudhury and Cole, 2012)
EgyptAbsent, Unconfirmed presence record(s)Rysavy and Moravec (1975)Record is of B. aegyptiacus, synonymized with B. acheilognathi, but synonymy should be validated.
EthiopiaPresentKuchta et al. (2012); Zekarias and Yimer (2007)
MauritiusPresentIntroducedPaperna (1996)Paperna reported a personal communication from J.G. Van As, University of the Free State, Bloemfontein, South Africa.
NigeriaPresentIntroducedEyo and Iyaji (2014)Niger and Benue River confluence, Lokoja
RwandaAbsent, Unconfirmed presence record(s)Baer and Fain (1958)Lake Kivu, as B. kivuensis, synonymized with B. acheilognathi, but synonymy requires validation (see Choudhury and Cole, 2012)
South AfricaPresentIntroducedInvasiveBoomker et al. (1980)

Asia

AfghanistanPresentIntroducedMoravec and Amin (1978)
AzerbaijanPresentIntroducedDubinina (1987)
ChinaPresentCABI (Undated a)Present based on regional distribution.
-ChongqingPresentNativeLuo et al. (2003)
-FujianPresentNativeLiao XiangHua (2007)In grass carp, but not in common carp
-GansuPresentNativeLiao XiangHua (2007)In grass carp, but not in common carp
-GuangdongPresentNativeLiao and Shih (1956)Originally described by Yeh, 1955, as B. gowkongensis
-GuizhouPresentNativeLiao XiangHua (2007)In common carp and grass carp
-HeilongjiangPresentNativeLiao XiangHua (2007)In common carp and grass carp
-HubeiPresentNativeNie et al. (2000); Luo et al. (2003)Lakes in the flood plains of the Yangtze River, in cultrinin cyprinids
-Inner MongoliaPresentNativeLiao XiangHua (2007)Dong Lake
-JiangxiPresentNativeXi et al. (2011)
-JilinPresentNativeLiao XiangHua (2007)
-LiaoningPresentNativeLiao XiangHua (2007)
-NingxiaPresentNativeLiao XiangHua (2007)In common carp, but not in grass carp
-ShanxiPresentNativeLiao XiangHua (2007)
-SichuanPresentNativeLiao XiangHua (2007)
-XinjiangPresentIntroducedLiao XiangHua (2007)In common carp, but not in grass carp
-YunnanPresentIntroducedLiao XiangHua (2007)
GeorgiaPresentIntroducedKurashvili (1990)In closed reservoirs
India
-Jammu and KashmirAbsent, Unconfirmed presence record(s)Akhter et al. (2008)Reported from hill streams in a native cyprinid. Species identification should be validated
-MeghalayaPresent2016Sunila Thapa and Das (2016)Found in aquacultured grass carp (Ctenopharyngodon idella)
-Uttar PradeshPresentIntroducedMalhotra (1984); Anshu Chaudhary et al. (2015)Reported in 1984 as B. teleostei -- that identification needs to be validated. In 2015 reported again, and identified by molecular methods, from introduced Xiphophorus hellerii.
IranPresentIntroducedInvasiveMokhayer (1976)
IraqPresentIntroducedInvasiveKhalifa (1986)
IsraelPresentIntroducedPaperna (1996)
JapanPresentYamaguti (1934); Choudhury and Cole (2012)See Choudhury & Cole (2012) for a discussion of its status as a native or introduced species
KazakhstanPresentIntroducedInvasiveDubinina (1987)
KyrgyzstanPresentIntroducedDubinina (1987)
MalaysiaPresentIntroducedFernando and Furtado (1964)
MongoliaAbsent, Unconfirmed presence record(s)Dubinina (1987)Dubinina does not specifically mention Mongolia, only the Amur River drainage
PhilippinesPresentIntroducedVelasquez (1982)
South KoreaPresentIntroducedKim et al. (1986)Kim et al. (1986) cited in Paperna (1996)
Sri LankaPresentIntroducedFernando and Furtado (1963)
TajikistanPresentIntroducedDubinina (1987)
TurkeyPresentIntroducedAydogdu et al. (2003)
TurkmenistanPresentIntroducedDubinina (1987)
UzbekistanPresentIntroducedOsmanov (1971)

