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nematodes in fish

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nematodes in fish

Summary

  • Last modified
  • 03 January 2018
  • Datasheet Type(s)
  • Animal Disease
  • Preferred Scientific Name
  • nematodes in fish
  • Pathogens
  • Nematoda (Nematodes)
  • Overview
  • Parasitic nematodes constitute one of the earliest known groups of helminths in fishes. They infect freshwater, marine and brackishwater fish species and sometimes cause substantial damage to the host. Although parasiti...

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PictureTitleCaptionCopyright
Philometra abdominalis filling the abdominal cavity of a gudgeon (Gobio gobio) (x 1).
TitleAbdominal cavity
CaptionPhilometra abdominalis filling the abdominal cavity of a gudgeon (Gobio gobio) (x 1).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Philometra abdominalis filling the abdominal cavity of a gudgeon (Gobio gobio) (x 1).
Abdominal cavityPhilometra abdominalis filling the abdominal cavity of a gudgeon (Gobio gobio) (x 1).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Hysterothylacium bidentatum specimens in the stomach of the sterlet (Acipenser ruthenus) (x 2). Photo by György Csaba.
TitleSpecimens in stomach of sterlet
CaptionHysterothylacium bidentatum specimens in the stomach of the sterlet (Acipenser ruthenus) (x 2). Photo by György Csaba.
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Hysterothylacium bidentatum specimens in the stomach of the sterlet (Acipenser ruthenus) (x 2). Photo by György Csaba.
Specimens in stomach of sterletHysterothylacium bidentatum specimens in the stomach of the sterlet (Acipenser ruthenus) (x 2). Photo by György Csaba.Kálmán Molnár, Kurt Buchmann & Csaba Székely
Heavy and moderate infections with Anguillicola crassus in opened swim bladders of European eels (x 1).
TitleBladder infections
CaptionHeavy and moderate infections with Anguillicola crassus in opened swim bladders of European eels (x 1).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Heavy and moderate infections with Anguillicola crassus in opened swim bladders of European eels (x 1).
Bladder infectionsHeavy and moderate infections with Anguillicola crassus in opened swim bladders of European eels (x 1).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Female specimen of Capillaria pterophylli in the gut of a discus fish (Symphysodon discus). Photo by Ferenc Baska.
TitleFemale specimen of Capillaria pterophylli
CaptionFemale specimen of Capillaria pterophylli in the gut of a discus fish (Symphysodon discus). Photo by Ferenc Baska.
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Female specimen of Capillaria pterophylli in the gut of a discus fish (Symphysodon discus). Photo by Ferenc Baska.
Female specimen of Capillaria pterophylliFemale specimen of Capillaria pterophylli in the gut of a discus fish (Symphysodon discus). Photo by Ferenc Baska.Kálmán Molnár, Kurt Buchmann & Csaba Székely
Anterior end of Anisakis simplex with interlabia (arrows) around mouth opening. Scanning electron microscope (SEM) x 200.
TitleAnterior end of Anisakis simplex
CaptionAnterior end of Anisakis simplex with interlabia (arrows) around mouth opening. Scanning electron microscope (SEM) x 200.
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Anterior end of Anisakis simplex with interlabia (arrows) around mouth opening. Scanning electron microscope (SEM) x 200.
Anterior end of Anisakis simplexAnterior end of Anisakis simplex with interlabia (arrows) around mouth opening. Scanning electron microscope (SEM) x 200.Kálmán Molnár, Kurt Buchmann & Csaba Székely
Buccal capsule of Camallanus truncatus from the gut of pike perch (Sander lucioperca) (´ 200).
TitleBuccal capsule of Camallanus truncatus
CaptionBuccal capsule of Camallanus truncatus from the gut of pike perch (Sander lucioperca) (´ 200).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Buccal capsule of Camallanus truncatus from the gut of pike perch (Sander lucioperca) (´ 200).
Buccal capsule of Camallanus truncatusBuccal capsule of Camallanus truncatus from the gut of pike perch (Sander lucioperca) (´ 200).Kálmán Molnár, Kurt Buchmann & Csaba Székely
The anterior part of Cucullanus heterochrous showing mouth opening. SEM x 600.
TitleThe anterior part of Cucullanus heterochrous
CaptionThe anterior part of Cucullanus heterochrous showing mouth opening. SEM x 600.
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
The anterior part of Cucullanus heterochrous showing mouth opening. SEM x 600.
The anterior part of Cucullanus heterochrousThe anterior part of Cucullanus heterochrous showing mouth opening. SEM x 600.Kálmán Molnár, Kurt Buchmann & Csaba Székely
(a) Ovoviviparous eggs of Anguillicola crassus. Second-stage larvae inside the extremely thin egg shells, (b) Viviparous larvae of Philometra abdominalis released from a ruptured female (x 150).
TitleLarvae
Caption(a) Ovoviviparous eggs of Anguillicola crassus. Second-stage larvae inside the extremely thin egg shells, (b) Viviparous larvae of Philometra abdominalis released from a ruptured female (x 150).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
(a) Ovoviviparous eggs of Anguillicola crassus. Second-stage larvae inside the extremely thin egg shells, (b) Viviparous larvae of Philometra abdominalis released from a ruptured female (x 150).
Larvae(a) Ovoviviparous eggs of Anguillicola crassus. Second-stage larvae inside the extremely thin egg shells, (b) Viviparous larvae of Philometra abdominalis released from a ruptured female (x 150).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Third-stage larvae of Anguillicola crassus (arrows) in the haemocoel of the intermediate host cyclops (x 100).
TitleLarvae of Anguillicola crassus
CaptionThird-stage larvae of Anguillicola crassus (arrows) in the haemocoel of the intermediate host cyclops (x 100).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Third-stage larvae of Anguillicola crassus (arrows) in the haemocoel of the intermediate host cyclops (x 100).
Larvae of Anguillicola crassusThird-stage larvae of Anguillicola crassus (arrows) in the haemocoel of the intermediate host cyclops (x 100).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Third-stage larvae of a Skrjabillanus sp. (arrow) entering the stylet of the intermediate host carp lice (Argulus foliaceus) (x 66).
TitleLarvae of a Skrjabillanus sp.
CaptionThird-stage larvae of a Skrjabillanus sp. (arrow) entering the stylet of the intermediate host carp lice (Argulus foliaceus) (x 66).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Third-stage larvae of a Skrjabillanus sp. (arrow) entering the stylet of the intermediate host carp lice (Argulus foliaceus) (x 66).
Larvae of a Skrjabillanus sp.Third-stage larvae of a Skrjabillanus sp. (arrow) entering the stylet of the intermediate host carp lice (Argulus foliaceus) (x 66).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Encapsulated dead and live third-stage larvae of Anguillicola crassus in the gut wall of the paratenic host ruffe (Gymnocephalus cernuus) (x 40).
TitleLarvae of Anguillicola crassus
CaptionEncapsulated dead and live third-stage larvae of Anguillicola crassus in the gut wall of the paratenic host ruffe (Gymnocephalus cernuus) (x 40).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Encapsulated dead and live third-stage larvae of Anguillicola crassus in the gut wall of the paratenic host ruffe (Gymnocephalus cernuus) (x 40).
Larvae of Anguillicola crassusEncapsulated dead and live third-stage larvae of Anguillicola crassus in the gut wall of the paratenic host ruffe (Gymnocephalus cernuus) (x 40).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Histological section of the gut of a fingerling of the paratenic host sheatfish (Silurus glanis). Third-stage larvae (arrows) of A. crassus encapsulated by epithelioid cells and connective tissue. Haematoxylin and eosin (H & E) x 135.
TitleGut of a fingerling
CaptionHistological section of the gut of a fingerling of the paratenic host sheatfish (Silurus glanis). Third-stage larvae (arrows) of A. crassus encapsulated by epithelioid cells and connective tissue. Haematoxylin and eosin (H & E) x 135.
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Histological section of the gut of a fingerling of the paratenic host sheatfish (Silurus glanis). Third-stage larvae (arrows) of A. crassus encapsulated by epithelioid cells and connective tissue. Haematoxylin and eosin (H & E) x 135.
Gut of a fingerlingHistological section of the gut of a fingerling of the paratenic host sheatfish (Silurus glanis). Third-stage larvae (arrows) of A. crassus encapsulated by epithelioid cells and connective tissue. Haematoxylin and eosin (H & E) x 135.Kálmán Molnár, Kurt Buchmann & Csaba Székely
Anisakis simplex (arrow) penetrating pyloric caecum of rainbow trout and causing mechanical damage of the wall (arrowheads) (experimental infection) (SEM x 90).
TitleAnisakis simplex
CaptionAnisakis simplex (arrow) penetrating pyloric caecum of rainbow trout and causing mechanical damage of the wall (arrowheads) (experimental infection) (SEM x 90).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Anisakis simplex (arrow) penetrating pyloric caecum of rainbow trout and causing mechanical damage of the wall (arrowheads) (experimental infection) (SEM x 90).
