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Digenean infections of fish

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Digenean infections of fish

Summary

  • Last modified
  • 09 November 2017
  • Datasheet Type(s)
  • Animal Disease
  • Preferred Scientific Name
  • Digenean infections of fish
  • Overview
  • The Digenea (previously termed digenetic trematodes) are one of the three major taxa of parasitic Platyhelminthes, the other two being the Cestoda and the Monogenea. Digeneans are heteroxenous (i.e. they require more th...

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A. Pirenella conica (actual size 15 mm). B. Bulinus truncatus (actual size 6-11 mm).
TitleMolluscs. Pirenella conica and Bulinus truncatus
CaptionA. Pirenella conica (actual size 15 mm). B. Bulinus truncatus (actual size 6-11 mm).
CopyrightIlan Paperna & Ronald Dzikowski
A. Pirenella conica (actual size 15 mm). B. Bulinus truncatus (actual size 6-11 mm).
Molluscs. Pirenella conica and Bulinus truncatusA. Pirenella conica (actual size 15 mm). B. Bulinus truncatus (actual size 6-11 mm).Ilan Paperna & Ronald Dzikowski
Melanoides tuberculata (actual size 20 mm).
TitleMelanoides tuberculata. Molluscs
CaptionMelanoides tuberculata (actual size 20 mm).
CopyrightIlan Paperna & Ronald Dzikowski
Melanoides tuberculata (actual size 20 mm).
Melanoides tuberculata. MolluscsMelanoides tuberculata (actual size 20 mm).Ilan Paperna & Ronald Dzikowski
Examples of a 'distome' digenean (A) Allocreadium isoporum (drawn with reference to Dawes, 1946, actual size 3-5 mm) and a 'monostome' digenean (B) Bucephalopsis gracilescens (Bucephalidae, drawn with reference to Dawes, 1946 and Bychowskaya-Pavlovskaya et al., 1962, actual size 6 mm). Abbreviations: a = acetabulum; cs = cirrus pouch (sac); dl = ductus laureri; g = Mehlis glands; gp = genital opening; in = intestine; o = ovary; od = oviduct; os = oral sucker; ot = ootype; ph = pharynx; pr = prostate; s = sucker; t = testis; vd = vas deferens; vid = vitelline ducts; vs = seminal vesicle; vt = vitellaria; x = excretory ducts/bladder.
TitleExamples of a 'distome' digenean
CaptionExamples of a 'distome' digenean (A) Allocreadium isoporum (drawn with reference to Dawes, 1946, actual size 3-5 mm) and a 'monostome' digenean (B) Bucephalopsis gracilescens (Bucephalidae, drawn with reference to Dawes, 1946 and Bychowskaya-Pavlovskaya et al., 1962, actual size 6 mm). Abbreviations: a = acetabulum; cs = cirrus pouch (sac); dl = ductus laureri; g = Mehlis glands; gp = genital opening; in = intestine; o = ovary; od = oviduct; os = oral sucker; ot = ootype; ph = pharynx; pr = prostate; s = sucker; t = testis; vd = vas deferens; vid = vitelline ducts; vs = seminal vesicle; vt = vitellaria; x = excretory ducts/bladder.
CopyrightIlan Paperna & Ronald Dzikowski
Examples of a 'distome' digenean (A) Allocreadium isoporum (drawn with reference to Dawes, 1946, actual size 3-5 mm) and a 'monostome' digenean (B) Bucephalopsis gracilescens (Bucephalidae, drawn with reference to Dawes, 1946 and Bychowskaya-Pavlovskaya et al., 1962, actual size 6 mm). Abbreviations: a = acetabulum; cs = cirrus pouch (sac); dl = ductus laureri; g = Mehlis glands; gp = genital opening; in = intestine; o = ovary; od = oviduct; os = oral sucker; ot = ootype; ph = pharynx; pr = prostate; s = sucker; t = testis; vd = vas deferens; vid = vitelline ducts; vs = seminal vesicle; vt = vitellaria; x = excretory ducts/bladder.