Europe

AlbaniaPresentStojanovski et al. (2013)Lake Ohrid
AustriaPresentIntroducedOtte et al. (1972)Carp ponds
BelarusPresentDubinina (1987)
Bosnia and HerzegovinaPresentIntroducedKiskaroly (1977)Carp ponds
CroatiaPresentIntroducedKezic et al. (1975)Kezic et al. (1975) cited in Hoffman (1999)
CzechoslovakiaPresentFaina and Par (1977)
FrancePresentIntroducedDenis et al. (1983)
GermanyPresentIntroducedInvasiveKorting (1974)
HungaryPresentMolnár (1968); Buza et al. (1970)Described as B. phoxini, in Phoxinus phoxinus
ItalyPresentIntroducedInvasiveMinervini et al. (1985)
LatviaPresentIntroducedVismanis and Jurkane (1967)Carp ponds. Vismanis and Jurkane (1967) cited in Kirjušina and Vismanis (2007)
North MacedoniaPresentIntroducedStojanovski et al. (2013)Lake Ohrid
PolandPresentIntroducedPanczyk and Zelazny (1974)
RomaniaPresentIntroducedRadulescu and Georgescu (1962)
RussiaPresentCABI (Undated a)Present based on regional distribution.
-Central RussiaPresentIntroducedInvasiveDubinina (1987)Mainly in aquaculture facilities
-Eastern SiberiaPresentNativeDubinina (1987)Amur River drainage
-Russian Far EastPresentNativeDubinina (1987)Amur River drainage
-Southern RussiaPresentIntroducedInvasiveDubinina (1987)Mainly in aquaculture facilities
-Western SiberiaPresentIntroducedInvasiveDubinina (1987)Mainly in aquaculture facilities
UkrainePresentIntroducedMALEVITSKAYA (1958)
United KingdomPresentIntroducedAndrews et al. (1981)

North America

CanadaPresentCABI (Undated a)Present based on regional distribution.
-British ColumbiaAbsent, Invalid presence record(s)Choudhury et al. (2006); Arai and Mudry (1983)Specimens reported by Arai and Mudry (1983) were misidentified
-ManitobaPresentIntroducedChoudhury et al. (2006)Lake Winnipeg below Lockport Dam, not likely upstream of Pine Falls on Winnipeg River or Grand Rapids on Saskatchewan River (Patrick Nelson, North South Consultants Inc., Winnipeg, Manitoba, Canada, personal communication)
-OntarioPresent, LocalizedIntroducedMarcogliese (2008)Detroit River, hence likely present in Lakes Huron, St. Clair and Erie. Muzzall et al. (2016) confirm its presence on the Michigan side of Lakes Huron and St. Clair.
HondurasPresentIntroducedSalgado-Maldonado et al. (2015)
MexicoPresent, WidespreadIntroducedInvasiveLópez-Jiménez (1981)Widespread
PanamaPresentIntroducedChoudhury et al. (2013)
Puerto RicoPresentIntroducedBunkley-Williams and Williams (1994)
United StatesPresentCABI (Undated a)Present based on regional distribution.
-ArizonaPresent, WidespreadIntroducedInvasiveClarkson et al. (1997)
-ArkansasPresentIntroducedScott and Grizzle (1979)In a hatchery
-CaliforniaPresentIntroduced1980Hoffman (1999); Warburton et al. (2002)Southern California
-ColoradoPresentIntroducedWard (2005)
-FloridaPresentIntroducedRogers (1976); Scott and Grizzle (1979)
-HawaiiPresentIntroducedFont and Tate (1994)
-IndianaPresentIntroducedBuckner et al. (1985)
-KansasPresentIntroducedPullen et al. (2009)
-KentuckyPresentIntroducedChoudhury et al. (2006)
-LouisianaPresentIntroducedCABI (Undated)In mosquitofish, Gambusia; Original citation: W. Font, Southeastern Louisiana University, Hammond, Louisiana, USA, personal communication, 2015
-MichiganPresentIntroducedMarcogliese (2008); Muzzall et al. (2016); Boonthai et al. (2017)In wild in Detroit River, Lake Huron and Lake St. Clair; likely present in Lake Erie. Widespread in mostly wild-caught baitfish in retail stores.
-NebraskaPresentIntroducedChoudhury et al. (2006)
-NevadaPresentIntroduced1987Heckmann et al. (1993)Muddy River, Virgin River, bait shops
-New HampshireAbsent, Unconfirmed presence record(s)Hoffman (1999)
-New MexicoAbsent, Unconfirmed presence record(s)Bean et al. (2007)Pecos River
-North CarolinaPresentIntroducedInvasiveRiggs and Esch (1987)In a reservoir
-TexasPresentIntroducedInvasiveBean et al. (2007)Rio Grande/Río Bravo del Norte, Pecos River
-UtahPresentIntroducedHeckmann et al. (1987)Virgin River
-WisconsinPresentIntroducedChoudhury et al. (2006)Land-o-Lakes lakes