Anisakis simplexAnisakis simplex (arrow) penetrating pyloric caecum of rainbow trout and causing mechanical damage of the wall (arrowheads) (experimental infection) (SEM x 90).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Damaged swim bladders of the European eels that died during the eel mortality in 1991 in Lake Balaton. One of the swim bladders contains numerous A. crassus specimens, while others have thickened fibrous walls as a consequence of past infection (x 0.8).
TitleDamaged swim bladders of the European eels
CaptionDamaged swim bladders of the European eels that died during the eel mortality in 1991 in Lake Balaton. One of the swim bladders contains numerous A. crassus specimens, while others have thickened fibrous walls as a consequence of past infection (x 0.8).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Damaged swim bladders of the European eels that died during the eel mortality in 1991 in Lake Balaton. One of the swim bladders contains numerous A. crassus specimens, while others have thickened fibrous walls as a consequence of past infection (x 0.8).
Damaged swim bladders of the European eelsDamaged swim bladders of the European eels that died during the eel mortality in 1991 in Lake Balaton. One of the swim bladders contains numerous A. crassus specimens, while others have thickened fibrous walls as a consequence of past infection (x 0.8).Kálmán Molnár, Kurt Buchmann & Csaba Székely
With heavy infections with A. crassus, the worms die and decay in the lumen of the thickened swim bladder (arrow) (x 1.2).
TitleInfections with A. crassus
CaptionWith heavy infections with A. crassus, the worms die and decay in the lumen of the thickened swim bladder (arrow) (x 1.2).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
With heavy infections with A. crassus, the worms die and decay in the lumen of the thickened swim bladder (arrow) (x 1.2).
Infections with A. crassusWith heavy infections with A. crassus, the worms die and decay in the lumen of the thickened swim bladder (arrow) (x 1.2).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Cross-section of a third-stage A. crassus larvae in the oedematous tissue of the submucosa of the eel's swim bladder (H & E x 250).
TitleThird-stage A. crassus larvae
CaptionCross-section of a third-stage A. crassus larvae in the oedematous tissue of the submucosa of the eel's swim bladder (H & E x 250).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Cross-section of a third-stage A. crassus larvae in the oedematous tissue of the submucosa of the eel's swim bladder (H & E x 250).
Third-stage A. crassus larvaeCross-section of a third-stage A. crassus larvae in the oedematous tissue of the submucosa of the eel's swim bladder (H & E x 250).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Fourth-stage larva of A. crassus accumulating in the gas gland, before entering the swim bladder (x 20).
TitleFourth-stage larva of A. crassus
CaptionFourth-stage larva of A. crassus accumulating in the gas gland, before entering the swim bladder (x 20).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Fourth-stage larva of A. crassus accumulating in the gas gland, before entering the swim bladder (x 20).
Fourth-stage larva of A. crassusFourth-stage larva of A. crassus accumulating in the gas gland, before entering the swim bladder (x 20).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Cross-section of a third-stage A. crassus larva inside the swim-bladder wall surrounded by granulation tissue and mononuclear cell infiltration (H & E x 150).
TitleThird-stage A. crassus larva
CaptionCross-section of a third-stage A. crassus larva inside the swim-bladder wall surrounded by granulation tissue and mononuclear cell infiltration (H & E x 150).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Cross-section of a third-stage A. crassus larva inside the swim-bladder wall surrounded by granulation tissue and mononuclear cell infiltration (H & E x 150).
Third-stage A. crassus larvaCross-section of a third-stage A. crassus larva inside the swim-bladder wall surrounded by granulation tissue and mononuclear cell infiltration (H & E x 150).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Females of Philometroides sanguinea in the blood vessels of the caudal fin of a gibel carp (Carassius gibelio) (x 1.5).
TitleFemales of Philometroides sanguinea
CaptionFemales of Philometroides sanguinea in the blood vessels of the caudal fin of a gibel carp (Carassius gibelio) (x 1.5).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Females of Philometroides sanguinea in the blood vessels of the caudal fin of a gibel carp (Carassius gibelio) (x 1.5).
Females of Philometroides sanguineaFemales of Philometroides sanguinea in the blood vessels of the caudal fin of a gibel carp (Carassius gibelio) (x 1.5).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Parasitic nodules (arrows) in the skin caused by Cystoopsis acipenseris in the abdominal side of sterlet (Acipenser ruthenus) (x 0.7). Photo by Ferenc Baska.
TitleParasitic nodules
CaptionParasitic nodules (arrows) in the skin caused by Cystoopsis acipenseris in the abdominal side of sterlet (Acipenser ruthenus) (x 0.7). Photo by Ferenc Baska.
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
Parasitic nodules (arrows) in the skin caused by Cystoopsis acipenseris in the abdominal side of sterlet (Acipenser ruthenus) (x 0.7). Photo by Ferenc Baska.
Parasitic nodulesParasitic nodules (arrows) in the skin caused by Cystoopsis acipenseris in the abdominal side of sterlet (Acipenser ruthenus) (x 0.7). Photo by Ferenc Baska.Kálmán Molnár, Kurt Buchmann & Csaba Székely
X-ray as a tool for diagnosis of A. crassus infection of eel swim bladder. Note the large convoluted worms (arrows) in the swim bladder and the ductus pneumaticus (x 1.3).
TitleDiagnosis of A. crassus infection
CaptionX-ray as a tool for diagnosis of A. crassus infection of eel swim bladder. Note the large convoluted worms (arrows) in the swim bladder and the ductus pneumaticus (x 1.3).
CopyrightKálmán Molnár, Kurt Buchmann & Csaba Székely
X-ray as a tool for diagnosis of A. crassus infection of eel swim bladder. Note the large convoluted worms (arrows) in the swim bladder and the ductus pneumaticus (x 1.3).
Diagnosis of A. crassus infectionX-ray as a tool for diagnosis of A. crassus infection of eel swim bladder. Note the large convoluted worms (arrows) in the swim bladder and the ductus pneumaticus (x 1.3).Kálmán Molnár, Kurt Buchmann & Csaba Székely
Female Capillaria philippinensis (without eggs, x 160) (courtesy of Dr John Cross).
TitleFemale Capillaria philippinensis (without eggs)
CaptionFemale Capillaria philippinensis (without eggs, x 160) (courtesy of Dr John Cross).
CopyrightRonald C. Ko
Female Capillaria philippinensis (without eggs, x 160) (courtesy of Dr John Cross).
Female Capillaria philippinensis (without eggs)Female Capillaria philippinensis (without eggs, x 160) (courtesy of Dr John Cross).Ronald C. Ko
Egg of Capillaria philippinensis showing two polar plugs (x 400, courtesy of Dr John Cross).
TitleEgg of Capillaria philippinensis
CaptionEgg of Capillaria philippinensis showing two polar plugs (x 400, courtesy of Dr John Cross).
CopyrightRonald C. Ko
Egg of Capillaria philippinensis showing two polar plugs (x 400, courtesy of Dr John Cross).
Egg of Capillaria philippinensisEgg of Capillaria philippinensis showing two polar plugs (x 400, courtesy of Dr John Cross).Ronald C. Ko
En face view of head of young male adult Gnathostoma spinigerum from stomach of experimentally infected cat, showing the large pseudolabia and cephalic spines (x 400, original).
TitleYoung male adult Gnathostoma spinigerum
CaptionEn face view of head of young male adult Gnathostoma spinigerum from stomach of experimentally infected cat, showing the large pseudolabia and cephalic spines (x 400, original).
CopyrightRonald C. Ko
En face view of head of young male adult Gnathostoma spinigerum from stomach of experimentally infected cat, showing the large pseudolabia and cephalic spines (x 400, original).
Young male adult Gnathostoma spinigerumEn face view of head of young male adult Gnathostoma spinigerum from stomach of experimentally infected cat, showing the large pseudolabia and cephalic spines (x 400, original).Ronald C. Ko
Egg of Gnathostoma spinigerum (original).
TitleEgg of Gnathostoma spinigerum
CaptionEgg of Gnathostoma spinigerum (original).
CopyrightRonald C. Ko
Egg of Gnathostoma spinigerum (original).
Egg of Gnathostoma spinigerumEgg of Gnathostoma spinigerum (original).Ronald C. Ko
Anterior region of third-stage larva of Gnathostoma spinigerum from catfish (Clarias fuscus) (original).
TitleAnterior region of third-stage larva of Gnathostoma spinigerum
CaptionAnterior region of third-stage larva of Gnathostoma spinigerum from catfish (Clarias fuscus) (original).
CopyrightRonald C. Ko
Anterior region of third-stage larva of Gnathostoma spinigerum from catfish (Clarias fuscus) (original).
Anterior region of third-stage larva of Gnathostoma spinigerumAnterior region of third-stage larva of Gnathostoma spinigerum from catfish (Clarias fuscus) (original).Ronald C. Ko