Examples of a 'distome' digeneanExamples of a 'distome' digenean (A) Allocreadium isoporum (drawn with reference to Dawes, 1946, actual size 3-5 mm) and a 'monostome' digenean (B) Bucephalopsis gracilescens (Bucephalidae, drawn with reference to Dawes, 1946 and Bychowskaya-Pavlovskaya et al., 1962, actual size 6 mm). Abbreviations: a = acetabulum; cs = cirrus pouch (sac); dl = ductus laureri; g = Mehlis glands; gp = genital opening; in = intestine; o = ovary; od = oviduct; os = oral sucker; ot = ootype; ph = pharynx; pr = prostate; s = sucker; t = testis; vd = vas deferens; vid = vitelline ducts; vs = seminal vesicle; vt = vitellaria; x = excretory ducts/bladder.Ilan Paperna & Ronald Dzikowski
Scanning electron micrographs of adult-stage Haplorchis pumilio (A) and Pygidiopsis geneta (B) from the gut of a cormorant (Phalacrocorax carbo), scales 50 µm.
TitleAdult stage trematodes
CaptionScanning electron micrographs of adult-stage Haplorchis pumilio (A) and Pygidiopsis geneta (B) from the gut of a cormorant (Phalacrocorax carbo), scales 50 µm.
CopyrightIlan Paperna & Ronald Dzikowski
Scanning electron micrographs of adult-stage Haplorchis pumilio (A) and Pygidiopsis geneta (B) from the gut of a cormorant (Phalacrocorax carbo), scales 50 µm.
Adult stage trematodesScanning electron micrographs of adult-stage Haplorchis pumilio (A) and Pygidiopsis geneta (B) from the gut of a cormorant (Phalacrocorax carbo), scales 50 µm.Ilan Paperna & Ronald Dzikowski
A. Clinostomum tilapiae eggs containing ready-to-hatch miracidium (actual size 125 µm x 83 µm). B. Redia of Mesorchis denticulatum (scanning electron micrograph, scale 10 µm, courtesy of M. Køie). C. Redia of C. tilapiae (live, actual size). D. Furcocercaria of C. tilapiae (live, actual size).
TitleEggs, rediae, furcocercaria
CaptionA. Clinostomum tilapiae eggs containing ready-to-hatch miracidium (actual size 125 µm x 83 µm). B. Redia of Mesorchis denticulatum (scanning electron micrograph, scale 10 µm, courtesy of M. Køie). C. Redia of C. tilapiae (live, actual size). D. Furcocercaria of C. tilapiae (live, actual size).
CopyrightIlan Paperna & Ronald Dzikowski
A. Clinostomum tilapiae eggs containing ready-to-hatch miracidium (actual size 125 µm x 83 µm). B. Redia of Mesorchis denticulatum (scanning electron micrograph, scale 10 µm, courtesy of M. Køie). C. Redia of C. tilapiae (live, actual size). D. Furcocercaria of C. tilapiae (live, actual size).
Eggs, rediae, furcocercariaA. Clinostomum tilapiae eggs containing ready-to-hatch miracidium (actual size 125 µm x 83 µm). B. Redia of Mesorchis denticulatum (scanning electron micrograph, scale 10 µm, courtesy of M. Køie). C. Redia of C. tilapiae (live, actual size). D. Furcocercaria of C. tilapiae (live, actual size).Ilan Paperna & Ronald Dzikowski
Sanguinicola dentata (Paperna, 1964) from Clarias gariepinus circulatory system (eggs are released into the kidneys). Actual size 3.4 mm, Abbreviations: EX = excretory duct; GP = genital pore; M = Mehlis glands; O = ovary; OD = oviduct; T = testis; U = uterus; V = vitellaria; VD = vas deferens; VID vitelline duct.
TitleCirculatory system
CaptionSanguinicola dentata (Paperna, 1964) from Clarias gariepinus circulatory system (eggs are released into the kidneys). Actual size 3.4 mm, Abbreviations: EX = excretory duct; GP = genital pore; M = Mehlis glands; O = ovary; OD = oviduct; T = testis; U = uterus; V = vitellaria; VD = vas deferens; VID vitelline duct.