Oceania

AustraliaPresentIntroducedInvasiveDove et al. (1997)
-New South WalesPresentIntroducedDove et al. (1997); Dove and Fletcher (2000)
-VictoriaPresentIntroducedDove and Fletcher (2000)
New ZealandAbsent, Intercepted onlyEdwards and Hine (1974)Arrived with grass carp imports from Hong Kong but intercepted during quarantine

South America

ArgentinaPresent, LocalizedIntroducedWaicheim et al. (2014)
BrazilPresent, LocalizedIntroducedRego et al. (1999)

Pathology

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The pathology of B. acheilognathi infection has been summarized by Choudhury and Cole (2012) and by Scholz et al. (2012). “It includes inflammation, haemorrhaging, destruction and dysfunction of the intestinal mucosa, necrosis and even perforation (Scott and Grizzle, 1979; Hoole and Nisan, 1994; Schäperclaus, 1991; Sinha and Mehrotra, 1991; Heckmann, 2000). At the cellular level, separation and shedding of microvilli occurred at the interface between the gut and the tapeworm’s bothridia (Hoole and Nisan, 1994). Inflammation is accompanied by migration and infiltration of lymphocytes, macrophages and eosinophils to the infected area and even out of the gut to the parasite surface (Hoole and Nisan, 1994).” (Choudhury and Cole, 2012).

Diagnosis

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There are no certain or exclusive external signs of bothriocephalosis unless worms protrude from the mouth (Han et al., 2010) or from the anus. In most cases, the disease is first suspected from the abdominal distension shown by smaller fish. Diagnosis is by examining faeces and finding eggs and segments, or by examining the gut of the fish. The gastrointestinal tract is either dissected open in a petri dish, or squashed between two glass plates if the gut is relatively thin walled.  Either way, the preparation is best examined with a stereo dissecting scope (Choudhury et al., 2004; Scholz et al., 2012). In small fish, the gut can be squashed between two slides and examined with a compound microscope to search for parasites (C. Banner, Oregon State University, Corvallis, Oregon, USA, personal communication, 2015). Dissections are preferred in larger fish because the variable thickness of various regions of the gut, especially the thicker stomach wall, as well the presence of mucus, may interfere with the examinations. The fleshy heart-shaped scolex (when viewed laterally) – unique among Bothriocephalus spp. - along with rounded edges of the segments, and the medial position of the genital pores, allow a tentative first identification. Extracted worms must be fixed in a steaming hot fixative such as buffered 10% hot formalin (3.8% - 4% formaldehyde solution), except for a small piece – not the scolex - that should be stored in 95% or 100% molecular grade ethanol for molecular diagnoses.  Fixed worms are stained and processed by standard methods, mounted permanently on slides and examined and identified using a compound microscope and available descriptions (Scholz, 1997). This identification can be supplemented by molecular data (Bean et al., 2007; Choudhury et al., 2013; Salgado-Maldonado et al., 2015). Choudhury and Cole (2012) discuss diagnosis in North America, while Kuchta et al. (2012) provide insight into issues in Africa.