Identity

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Preferred Scientific Name

  • nematodes in fish

Pathogen/s

Top of page Nematoda (Nematodes)

Overview

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Parasitic nematodes constitute one of the earliest known groups of helminths in fishes. They infect freshwater, marine and brackishwater fish species and sometimes cause substantial damage to the host. Although parasitic nematodes can infect almost all organs in a fish, the majority of the currently known species have been described from the intestine. Most nematodes infect fish as adults, but a large proportion of them occur as larval stages. These are usually parasites of piscivorous birds, mammals or reptiles, or less frequently of predatory fishes.

The majority of nematodes reach sexual maturity through a complicated developmental cycle involving an intermediate or possibly paratenic hosts. Species living in the temperate zone are usually characterized by seasonal occurrence, and annual life cycles are common. Because of the complicated, multi-host life cycle, the development of fish nematodes is successful in non-disrupted ecosystems. In fish taken from their natural surroundings, nematode infections are less likely to develop. For these reasons, nematodes cause less damage in cultured fishes than do other helminths. At the same time, certain nematodes can give rise to massive infections with high fish mortality in natural waters.

Although nematodes are important pathogens, their direct pathogenic effect on fishes is much less important than the role played by them as causative agents of zoonoses. Nematode infections in marine fishes cause a range of problems. Some of these are associated with pathogenicity of the parasite to the fish host, while others are health hazards connected to human ingestion of live nematodes in fresh or undercooked fish. Furthermore, consumer attitudes to the presence of nematodes in foods also have a great impact on the market value of fish products and may stress the need for strict monitoring and diagnosis of nematode infections in marine fish stocks.

The prevalence of parasitic nematodes on the different continents is not equally well known. The majority of species have been described from Europe. Detailed data on nematodes of freshwater fishes (Avdeev et al., 1987; Moravec, 1994) and those in North American fishes (Hoffman, 1999) are readily available. Recently major progress has been made in the study of fish-parasitic nematodes in the Neotropical region (Moravec, 1998). Of the nematodes parasitizing marine fishes, anisakid species causing human infections are relatively well documented (Hauck and May, 1977; Smith and Wootten, 1978; Berland, 1981; Sindermann, 1990; Koie et al., 1995). For relevant data on the occurrence and pathological role of fish-parasitic nematodes see Dick and Choudhury (1995); since the publication of that work, relatively little new information has been added to our existing knowledge of fish-parasitic nematodes. The new data concern primarily the development, pathological effect and human health implications of a few selected groups (Anguillicola, Philometra, Skrjabillanus, Anisakis).