CopyrightIlan Paperna & Ronald Dzikowski
Sanguinicola dentata (Paperna, 1964) from Clarias gariepinus circulatory system (eggs are released into the kidneys). Actual size 3.4 mm, Abbreviations: EX = excretory duct; GP = genital pore; M = Mehlis glands; O = ovary; OD = oviduct; T = testis; U = uterus; V = vitellaria; VD = vas deferens; VID vitelline duct.
Circulatory systemSanguinicola dentata (Paperna, 1964) from Clarias gariepinus circulatory system (eggs are released into the kidneys). Actual size 3.4 mm, Abbreviations: EX = excretory duct; GP = genital pore; M = Mehlis glands; O = ovary; OD = oviduct; T = testis; U = uterus; V = vitellaria; VD = vas deferens; VID vitelline duct.Ilan Paperna & Ronald Dzikowski
Blood flukes. A. Scanning electron micrograph of Sanguinicola fontianalis (from Hoffman et al., 1985, courtesy of the author; bar = 100 µm). B. Eggs of Sanguinicola sp. in gills of Oreochromis aureus, Lake Kinnereth. C. Egg in gill tissue of Baryanchistus sp. (Plecostomidae), Amazon, Brazil. D. Eggs in spleen of O. aurea, Lake Kinnereth.
TitleBlood flukes
CaptionBlood flukes. A. Scanning electron micrograph of Sanguinicola fontianalis (from Hoffman et al., 1985, courtesy of the author; bar = 100 µm). B. Eggs of Sanguinicola sp. in gills of Oreochromis aureus, Lake Kinnereth. C. Egg in gill tissue of Baryanchistus sp. (Plecostomidae), Amazon, Brazil. D. Eggs in spleen of O. aurea, Lake Kinnereth.
CopyrightIlan Paperna & Ronald Dzikowski
Blood flukes. A. Scanning electron micrograph of Sanguinicola fontianalis (from Hoffman et al., 1985, courtesy of the author; bar = 100 µm). B. Eggs of Sanguinicola sp. in gills of Oreochromis aureus, Lake Kinnereth. C. Egg in gill tissue of Baryanchistus sp. (Plecostomidae), Amazon, Brazil. D. Eggs in spleen of O. aurea, Lake Kinnereth.
Blood flukesBlood flukes. A. Scanning electron micrograph of Sanguinicola fontianalis (from Hoffman et al., 1985, courtesy of the author; bar = 100 µm). B. Eggs of Sanguinicola sp. in gills of Oreochromis aureus, Lake Kinnereth. C. Egg in gill tissue of Baryanchistus sp. (Plecostomidae), Amazon, Brazil. D. Eggs in spleen of O. aurea, Lake Kinnereth.Ilan Paperna & Ronald Dzikowski
Didymozoidae. A. Palate of Platycephalus fuscus infected with Neometadidymozoon helicis (in life, bright yellow). From Lester, 1980, courtesy of the author. B. Lobatozoum multidacculatum on gills of Katsumonus pelamis, New Zealand. C. Larval stages of didymozoids encysted on the surface of the intestine of Favonigobius exquisitus. D. Section of Nematobothrium spinneri in the body wall muscle of Acanthocybium soalndri, Queensland. E. N. helicis capsule in the body wall of P. fuscus. (B-E courtesy of B. Lester.)
TitleDidymozoidae
CaptionDidymozoidae. A. Palate of Platycephalus fuscus infected with Neometadidymozoon helicis (in life, bright yellow). From Lester, 1980, courtesy of the author. B. Lobatozoum multidacculatum on gills of Katsumonus pelamis, New Zealand. C. Larval stages of didymozoids encysted on the surface of the intestine of Favonigobius exquisitus. D. Section of Nematobothrium spinneri in the body wall muscle of Acanthocybium soalndri, Queensland. E. N. helicis capsule in the body wall of P. fuscus. (B-E courtesy of B. Lester.)