List of Symptoms/Signs

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SignLife StagesType
Finfish / Change in body shape - Body Aquatic:Fry Sign
Finfish / Change in feed-conversion ratio - Miscellaneous Sign
Finfish / Change in length-weight ratio - Body Aquatic:Fry Sign
Finfish / 'Dropsy' - distended abdomen, 'pot belly' appearance - Body Aquatic:Adult,Aquatic:Fry Sign
Finfish / Fish swimming near surface - Behavioural Signs Aquatic:Fry Sign
Finfish / Intestines swelling / oedema - Organs Sign
Finfish / Intestines white-grey patches (haemorrhage / necrosis / tissue damage) - Organs Sign
Finfish / Mortalities -Miscellaneous Aquatic:Adult,Aquatic:Fry Sign

Disease Course

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The disease caused by B. acheilognathi is called ‘bothriocephalosis’ and has been reviewed by Hoffman and Schubert (1984), Schäperclaus (1991), Paperna (1991, 1996), Hansen et al. (2006), Borucinska (2008), Choudhury and Cole (2012), and Scholz et al. (2012).  The parasite causes a decrease in weight of carp (Bauer et al., 1969; Britton et al., 2011); mortality of young fish caused by moderate to heavy infections has been documented in cultured grass carp (Ctenopharyngodon idella), common carp (Cyprinus carpio) and koi (Yeh, 1955; Bauer et al., 1969; Han et al., 2010), as well as in golden shiners (Notemigonus chrysoleucas) and mosquitofish (Gambusia) (Hoffman, 1980; Granath and Esch, 1983b).

Several authors have reviewed the common lesions seen in infections in various species of fish (Scott and Grizzle, 1979; Granath and Esch 1983a; Hoole and Nissan, 1994; Hansen et al., 2006, Han et al., 2010). Stunted growth in infected fish will limit mouth gape size which increases the time the fish relies on zooplankton as prey, thus extending the window of exposure to the infectious larvae (Hansen et al., 2006). As the worms grow, they cause mechanical obstruction and distension of the gut, especially in smaller fish (those which are small when mature, and the young of larger species) in which just a few worms can cause a significant problem. This is accompanied by gut inflammation which can have severe effects. Pathogenic changes are mainly due to the interaction between the worms’ scoleces (attachment organs) and the gut mucosa at the attachment site, but the gut mucosa can also be altered by the rest of the worm’s body (its strobila) in heavy infections, resulting in pressure necrosis and rupture.  If left untreated, heavily infected fish suffer distress, which is manifested in altered behaviour and physiological complications that may affect other organs such as the liver. The condition worsens as the disease progresses, and eventually emaciation and pathology lead to death. On the other hand, light chronic infections may not have such a detrimental effect.

Hoffman (1999) suggests that in an aquaculture setting the higher susceptibility of smaller fish to heavy infections may decrease; however there are no data for this decrease in susceptibility in wild populations. Britton et al. (2011) showed that infected fish in experimental settings have a suppressed feeding rate when compared with paired controls of uninfected fish. In wild populations of fish, B. acheilognathi has been implicated in poor body condition (Hoffnagle et al., 2006).