Knowledge of fish nematodes varies greatly. The nematode fauna of fish in Europe and the northern parts of Asia and America is relatively well studied. Little is known, however, about fish nematodes of South Asia, Africa, Australia and Latin America. All nematodes in marine fishes with zoonotic relevance have been studied and, of these, large-sized ascaridoid and philometrid nematodes are the best known.

Small-sized nematodes, such as the skrjabillanids, are known mostly from Europe, although recent studies on fishes of Central America show that their distribution may also be high in other continents. In most cases, only the intestinal tract is examined and histozoic nematodes infecting inner organs, serous membranes and muscles may not be found by inexperienced examiners.

There is an urgent need for the introduction of molecular methods in the research of fish nematodes. This method can be useful in synonymizing or separating morphologically similar species and finding latent infections with small histozoic nematodes.

Pathogenicity of fish parasites is mostly based on field observations, except studies done on A. crassus infections. R. acus and A. simplex also seem to be experimental tools for studying pathogenic effect of nematodes.

There is no effective drug against fish nematodes. Although nematodes cause relatively few problems in propagated fishes in fish farms or cage cultures, they are relatively important in aquarium fishes.

[Derived from: Woo, PTK, ed., 2006. Fish diseases and disorders, Volume 1: Protozoan and Metazoan infections. (2nd edition) Wallingford, UK: CAB International]

Hosts/Species Affected

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Host Range

Nematode infection occurs in practically all fish species and in all locations. The prevalence, intensity and economic importance of nematode infections, however, vary by region and by fish species.

The host range of these parasites is primarily influenced by the host specificity of the nematode species for the final host and for the intermediate host. Species having a narrow host range are usually in a fish host that is prevalent in a given habitat, while species with a broad host range are in fish species all over the world. These nematodes can often cause mild disease in all members of a fish genus and, after colonizing a new host species, they can give rise to much more severe disease. Such a species is A. crassus, a nematode of Japanese eel (Anguilla japonica), which causes a mild disease in its original hosts. However, this nematode is much more pathogenic in European and American eel species (Anguilla rostrata).

The typical examples of highly specific nematodes are the members of different skrjabillanid genera (Skrjabillanus tincae, Skrjabillanus cyprini, Molnaria intestinalis, Sinoichthyonema amuri), which in most cases infect a single fish species. Nematodes with a broad host range are represented by Capillaria species, which can colonize numerous fish species of different taxonomic positions. Fish parasites characterized by a global range include the larval stages of nematodes of seabirds and marine mammals in their adult stage (Anisakis, Pseudoterranova). The distribution of the majority of freshwater nematode species is mostly restricted to a single continent or specific zoogeographical zones thereof.

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

North America

USAPresentPresent based on regional distribution.
-FloridaPresentGaines and Rogers, 1972

South America

BrazilPresentFreitas and Lent, 1946

Europe

HungaryPresentMolnár et al., 1991; Molnár et al., 1993
Russian FederationPresentDogiel and Bykhovskiy, 1939; Dubinin, 1952

Pathology

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Immune reactions



Nematodes elicit specific antibody production in the host. Migration of the larval stages in host cavities and host tissue may expose both structural and metabolic antigens to the host immune system. It has been demonstrated by immunoblotting that the European eel produces specific antibodies against a number of antigens of the swim bladder nematode A. crassus (Buchmann et al., 1991; Hoglund and Pilström, 1994; Békési et al., 1997; Nielsen and Buchmann, 1997; Knopf et al., 2000). Third-stage larvae of A. simplex in the fish host provöke the production of specific antibodies in naturally infected saithe (Priebe et al., 1991). In addition, a range of Antarctic teleosts has been shown to possess reactive antibodies against molecules from C. osculatum (Coscia and Oreste, 1998). Similar positive plasma and bile antibody reactivities in fish against P. decipiens larvae were recorded by Coscia and Oreste (2000). Two main types of antigens are generally recognized in nematodes. These are the soluble excretory and secretory (E/S) antigens and the somatic antigens associated with surfaces on the outer or inner part of the worm. These may be partly protective. However, it is generally agreed that cellular reactions play a major role in protection of the host against nematode infections. Both humoral and cellular host reactions have been detected in various hosts against invading nematodes and these immune factors may function in host elimination and killing of infective stages. However, the roundworms seem to have some evasion mechanisms. Thus, the cuticle of nematodes consists in most cases of a protective proteinaceous layer, which protects the inner vital organs of the worm against aggressive immune effectors. Even encapsulated Anisakis and Anguillicola worms recovered from infected fish are fully capable of moving vividly upon removal of the host encapsulation (Santamarina et al., 1994; Székely, 1994; Larsen et al., 2002). These mechanisms have not been adequately investigated.

Pathogenicity

Pathological effect

The pathological effect of nematode infection in fish is little studied, and most information is based on field observations. There are only a few reported cases of mortality due to nematode infections. Most authors (Bauer et al., 1977; Moravec, 1994; Dick and Choudhury, 1995) agree that fish nematodes damage the hosts by depriving the fish of digested food; by feeding on host tissues, sera or blood; and by direct mechanical damage through fixing to host tissues and developing or migrating in them (Fig. 14). Nematodes generally possess a range of enzymes, such as proteases, which may have tissue-degrading functions (Newton and Munn, 1999). Large-sized parasites compress organs (Platzer and Adams, 1967), deform the shape of the body (Molnár, 1966a), increase or reduce the size of organs (Paperna, 1974) and cause haemorrhages (Jilek and Crites, 1982; Dunn et al., 1983), inflammation (Measures, 1988; Molnár et al, 1993), granulomas (Hauck and May, 1977; Sindermann, 1990), ascites (Bauer et al., 1977) and mesenteric and visceral adhesions (Sindermann, 1990). The pathogenic effect depends on the species and the size and the number of parasites (Fig. 3), and survival of the fish also depends on the site of infection.