CopyrightIlan Paperna & Ronald Dzikowski
Didymozoidae. A. Palate of Platycephalus fuscus infected with Neometadidymozoon helicis (in life, bright yellow). From Lester, 1980, courtesy of the author. B. Lobatozoum multidacculatum on gills of Katsumonus pelamis, New Zealand. C. Larval stages of didymozoids encysted on the surface of the intestine of Favonigobius exquisitus. D. Section of Nematobothrium spinneri in the body wall muscle of Acanthocybium soalndri, Queensland. E. N. helicis capsule in the body wall of P. fuscus. (B-E courtesy of B. Lester.)
DidymozoidaeDidymozoidae. A. Palate of Platycephalus fuscus infected with Neometadidymozoon helicis (in life, bright yellow). From Lester, 1980, courtesy of the author. B. Lobatozoum multidacculatum on gills of Katsumonus pelamis, New Zealand. C. Larval stages of didymozoids encysted on the surface of the intestine of Favonigobius exquisitus. D. Section of Nematobothrium spinneri in the body wall muscle of Acanthocybium soalndri, Queensland. E. N. helicis capsule in the body wall of P. fuscus. (B-E courtesy of B. Lester.)Ilan Paperna & Ronald Dzikowski
Didymozoidae. A. Didymozoon faciale (actual size 16.3 mm). B. Neodiplotrema pelamydis - fused pair (drawn with reference to Dawes, 1946 (A) and Yamaguti, 1958 (B); for abbreviations see legend to Fig. 10.3).
TitleDidymozoidae
CaptionDidymozoidae. A. Didymozoon faciale (actual size 16.3 mm). B. Neodiplotrema pelamydis - fused pair (drawn with reference to Dawes, 1946 (A) and Yamaguti, 1958 (B); for abbreviations see legend to Fig. 10.3).
CopyrightIlan Paperna & Ronald Dzikowski
Didymozoidae. A. Didymozoon faciale (actual size 16.3 mm). B. Neodiplotrema pelamydis - fused pair (drawn with reference to Dawes, 1946 (A) and Yamaguti, 1958 (B); for abbreviations see legend to Fig. 10.3).
DidymozoidaeDidymozoidae. A. Didymozoon faciale (actual size 16.3 mm). B. Neodiplotrema pelamydis - fused pair (drawn with reference to Dawes, 1946 (A) and Yamaguti, 1958 (B); for abbreviations see legend to Fig. 10.3).Ilan Paperna & Ronald Dzikowski
Transversotrema haasi (drawn with reference to Witenberg, 1944a; for abbreviations see legend to Fig. 10.3).
TitleTransversotrema haasi
CaptionTransversotrema haasi (drawn with reference to Witenberg, 1944a; for abbreviations see legend to Fig. 10.3).
CopyrightIlan Paperna & Ronald Dzikowski
Transversotrema haasi (drawn with reference to Witenberg, 1944a; for abbreviations see legend to Fig. 10.3).
Transversotrema haasiTransversotrema haasi (drawn with reference to Witenberg, 1944a; for abbreviations see legend to Fig. 10.3).Ilan Paperna & Ronald Dzikowski
A. Bolbophorus levantinum daughter sporocyst with cercariae, characteristic of strigeoid and schistosomatid digeneans (redrawn from Paperna and Lengy, 1963). B.Young and C, mature daughter redia containing cercariae of Clinostomum tilapiae. Abbreviations: b = birth pore; c = cercariae; gr = germinating stages; in = intestine; os = oral sucker.
TitleDigenean cercariae
CaptionA. Bolbophorus levantinum daughter sporocyst with cercariae, characteristic of strigeoid and schistosomatid digeneans (redrawn from Paperna and Lengy, 1963). B.Young and C, mature daughter redia containing cercariae of Clinostomum tilapiae. Abbreviations: b = birth pore; c = cercariae; gr = germinating stages; in = intestine; os = oral sucker.
CopyrightIlan Paperna & Ronald Dzikowski
A. Bolbophorus levantinum daughter sporocyst with cercariae, characteristic of strigeoid and schistosomatid digeneans (redrawn from Paperna and Lengy, 1963). B.Young and C, mature daughter redia containing cercariae of Clinostomum tilapiae. Abbreviations: b = birth pore; c = cercariae; gr = germinating stages; in = intestine; os = oral sucker.