Epidemiology

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B. acheilognathi typically has two hosts in its life cycle. As an adult, it lives in the gut of its fish host where it reproduces sexually and releases its eggs.  The eggs pass out with the faeces into the water, where a ciliated larva, the coracidium, develops and eventually hatches out.  Development is temperature dependent; at 28-30°C, larvae develop and hatch after a day, whereas at 14-15°C, it takes 10-28 days for development and hatching (Hoffman, 1980; Granath and Esch, 1983c; Paperna, 1996). Eggs do not develop below 12°C (Paperna, 1996). The hatched coracidium larva swims around, using its stored energy reserves, until it is eaten by a copepod (Marcogliese and Esch, 1989b; Paperna, 1996).  Numerous species of cyclopoid copepods are suitable intermediate hosts (see Vectors and Intermediate Hosts table in the Bothriocephalus acheilognathi datasheet)). Coracidia survive longer (5-6 days) at lower temperatures (15-16°C -- Hanzelova and Zitnan, 1986), but may become sluggish (Granath and Esch, 1983c). Once consumed by a copepod, the larva burrows through the gut into the body cavity where it develops into a larval stage called a procercoid. Copepods may have several procercoids at the same time. In a week or so at 20°C, the procercoid develops a tiny rounded tail called a ‘cercomer’ and is now ready to infect a fish host. Small fish are more likely to become infected because of the higher proportion of zooplankton, and therefore of infected copepods, in their diet. When susceptible fish hosts ingest infected copepods, the procercoid larvae emerge from the copepod body and occupy the folds of the gut mucosa where they begin developing into typical, segmented, ‘strobilate’ tapeworms. In north-temperate regions, recruitment of larval worms into fish typically occurs in the autumn (fall) and worms produce eggs by the following spring. This seasonality may not be pronounced in the tropics. Strobilation – producing segments - is also temperature dependent. In a study in common carp (Cyprinus carpio), 89% of the worms became segmented in 16-20 days at 25°C, while 80% remained unsegmented 4 months later at 15°C (Oskinis, 1994). In barbs (also members of the Cyprinidae), maturation takes 1.5 -2 months at 15-22°C (Davydov 1978), while temperatures below 15°C hinder development (6-8 months). In grass carp (Ctenopharyngodon idella), worms mature in about three weeks at 28-29°C (Liao and Shih, 1956). Granath and Esch (1983c) found that worms in mosquitofish (Gambusia affinis) did not mature at 20°C. Such differences may be due to parasite strain/population differences along with differences in host physiology.

Hansen et al. (2007) showed, using different size classes of bonytail chub (Gila elegans) in experimental infections, that larger, suitable, predatory fish hosts can acquire and accumulate infections by consuming smaller infected fish.

Impact: Economic

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B. acheilognathi was first reported in Asia and Eastern Europe as an important disease agent (Bauer et al., 1969; Heckmann, 2009).  Moderate to heavy B. acheilognathi infections can be fatal to fish fingerlings. This has been documented in cultured grass carp (Ctenopharyngodon idella), common carp (Cyprinus carpio) and koi (Yeh, 1955; Bauer et al. 1969; Han et al., 2010); Liao and Shih (1956) reported severe losses in grass carp of the young-of-the-year age class in China, with a mortality rate of 90% in winter months (Liao and Shih, 1956). Costs to aquaculture, beyond loss of fish, are due to disruption of normal hatchery and fish farming operations, increased cost of operation as a result of treating fish (with medication administered in diet), and draining and disinfecting tanks and ponds. Aquaculture operations have to invest in additional quarantine and inspection infrastructure and personnel.

Impact: Environmental

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Impact on Biodiversity

The impact of B. acheilognathi on biodiversity is still largely unknown, and assessing the population-level impact is difficult, but it is suspected of impacting populations of two IUCN-listed endangered North American fish species, the woundfin (Plagopterus argentissimus) and the humpback chub (Gila cypha), and potentially threatens others such as the U.S. federally listed endangered Tui Mohave chub Siphateles bicolor mohavensis (Heckmann, 2000, 2009; Hoffnagle et al., 2006; Choudhury and Cole, 2012; Archdeacon et al., 2008; Stone et al., 2007).  Also in the USA, it hinders growth of the Near Threatened roundtail chub Gila robusta (Brouder, 1999) and the US-listed Topeka shiner Notropis topeka (Koehle and Adelman, 2007); experimental infections in the Critically Endangered bonytail chub (Gila elegans) resulted in reduced growth, decline in health condition indices, and accelerated mortality when food was reduced (Hansen et al., 2006).  Velázquez-Velázquez et al. (2011) found a high prevalence of infection in the Endangered Chiapas killifish (Profundulus hildebrandi) in Mexico.

Zoonoses and Food Safety

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Yera et al. (2013) report eggs of B. acheilognathi in the faeces of a patient presenting with stomach pains.  The eggs were identified by molecular sequencing and the infection was reported as accidental and was the result of the tapeworm passing through the intestine after an infected fish was consumed. Apart from that, this parasite has never been described from mammals.