Mortality

Mortality caused by nematodes was described by Bauer and Zmerzlaya (1972), who indicated that R. acus larvae caused heavy mortalities in bream (Abramis brama). According to Bauer et al. (1977), the heavily infected bream lost their balance and swam on their side, their body was covered by a thick layer of slime, there was a local destruction of their sexual glands and bloody exudates accumulated in their abdominal cavity. Eiras and Reichenbach- Klinke (1982) also described heavy infection and deformation of the rainbow trout's intestine due to large parasitic nodules caused by R. acus.Moravec and Gut (1982) and Moravec et al. (1984) reported on the mortality of ornamental fishes due to massive infection with Pseudocapillaria brevispicula and Capillaria pterophylli (Fig. 5). Schäperclaus (1992) also found that Pseudocapillaria tomentosa can severely damage tench (Tinca tinca). A devastating effect of nematode infection was observed by Molnár et al. (1991, 1993), who reported on a massive eel die in Lake Balaton, Hungary, due to A. crassus (Figs. 15 and 16) infection. In this lake an estimated 400 tons of eels died in 1991, 1992 and 1995. Similar heavy mortalities caused by this nematode were observed in the Czech Republic by Barus (1995). Goezia spp. infecting the stomach of the fish seem to have a relatively high pathogenic effect. These worms bore their anterior ends deep into mucosa up to the muscularis layer. Gaines and Rogers (1972) observed that they form deep nodules in the stomach wall. These authors also reported mortalities in striped bass (Morone saxatilis) and tilapia in Florida. Mortalities were observed by Freitas and Lent (1946) in Arapaima gigas in Brazil caused by Goezia spinulosa.

Fig. 14. Anisakis simplex (arrow) penetrating pyloric caecum of rainbow trout and causing mechanical damage of the wall (arrowheads) (experimental infection) (SEM).

Fig. 15. Damaged swim bladders of the European eels that died during the eel mortality in 1991 in Lake Balaton. One of the swim bladders contains numerous A. crassus specimens, while others have thickened fibrous walls as a consequence of past infection.

Fig. 16. With heavy infections with A. crassus, the worms die and decay in the lumen of the thickened swim bladder (arrow).

Trichuroid nematodes are generally considered pathogenic. These nematodes damage epithelial cells by feeding and penetrating deeply into the intestinal mucosa. The necrotic changes they cause often lead to morbidity of the hosts. Dubinin (1952) in Russia reported on disease through gut inflammation of acipenserids (A. ruthenus, Acipenser nudiventris) caused by Pseudocapillaria tuberculata. Severe debilitating conditions also developed on some North American centrarchid fishes. Capillaria catostomi caused enteritis in the caeca and intestine (Hoffman, 1982). Eustrongylides spp. are also pathogenic. The pathogenicity of Eustrongylides spp. in sturgeons (A. nudiventris) was studied by Dubinin (1952) and Dogiel and Bykhovskiy (1939) in Russia, who considered Eustrongylides larvae to be highly pathogenic, causing heavy infections leading to complete destruction of gonads and to parasitic castration of infected fishes. It was also Dubinin (1952) who found that, in heavy infections with H. bidentatum, inflammatory processes were found in the intestinal walls of sterlets (A. ruthenus) and sometimes perforation of the swim bladder by migrating worms was recorded. Kall et al. (2004), who examined P. obturans infection of pike (Esox lucius), reported that the pikes infected by this large worm inhabiting gill arteries were less active, showed lethargy and died in aquaria shortly after transportation to the laboratory. Sprengel and Lüchtenberg (1991) experimentally proved that A. crassus infection in eels reduced swimming performance, and in another experiment Molnár (1993) found that heavily infected eels were more susceptible to decreased oxygen content in the water.

Intestinal tract

Most Nematoda species infect the intestinal tract. Some of them (e.g. Hysterothylacium, Goezia, Cystidicola spp.) prefer the anterior part of the alimentary tract (oesophagus, stomach, pyloric region), while others are in the intestine. The major damage caused by these worms is associated with their consumption of intestinal contents, thereby depriving the host of nutrients. A less important effect comes from direct mechanical blockage by the worms.

H. bidentatum (Fig. 2), a common parasite of the acipenserids, may cause heavy infections in sterlet. These large-sized nematodes can completely occlude the stomach and thus reduce digestion and block the passage of food. After the death of heavily infected fish, these nematodes frequently migrate out through the mouth or the gill slit.

At less intensive infections, nematodes may evoke pathological changes, mostly around their attachment sites. Damage to the mucosa or deeper tissues is usually caused by lips, the buccal capsule, teeth or spines.

Rhabdochona species, when in high numbers, can cause perforation of the intestinal wall at attachment points (Moravec, 1975). Similar damage occurs with Camallanus, Procamallanus and Paracamallanus species as they 'grab' the intestinal wall with their buccal capsules while feeding on blood. Usually there is a local inflammatory reaction at the attachment site. Thatcher (1991), as well as Sinha and Sinha (1988), suggested that nematodes could cause primary anaemia by feeding on blood. In intensive infections, especially in small fishes, these camallanids can reduce growth rates and also cause intestinal blockage. More severe changes were recorded in ornamental fishes. Several authors (Petter et al., 1974; Stumpp, 1975; Campana-Rouget et al., 1976; Schäperclaus, 1992) reported on complete destruction of the intestinal mucosa and death of the fish in the presence of large number of Camallanus fotedari or Camallanus moraveci. Heavy infection with Camallanus cotti caused a reduced sexual display rate in Poecilia reticulata (McMinn, 1990).

Cucullanus truttae has similar effects on rainbow trout. According to Dunn et al. (1983), there is loss of epithelium and mucosal hyperplasia, as well as haemorrhage and fibrosis in the laminar propria at the point of attachment. Growth rate, food consumption and swimming activity are reduced in infected fish. The spiruroid nematodes Camallanus oxycephalus and Spinitechtus carolini of the green sunfish penetrate to the mucosal layer of the gut and cause damage to the columnar epithelium. At the site of penetration, ulcers developed in the mucosal and submucosal layers and there was growth of granulomatous tissue with extensive fibrosis (Meguid and Eure, 1996). Local changes in the intestine can also be provoked by seemingly less pathogenic nematodes.

In the case of Echinocephalus daileyi, where there is a special cephalic inflation and rows of hooks for attachment of the worm to the intestinal mucosa, Thatcher (1991) observed inflammation and formation of a fibrous capsule around the head bulb. Formation of capsules filled by tissue debris, oedematous fluid, fibrous exudate and leucocytes at the attachment point of the nematodes was also observed by Ko et al. (1975) with Echinocephalus sinensis in the ray Aetobatus flagellum.

Deardorff and Overstreet (1980) remarked that Goezia pelagia apparently feeds on both the host elements and partially digested food. The parasite forms a deep nodule in the intestinal wall, with the development of ulcers. A connective-tissue capsule surrounds the head part of the parasite, and primary exudates, inflammatory cells, red blood cells and necrotic tissue are in the nodules. Aeromonas bacteria could be cultured from some of the nodules. It is often larval penetration and migration that cause severe reactions. Thus, invasion of host tissue by marine nematode larvae has been described to cause pathological reactions. Haemorrhages, tissue compression and necrosis are often found in tissue with invading P. decipiens larvae (Ramakrishna et al., 1993). The host reaction may be expressed in tissue proliferation, degeneration and inflammation.