Digenean cercariaeA. Bolbophorus levantinum daughter sporocyst with cercariae, characteristic of strigeoid and schistosomatid digeneans (redrawn from Paperna and Lengy, 1963). B.Young and C, mature daughter redia containing cercariae of Clinostomum tilapiae. Abbreviations: b = birth pore; c = cercariae; gr = germinating stages; in = intestine; os = oral sucker.Ilan Paperna & Ronald Dzikowski
Transmission electron micrographs of Centrocestus sp. metacercariae from gills of Oreochromis aureus. A. View of the host-produced cartilaginous cyst (c), the parasite-produced wall (pw) enclosing the parasite (p) with the spiny tegument (s). B. Parasite's wall (pw) and the syncytial tegument. C. Enlarged view of the tegument with its spine-carrying border. Abbreviations: cl = circular muscles; ll = longitudinal muscles; m = mitochondrion; n = nuclei of the tegument; s = spines.
TitleCentrocestus sp.
CaptionTransmission electron micrographs of Centrocestus sp. metacercariae from gills of Oreochromis aureus. A. View of the host-produced cartilaginous cyst (c), the parasite-produced wall (pw) enclosing the parasite (p) with the spiny tegument (s). B. Parasite's wall (pw) and the syncytial tegument. C. Enlarged view of the tegument with its spine-carrying border. Abbreviations: cl = circular muscles; ll = longitudinal muscles; m = mitochondrion; n = nuclei of the tegument; s = spines.
CopyrightIlan Paperna & Ronald Dzikowski
Transmission electron micrographs of Centrocestus sp. metacercariae from gills of Oreochromis aureus. A. View of the host-produced cartilaginous cyst (c), the parasite-produced wall (pw) enclosing the parasite (p) with the spiny tegument (s). B. Parasite's wall (pw) and the syncytial tegument. C. Enlarged view of the tegument with its spine-carrying border. Abbreviations: cl = circular muscles; ll = longitudinal muscles; m = mitochondrion; n = nuclei of the tegument; s = spines.
Centrocestus sp.Transmission electron micrographs of Centrocestus sp. metacercariae from gills of Oreochromis aureus. A. View of the host-produced cartilaginous cyst (c), the parasite-produced wall (pw) enclosing the parasite (p) with the spiny tegument (s). B. Parasite's wall (pw) and the syncytial tegument. C. Enlarged view of the tegument with its spine-carrying border. Abbreviations: cl = circular muscles; ll = longitudinal muscles; m = mitochondrion; n = nuclei of the tegument; s = spines.Ilan Paperna & Ronald Dzikowski
Scanning electron micrographs of adult-stage trematodes from cormorant's gut: A. Paryphostomum radiatum (Echinostomatidae) with perioral spines and smooth tegument, scale bar 100 µm. B. Centrocestus sp. (Heterophyiidae) with perioral and tegumental spines, scale bar 10 µm.
TitleAdult stage trematodes
CaptionScanning electron micrographs of adult-stage trematodes from cormorant's gut: A. Paryphostomum radiatum (Echinostomatidae) with perioral spines and smooth tegument, scale bar 100 µm. B. Centrocestus sp. (Heterophyiidae) with perioral and tegumental spines, scale bar 10 µm.
CopyrightIlan Paperna & Ronald Dzikowski
Scanning electron micrographs of adult-stage trematodes from cormorant's gut: A. Paryphostomum radiatum (Echinostomatidae) with perioral spines and smooth tegument, scale bar 100 µm. B. Centrocestus sp. (Heterophyiidae) with perioral and tegumental spines, scale bar 10 µm.
Adult stage trematodesScanning electron micrographs of adult-stage trematodes from cormorant's gut: A. Paryphostomum radiatum (Echinostomatidae) with perioral spines and smooth tegument, scale bar 100 µm. B. Centrocestus sp. (Heterophyiidae) with perioral and tegumental spines, scale bar 10 µm.Ilan Paperna & Ronald Dzikowski
Eggs of a sanguinicolid in gills of juvenile Liza sp., Kowie estuary, south-east Cape, South Africa.