Disease Treatment

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Various herbal extracts, such as from Cucurbita (gourds) and Areca (areca nut), were successfully used to treat Asian fish tapeworm infections before the advent of modern anthelminthics (Liao and Shih, 1956; Nie and Pan, 1985). Scholz et al. (2012) also mention horseradish (Armoracia rusticana) leaves and conifer needles.

Such herbal extracts were replaced by niclosamide-based drugs (see Dick and Choudhury, 1995, and table), which were quite successful. Molnár (1970) reported that niclosamide was effective in clearing Asian tapeworm infections in aquaria and experimental ponds when added to feed at doses between 0.1 and 20 g/kg body weight of fish. Par et al (1977) successfully used a dose of 1 g/kg body weight of fish. Zitnan et al. (1981) used ‘Taenifugin Carp’, a granulated medicated feed containing 0.7% niclosamide salt, and reported that administering this feed at 2% of body weight of fish (=0.14 g niclosamide/kg body weight of fish) was effective in clearing tapeworm infections. However, niclosamide was reported to be highly toxic to fish when administered in water (Molnár, 1970).

Praziquantel, which is administered either by water bath or in food, seems to be the current drug of choice. Time and dosage are important; Ward (2007) was able to clear infections in bonytail chub (Gila elegans) exposed to a dose of 1.5 mg/l for 24 hours. Mitchell and Darwish (2009) found that fish density during treatment also affected efficacy; grass carp (Ctenopharyngodon idella) immersed for 24 hrs at a density of 60 g/l, and 0.75 mg/l of praziquantel or higher, were cleared of the tapeworm. Their study showed that dosage must be adjusted to fish densities for effective treatment. Drug delivery must also take into account potential harm to fish at higher dosages (Mitchell and Hobbs, 2007).

Prevention and Control

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Immunization and Vaccines

There is currently no effective fish vaccine against B. acheilognathi.

Management Strategies in Aquaculture

(reviewed by Choudhury and Cole, 2012)

Infected fish should not be allowed into aquaculture facilities. Imported fish and new shipments should be quarantined and examined carefully for B. acheilognathi and other parasites and pathogens. ‘Disease free’ certification should include this species. Aquaculture operations may wish to deworm fish before introducing them to their aquaculture facilities. However, praziquantel-treated fish cannot be sold as food in the USA. Praziquantel will not kill the eggs or free coracidia swimming larvae (Kline et al., 2009). Copepod densities can be controlled in small indoor facilities, but probably not in outdoor ponds. Stocking of fish and movement between hatcheries should be done carefully. Bauer et al. (1969) noted success in treating the pond sediment after fish have been harvested.  Drying the pond in the spring or freezing to a depth of 5-10 cm was recommended. Chemical treatment of drained ponds with chloride of lime (0.5-0.6 ton /hectare) in autumn decreased transmission. Chlorination (50 p.p.m bleaching powder -- Liao and Shih, 1956) or a combination of chlorination and freezing is even more efficacious (Bauer et al., 1969).

Management Strategies in the Wild

(reviewed by Choudhury and Cole, 2012)

Eradicating a parasite like B. acheilognathi in open systems such as interconnected lakes and streams is improbable. In systems where the parasite depends on the continued presence of an introduced host, as in Australia, eradicating that host (common carp, Cyprinus carpio) may eliminate the parasite. However, given the abundance of carp in eastern Australia, this seems unlikely to be feasible. Ward (2007) discussed a strategy for systematically deworming humpback chub (Gila cypha) from the Little Colorado River in the U.S. southwest, a river that is isolated by seasonally dry upstream regions and the cold Colorado River downstream. The abundance of the tapeworm from the Little Colorado River appears to have declined following this strategy.

References

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18/05/2015 Original text by:

Rebecca A. Cole, US Geological Survey, National Wildlife Health Center, Madison, Wisconsin, U.S.A.

Anindo Choudhury, Division of Natural Sciences, St. Norbert College, De Pere, Wisconsin, U.S.A.

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