Molnár (1994) found hundreds of A. crassus larvae in nodules in the intestinal wall; some of the larvae were alive while others were dead and calcified. Similar observations were reported by Janiszewska (1939) with D. minutus in flatfishes. Jilek and Crites (1982), who studied the pathogenicity of the habronematoid S. carolini in centrarchid fishes, described the third-stage larvae penetrating the intestinal wall, causing traumatic enteritis, the growth of epithelioid fibroblasts around worms, and accumulation of granule cells, leucocytes and macrophages. An expanding fibrocytic layer formed a capsule around the larvae; the innermost layer became necrotic but encapsulated worms were able to develop into adults. A. simplex larvae have been seen to induce severe inflammatory reactions in the wall of the stomach of cod. Thus clusters of larvae gathered in local inflammatory foci in the stomach wall of the fish host (Berland, 1981). Arai (1969) found large ulcers caused by Anisakis larvae in the stomach wall of Ophiodon elongatus, and Williams and Richards (1968) observed prominent host reactions in Raja radiata against Pseudanisakis rotunda, especially the head, which was in the lamina submucosa granulation tissue.

Body cavity

The body cavity is the frequent location of Philometra and Philonema; however, little is known about their pathogenic effect. Philonema spp. are known to be responsible for multiple mesenteric and visceral adhesions in salmonid fishes (Nagasawa, 1985; Garnick and Margolis, 1990; Sindermann, 1990). Both Philonema and Philometra infection can cause atrophy or destruction of gonads, ascites and extension of the abdomen (Molnár, 1966a, 1967; Platzer and Adams, 1967; Williams, 1967; Hoffman, 1975; Moravec et al., 2003). While studying Eustrongylides sp. larvae infecting mesenteries and inner organs of African fishes of the genera Haplochromis, Bagrus and Clarias, Paperna (1974) found that the most heavily infected fish were emaciated. In these fish an extensive lysis was observed around the worms, which penetrated the somatic muscles. Inner organs were infiltrated by lymphocytes and macrophages and inflammatory necrosis was observed. Eiras and Rego (1988) reported on similar changes in South American fishes (Pygocentrus natteri), while Measures (1988) in North America found granulomatous inflammation and exudates containing erythrocytes and macrophages. However, Kennedy and Lie (1976) reported that heavy infections of encapsulated Eustrongylides did not cause weight loss or changes in the outward appearance and condition factor of fish. Experimental infections of rainbow trout with A. simplex third-stage larvae are known to cause a pronounced cellular reaction in the body cavity and in tissue penetrated, especially in the pyloric caeca (Santamarina et al., 1994; Larsen et al., 2002). Larvae of the genera Hysterothylacium, Anisakis, Pseudoterranova and Contracaecum and of Spiroxis contortus are frequently found on the serosa of the gut, where they are coiled and covered only by a thin serosa layer; in other cases they are encapsulated by a row of connective-tissue layers (Smith and Wootten, 1978).

Liver

Of the nematodes damaging the liver, Schulmanela (Hepaticola) petruschewskii is the best-known species. Kutzer and Otte (1966) studied the pathological effects of H. petruschewskii in salmonid, percid and cyprinid fishes. Macroscopic changes of the severely infected fish included greyish discoloration of the liver, with the appearance of nodules the size of a pinhead or larger, which was sometimes accompanied by hyperaemia, petechial haemorrhages and icterus. Histologically, in addition to the presence of helminths in numerous convoluted passages, haemorrhage and hyperaemia of the liver capillaries, aggravated by the appearance of fibrinous-serous exudate, were found. Leucocytic infiltration, epithelioid cell proliferation and even the appearance of giant cells could also be observed.

Devastating effects on host liver tissue were described by Petrushevski and Shulman (1961). Nematode larvae, probably C. osculatum, which normally reside in the liver of Baltic cod (G. morhua), were suggested to affect the size and function of the liver. Thus, heavy parasite burdens were seen in small livers and only a few nematode larvae were in large livers. Although tissue-penetrating worms may cause adverse reactions, further studies should be conducted to elucidate this question. In this context it should be recalled that liver size in gadoids is highly influenced by food composition and energy content (Buchmann and Borresen, 1988). Thus, feeding on low-energy diets such as crustaceans (serving as an intermediate host for C. osculatum) instead of high-energy feed (clupeids) may lead to high parasite burdens and, secondarily, low liver weight.

Of the nematodes causing heavy pathological changes of the liver in the larval stage, R. acus is the best-studied species. In the adult stage this parasite infects the gut of piscivorous fishes, primarily pike. In the latter fish, only local inflammations were registered in the gut. In intermediate host fishes, however, heavy infections and mass mortalities were often registered in rainbow trout, common bream, yellow perch and experimentally infected loach (Barbatula barbatula) (Moravec, 1970; Bauer et al., 1977; Poole and Dick, 1984). Third- and fourth-stage larvae in their true intermediate hosts invaded various internal organs, destroyed blood vessels during larval migration and caused numerous nodules in the intestinal wall, liver, peritoneum and mesentery. Valtonen et al. (1994), who followed R. acus migration in roach (Rutilus rutilus), recorded a chronic granulomatous inflammatory reaction. They remarked that larvae occurred more often in pancreatic tissue than in the liver. These authors also remarked that the infection rate in polluted eutrophic lakes was much higher than in oligotrophic ones. Dezfuli et al. (2000), who studied the effect of larval raphidascariosis in the liver of Phoxinus phoxinus, reported that the inflammatory process was characterized by an increase of rodlet cells, besides granulocytes and epithelioid granulomata.

Authors who examined the effect of Anisakis larvae on host tissues (Mikailova et al., 1964; Prusevich, 1964; Hauck and May, 1977; Smith, 1984) reported that Anisakis larvae invading tissues caused destruction of the liver parenchyma, ruptures of the wall of the blood vessels, small haemorrhages and thrombus formation. The larvae were covered with fibrin infiltrated by leucocytes. At a later stage, a connective-tissue capsule was formed around the larvae.