TitleEggs of a sanguinicolid
CaptionEggs of a sanguinicolid in gills of juvenile Liza sp., Kowie estuary, south-east Cape, South Africa.
CopyrightIlan Paperna & Ronald Dzikowski
Eggs of a sanguinicolid in gills of juvenile Liza sp., Kowie estuary, south-east Cape, South Africa.
Eggs of a sanguinicolidEggs of a sanguinicolid in gills of juvenile Liza sp., Kowie estuary, south-east Cape, South Africa.Ilan Paperna & Ronald Dzikowski
Degenerate eggs of the sanguinicolid Pearsonellum corventum in the heart of the fish Plectropomus leopardus. A. Encapsulated egg mass enclosed by infiltrated macrophages. B. Granuloma encased egg residue within a melanomacrophage. (From Overstreet and Thulin, 1989, courtesy of R. Overstreet.)|Degenerate eggs of the sanguinicolis Pearsonellum corventum in the heart of the fish Plectropomus leopardus. A. Encapsulated egg mass enclosed by infiltrated macrophages. B. Granuloma encased egg residue within a melanomacrophage. (From Overstreet and Thulin, 1989, courtesy of R. Overstreet.)
TitleDegenerate sanguinicolid eggs
CaptionDegenerate eggs of the sanguinicolid Pearsonellum corventum in the heart of the fish Plectropomus leopardus. A. Encapsulated egg mass enclosed by infiltrated macrophages. B. Granuloma encased egg residue within a melanomacrophage. (From Overstreet and Thulin, 1989, courtesy of R. Overstreet.)|Degenerate eggs of the sanguinicolis Pearsonellum corventum in the heart of the fish Plectropomus leopardus. A. Encapsulated egg mass enclosed by infiltrated macrophages. B. Granuloma encased egg residue within a melanomacrophage. (From Overstreet and Thulin, 1989, courtesy of R. Overstreet.)
CopyrightIlan Paperna & Ronald Dzikowski
Degenerate eggs of the sanguinicolid Pearsonellum corventum in the heart of the fish Plectropomus leopardus. A. Encapsulated egg mass enclosed by infiltrated macrophages. B. Granuloma encased egg residue within a melanomacrophage. (From Overstreet and Thulin, 1989, courtesy of R. Overstreet.)|Degenerate eggs of the sanguinicolis Pearsonellum corventum in the heart of the fish Plectropomus leopardus. A. Encapsulated egg mass enclosed by infiltrated macrophages. B. Granuloma encased egg residue within a melanomacrophage. (From Overstreet and Thulin, 1989, courtesy of R. Overstreet.)
Degenerate sanguinicolid eggsDegenerate eggs of the sanguinicolid Pearsonellum corventum in the heart of the fish Plectropomus leopardus. A. Encapsulated egg mass enclosed by infiltrated macrophages. B. Granuloma encased egg residue within a melanomacrophage. (From Overstreet and Thulin, 1989, courtesy of R. Overstreet.)|Degenerate eggs of the sanguinicolis Pearsonellum corventum in the heart of the fish Plectropomus leopardus. A. Encapsulated egg mass enclosed by infiltrated macrophages. B. Granuloma encased egg residue within a melanomacrophage. (From Overstreet and Thulin, 1989, courtesy of R. Overstreet.)Ilan Paperna & Ronald Dzikowski
Metacercariae in fish tissues. A. Cartilaginous cyst around Centrocestus sp. metacercariae in gills of Oreochromis aurea, Lake Kinnereth. B. Ascocotyle coelostoma (live) in the truncus arteriosus of Liza ramada. C. Encysted Phagicola nana in the gut wall of Micropterus salmoides. D. Pigmented Neascus ('black spot') encysted on fins of young Oreochromis hybrids.
TitleMetacercariae in fish tissues
CaptionMetacercariae in fish tissues. A. Cartilaginous cyst around Centrocestus sp. metacercariae in gills of Oreochromis aurea, Lake Kinnereth. B. Ascocotyle coelostoma (live) in the truncus arteriosus of Liza ramada. C. Encysted Phagicola nana in the gut wall of Micropterus salmoides. D. Pigmented Neascus ('black spot') encysted on fins of young Oreochromis hybrids.
CopyrightIlan Paperna & Ronald Dzikowski
Metacercariae in fish tissues. A. Cartilaginous cyst around Centrocestus sp. metacercariae in gills of Oreochromis aurea, Lake Kinnereth. B. Ascocotyle coelostoma (live) in the truncus arteriosus of Liza ramada. C. Encysted Phagicola nana in the gut wall of Micropterus salmoides. D. Pigmented Neascus ('black spot') encysted on fins of young Oreochromis hybrids.
Metacercariae in fish tissuesMetacercariae in fish tissues. A. Cartilaginous cyst around Centrocestus sp. metacercariae in gills of Oreochromis aurea, Lake Kinnereth. B. Ascocotyle coelostoma (live) in the truncus arteriosus of Liza ramada. C. Encysted Phagicola nana in the gut wall of Micropterus salmoides. D. Pigmented Neascus ('black spot') encysted on fins of young Oreochromis hybrids.Ilan Paperna & Ronald Dzikowski
Diplostomatid metacercariae: A. Diplostomum sp. removed from a visceral cyst in Clarias gariepinus, Uganda. B. Heavy Bolbophorus levantinum metacercaria in muscles of young Oreochromis aureus from Lake Kinnereth. C. Early-stage B. levantinum metacercaria with expanded posterior half. D. Later-stage B. levantinum metacercaria with emptied posterior end (actual size 0.6-0.8 mm).
TitleDiplostomatid metacercariae
CaptionDiplostomatid metacercariae: A. Diplostomum sp. removed from a visceral cyst in Clarias gariepinus, Uganda. B. Heavy Bolbophorus levantinum metacercaria in muscles of young Oreochromis aureus from Lake Kinnereth. C. Early-stage B. levantinum metacercaria with expanded posterior half. D. Later-stage B. levantinum metacercaria with emptied posterior end (actual size 0.6-0.8 mm).
CopyrightIlan Paperna & Ronald Dzikowski
Diplostomatid metacercariae: A. Diplostomum sp. removed from a visceral cyst in Clarias gariepinus, Uganda. B. Heavy Bolbophorus levantinum metacercaria in muscles of young Oreochromis aureus from Lake Kinnereth. C. Early-stage B. levantinum metacercaria with expanded posterior half. D. Later-stage B. levantinum metacercaria with emptied posterior end (actual size 0.6-0.8 mm).
Diplostomatid metacercariaeDiplostomatid metacercariae: A. Diplostomum sp. removed from a visceral cyst in Clarias gariepinus, Uganda. B. Heavy Bolbophorus levantinum metacercaria in muscles of young Oreochromis aureus from Lake Kinnereth. C. Early-stage B. levantinum metacercaria with expanded posterior half. D. Later-stage B. levantinum metacercaria with emptied posterior end (actual size 0.6-0.8 mm).Ilan Paperna & Ronald Dzikowski
Histopathology of Bolbophorus levantinum infection in young Oreochromis aureus. A. Macrophage infiltration around young unencysted metacercaria. B. Maturing metacercaria with 'reserve cells' (r) enclosed in a parasite-derived wall (w) and foci of macrophage proliferations in the surrounding muscles (arrows).
TitleHistopathology of Bolbophorus levantinum infection
CaptionHistopathology of Bolbophorus levantinum infection in young Oreochromis aureus. A. Macrophage infiltration around young unencysted metacercaria. B. Maturing metacercaria with 'reserve cells' (r) enclosed in a parasite-derived wall (w) and foci of macrophage proliferations in the surrounding muscles (arrows).
CopyrightIlan Paperna & Ronald Dzikowski
Histopathology of Bolbophorus levantinum infection in young Oreochromis aureus. A. Macrophage infiltration around young unencysted metacercaria. B. Maturing metacercaria with 'reserve cells' (r) enclosed in a parasite-derived wall (w) and foci of macrophage proliferations in the surrounding muscles (arrows).