Swim bladder

The swim bladder of fishes is damaged mostly by Cystidicola and Anguillicola species. Van Banning and Haenen (1990) and Molnár et al. (1993) reported that the acute process of A. crassus larval migration was characterized by epithelial hyperplasia and hyperaemia of the swim bladder wall. In cases of chronic swim bladder inflammation, oedema and hyperplasia of tissues of the tunica propria, submucosa and serosa were observed, as well as granulomatoid infiltration by mononuclear cells and fibrinoid degeneration around larvae. Molnár (1994) remarked that inside the oedematous connective tissue of the swim bladder larvae migrate without causing an observable host reaction in the swimbladder wall or the gas glands (Figs. 17 and 18), but in more advanced cases granulation tissue containing mononuclear cells stops larvae (Fig. 19.) The granulation tissue, built up from epithelioid macrophages, forms a nodule around the larvae, which become surrounded by a capsule. Larvae die and, together with the necrotized epithelioid cells, give rise to amorphous tissue debris. Secondary bacterial infections may change nodules into pustules filled with degenerate cells, inflammatory cells and serum. Kamstra (1990), Van Banning and Haenen (1990) and Molnár et al. (1993) reported that adult A. crassus specimens filling the whole lumen of the swim bladder feed on blood, causing anaemia, epithelial hyperaemia and dilatation of the ductus pneumaticus. Due to the damage caused by larvae and adult worms, the wall of the swim bladder thickens and becomes fibrotic (Fig. 15). Worms in the lumen either leave the bladder and migrate to the gut through the ductus pneumaticus or they die. Dead worms and blood form a hard brown-black mass in the lumen of the swim bladder (Fig. 16), which will eventually contain facultative pathogenic bacteria.

Fig. 17. Cross-section of a third-stage A. crassus larva in the oedematous tissue of the submucosa of the eel's swim bladder.

Fig. 18. Fourth-stage larvae of A. crassus accumulating in the gas gland, before entering the swim bladder.

Fig. 19. Cross-section of a third-stage A. crassus larva inside the swim-bladder wall surrounded by granulation tissue and mononuclear cell infiltration.

Skin

The best-known parasites of the skin come from the genera Philometra and Philometroides. By the time they reach full maturity, the large females of Philometra rischta destroy the subepithelial layer of the gill cover of bleak (Alburnus alburnus), and the worms are separated from both the gill and the aquatic environment by an epithelium bordered by basement membrane. Worms cause ulceration of the gill cover, which becomes completely disintegrated in some cases (Molnár, 1966a; K. Molnár, unpublished). Similar injuries may arise after colonization of the skin of sucker (Catostomus commersonii) by P. nodulosa (Dailey, 1966; Hoffman, 1975, 1999). The very tiny S. cyprini is not pathogenic, while the large females (up to 20 cm in length) of Philometroides cyprini (Fig. 4), a parasite of the Amur wild carp, can cause swellings of the scale sacs, accumulation of serous infiltration and haemorrhages under the scales. After the expulsion of worms, the areas without scales and with ulcers may have secondary microbial infections (Vasilkov, 1967, 1975, Sekretaryuk, 1983). Similar changes in the fins are caused by P. sanguinea infection in Carassius species (Fig. 20) and by Philometra huronensis infection in sucker. These changes lead to fraying of the fin rays, which may break and come off in fragments (Molnár, 1966a; Uhazy, 1978). In sturgeon fishes, especially in the sterlet, C. acipenseris forms large nodules in the connective tissue on the ventral surface of the skin (Fig. 21), causing capsule formation and oedema (Bauer et al., 1977). There is usually a female and a male worm in each nodule. Rockfish infected with adult trichuroid nematodes in the skin have inflammatory reactions with intraepithelial deposition of eggs (Conboy and Speare, 2002).

Fig. 20. Females of Philometroides sanguinea in the blood vessels of the caudal fin of a gibel carp (Carassius gibelio).

Fig. 21. Parasitic nodules (arrows) in the skin caused by Cystoopsis acipenseris in the abdominal side of sterlet (Acipenser ruthenus).

Gills

Nematode infection of the gills is relatively rare. The best-known nematode is P. obturans, a parasite of the gill arteries of the European pike. This large-sized nematode, measuring 20 cm in length, is known to obstruct blood circulation, to feed on blood and to perforate gill arteries when releasing larvae. Moravec and Dyková (1978) and Káli et al. (2004) found parietal trombi in the bulbus arteriosus and endothelial hypertrophy in the ventral aorta. The worms clearly obstructed the arteries; the elastic wall of the vessel was stretched around the parasite and appeared thinner. In cases when the female worm was damaged and only the gonads filled the arteries, the arterial wall became irregular, showing signs of hypertrophy and hyperplasia. In contrast to this large Nematoda species, Ribu and Lester (2004) found tiny histozoic nematodes in gill filaments and described them as Moravecia australiensis.

Eyes

The eyes of fishes may frequently have worms. For example, Parukhin (1975) found the females of Philometra oveni in the orbits of a percid fish (Serranelus hepatus) and Moravec and Dyková (1978) found larval stages of P. obturans in the vitreous humour.

Diagnosis

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Identification and Diagnosis of Infection

General methods for identifying nematodes are described in textbooks by Bykhovskaya- Pavlovskaya (1969), Fernando et al. (1972), Bauer et al. (1977), Bauer (1987), Sindermann (1990) and Moravec (1994).

Methods for diagnosis of nematodes much depend on the size and location of the parasites. In live fish, only large parasites on or close to the surface of the body are recognized. Red-coloured Philometra species in the opercula and the fins or parasite nodules in the skin around C. acipenseris can easily be observed. Large nematodes inhabiting the gut and inner organs are easily detected on dissection of freshly killed, frozen or formalin-fixed animals; in the case of small nematodes, a dissecting microscope is necessary. Infection with histozoic species, such as the small skrjabillanid nematodes, can be diagnosed on live material under a microscope. Scrapings of intestinal serosa can be examined under a dissecting microscope for the delicate capillariid nematodes and larval stages. Larval nematodes in the internal organs are usually found in squash preparations between two glass plates. Physiological 0.6% fish saline is necessary to keep nematodes alive. Before fixing, nematodes are rinsed in saline. For fixation, a hot mixture of 70% ethanol and glycerine (9 : 1 part) or a hot mixture of saline and 40% formalin can be used. For diagnosis of Anguillicola infection in the lumen of the swim bladder, Beregi et al. (1998) suggested X-ray, (Fig. 22) and Székely et al. (2004) used computer tomographic methods.

Fig. 22. X-ray as a tool for diagnosis of A. crassus infection of eel swim bladder. Note the large convoluted worms (arrows) in the swim bladder and the ductus pneumaticus.

Male genital organs can be studied by placing the worms in glycerine or lactophenol under a cover slip for clearing. For staining permanent mounts, carmine staining or Thatcher's method (Petter and Thatcher, 1988) is suggested. Oral papillae can be studied by en face preparations (e.g. as suggested by Anderson, 1958). Identification is based on morphological characters. Thus, relative proportion of length and width is used. An important feature is the overall shape of the worm, specifically the presence of papillae, alae, boring tooth, striations, oral opening, excretory pore opening location, caeca and appendages on oesophagus, ventriculus and intestine. As an example, it can be mentioned that the identification of a number of species (P. decipiens, Pseudoterranova krabbei, Pseudoterranova bulbosa and Pseudoterranova azarasi), comprising a species complex previously referred to as P. decipiens (Paggi et al., 2000), has traditionally been based on such structures. Morphologically, the worm is characterized by an anteriorly directed intestinal caecum, no ventricular caecum and an excretory pore opening at the nerve ring. Most ascaridoids are clearly differentiated from Anisakis spp. Anisakis larvae are easily identified by the lack of caeca on the intestine and ventriculus. Likewise a characteristic tail spine is found. The excretory pore is located anterior to the nerve ring (Smith and Wootten, 1978). The differential diagnosis of C. osculatum is also feasible using morphology. Both ventricular and intestinal caeca are found and the excretory pore opens anterior to the nerve ring. Species of Porrocaecum and Phocascaris are difficult to distinguish morphologically, but the former genus is presumed to be in birds while the latter genus is in seals (Paggi et al., 2000). Some similarity exists between Contracaecum spp. and Hysterothylacium spp. larvae. However, several differences are found, among which the location of the opening of the excretory pore is important. The third-stage larva of Hysterothylacium has no lips and carries characteristic caeca on both the intestine and ventriculus, but the excretory pore opens below the nerve ring (Fagerholm, 1982). Likewise, morphological differences are obviously used to diffentiate adult nematodes in fish. The identification of cucullanids is based on the shape and length of the worms, the location of the vulva and the shape of the spicule (Berland, 1970).

Molecular tools

The recent development of molecular techniques has provided alternative and, in some cases, more accurate diagnostic tools. Thus, PCR-based methods include amplification of genes encoding ribosomal RNA. Direct alignment and comparison of DNA sequences may reveal differences between species. By using the PCR-restriction fragment length polymorphism (RFLP) method, various closely related nematodes can be differentiated on their banding pattern in an agarose gel (Kijewska et al., 2002; Szostakowska et al., 2002). Further, mitochondrial DNA sequences have been found to be valuable for differentiation between sibling species within Contracaecum ogmorhini (Mattiucci et al., 2003). In addition, electrophoresis of enzymes from various nematodes and subsequent comparison of gel migration distances allow separation of such species. These techniques were also used for confirmation of mitochondrial DNA results from C. ogmorhini studies (Mattiucci et al., 2003). These have been found feasible for differentiation between the genera Anisakis, Contracaecum and Pseudoterranova (Mattiucci et al., 1998). P. decipiens was formerly considered to be a single, well-defined species, but recent allozyme work has shown that this entity comprises a species complex like P. decipiens (sensu stricto), P. krabbei, P. bulbosa and P. azarasi (Paggi et al., 2000). Similar techniques were used by Mattiucci et al. (2002) for differentiation between Anisakis typica and other species within the genus.

Impact: Economic

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In some cases, parasitic nematodes can produce very spectacular and massive infections in fishes. The masses of Philometra abdominalis (Fig. 1.) in the abdominal cavity of gudgeon or the white-coloured Hysterothylacium bidentatum (Fig. 2.) in the stomach of sterlet (Acipenser ruthenus) definitely indicate that nematodes are important pathogens. In spite of this, Williams (1967) rightly stated that among the helminth parasites of fishes the pathological effect of nematodes had been studied the least. The situation has not changed. Although the number of known species has gradually increased, there are still few new data on the pathological effects of nematodes. In freshwater fish species, the new data mostly concern Anguillicola crassus (Fig. 3) infection in eel (Anguilla anguilla). As regards marine fish parasites, the anisakid species (Anisakis, Contracaecum, Hysterothylacium, Pseudoterranova) have come into prominence because of infections in humans.

Fish nematodes might harm their host in a variety of ways. They can cause mechanical injuries, atrophy of tissues, occlusion of the alimentary canal, blood vessels and other ducts and toxication from their metabolic products, and they can deprive the host of food, enzymes and vitamins.

Fig. 1. Philometra abdominalis filling the abdominal cavity of a gudgeon (Gobio gobio).

Fig. 2. Hysterothylacium bidentatum specimens in the stomach of the sterlet (Acipenser ruthenus).

Fig. 3. Heavy and moderate infections with Anguillicola crassus in opened swim bladders of European eels.

Zoonoses and Food Safety

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The presence of nematode larvae in fish may present a risk for humans. Third-stage larvae of Anisakis spp., Pseudoterranova spp., Phocascaris spp. and Contracaecum spp. occur in fish musculature and their adult stage is in homeothermic animals such as whales and seals. Thus, the stimuli for further development of the larva to the final moult are high temperature, acidic pH and pepsinogen, which occur in the stomach of homeothermic animals. Following ingestion by humans of live larvae in fish products, the larvae get activated and penetrate the gastric or intestinal mucosa, eliciting abdominal symptoms. Extra-intestinal migration of worms also occurs. This is anisakiasis (anisakiosis) or pseudoterranoviasis (pseudoterranovosis) (Smith and Wootten, 1978; Moller and Anders, 1986; McClelland, 2002).

Freezing (-20°C for 24 h), heating above 60°C or salting (250 g NaCl/l) will kill Anisakis larvae. Likewise, storage of Pseudoterranova larvae in fish meat at 30°C for 15 h or at 20°C for 7 days is lethal for the worms (McClelland, 2002). Even dead worms (killed by freezing, heating or salting) still contain immunogenic molecules, which can trigger severe allergic reactions following ingestion of fish products. In fact, the disease provoked by the marine larvae is to a high degree associated with the host reaction, such as immunoglobulin E (IgE) production, mast-cell degranulation, eosinophilia, oedema and urticaria. It has been speculated that nematodes living in the adult stage in fish, such as Hysterothylacium, could elicit disease in humans following ingestion of live larvae (Norris and Overstreet, 1976). It has been reported that adult nematodes of Philometra from fish tissue can infect humans through open wounds in skin and elicit disease (Deardorff et al., 1986). Considering the high prevalence of philometrids in fishes (Moravec et al., 2003), this suggestion should be further studied.

Spoilage of fish products by marine nematode larvae

Nematode larvae in fish fillets may lead to rejection by consumers due to the unattractive appearance of the product. Therefore considerable efforts are being made to detect and remove larval nematodes from fish products (Moller and Anders, 1986). It has been estimated that up to half of the production costs in certain processing plants consists in the detection and removal of Pseudoterranova from cod fillets; the problem is worldwide due to the extensive distribution of these anisakid larvae (McClelland, 2002). These concerns are mainly with wild fishes because several studies have shown that aquacultured salmon are free from infection, since in this case fish conduct their entire life in captivity and are fed by non-infected feed (Lunestad, 2003).

References

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