Histopathology of Bolbophorus levantinum infectionHistopathology of Bolbophorus levantinum infection in young Oreochromis aureus. A. Macrophage infiltration around young unencysted metacercaria. B. Maturing metacercaria with 'reserve cells' (r) enclosed in a parasite-derived wall (w) and foci of macrophage proliferations in the surrounding muscles (arrows).Ilan Paperna & Ronald Dzikowski
Metacercarial skin infection in Lake Kinnereth cichlids. A, B. Low and heavy 'black spot' (Neascus) infection in young pond-reared Oreochromis aureus x nilotica.
TitleMetacercarial skin infection
CaptionMetacercarial skin infection in Lake Kinnereth cichlids. A, B. Low and heavy 'black spot' (Neascus) infection in young pond-reared Oreochromis aureus x nilotica.
CopyrightIlan Paperna & Ronald Dzikowski
Metacercarial skin infection in Lake Kinnereth cichlids. A, B. Low and heavy 'black spot' (Neascus) infection in young pond-reared Oreochromis aureus x nilotica.
Metacercarial skin infectionMetacercarial skin infection in Lake Kinnereth cichlids. A, B. Low and heavy 'black spot' (Neascus) infection in young pond-reared Oreochromis aureus x nilotica.Ilan Paperna & Ronald Dzikowski
Metacercarial skin infection in Lake Kinnereth cichlids. Clinostomum 'cutaneus'-infected Tristramella simonis (top) and Tilapia zilli (bottom).
TitleMetacercarial skin infection
CaptionMetacercarial skin infection in Lake Kinnereth cichlids. Clinostomum 'cutaneus'-infected Tristramella simonis (top) and Tilapia zilli (bottom).
CopyrightIlan Paperna & Ronald Dzikowski
Metacercarial skin infection in Lake Kinnereth cichlids. Clinostomum 'cutaneus'-infected Tristramella simonis (top) and Tilapia zilli (bottom).
Metacercarial skin infectionMetacercarial skin infection in Lake Kinnereth cichlids. Clinostomum 'cutaneus'-infected Tristramella simonis (top) and Tilapia zilli (bottom).Ilan Paperna & Ronald Dzikowski
Abdominal dropsy in Posthodiplostomum minimum-infected Pimephales promelas (43 mm long) (after Mitchell et al., 1982, courtesy of G.L. Hoffman).
TitleAbdominal dropsy
CaptionAbdominal dropsy in Posthodiplostomum minimum-infected Pimephales promelas (43 mm long) (after Mitchell et al., 1982, courtesy of G.L. Hoffman).
CopyrightIlan Paperna & Ronald Dzikowski
Abdominal dropsy in Posthodiplostomum minimum-infected Pimephales promelas (43 mm long) (after Mitchell et al., 1982, courtesy of G.L. Hoffman).
Abdominal dropsyAbdominal dropsy in Posthodiplostomum minimum-infected Pimephales promelas (43 mm long) (after Mitchell et al., 1982, courtesy of G.L. Hoffman).Ilan Paperna & Ronald Dzikowski
Eye diplostomiasis in Ictalurus melas. A. Cross-section of the eye showing herniation (h) of the lens (l) and the metacercariae inside (arrows). B. Damaged lens (l) with proliferated epithelium around a necrotic core (i); note vascularization of the vitreous humour (arrow). (After Larson, 1965, courtesy of the author.)|Eye diplostomiasis in Ictalurus melas. A. Cross-section of the eye showing herniation (h) of the lens (l) and the metacercariae inside (arrows). B. Damaged lens (l) with proliferated epithelium around a necrotic core (i); note vascularization of the vitreous humour (arrow). C, D. Cross-section of herniated lenses; empty spaces contained worms before processing. (After Larson, 1965, courtesy of the author.)
TitleEye diplostomiasis
CaptionEye diplostomiasis in Ictalurus melas. A. Cross-section of the eye showing herniation (h) of the lens (l) and the metacercariae inside (arrows). B. Damaged lens (l) with proliferated epithelium around a necrotic core (i); note vascularization of the vitreous humour (arrow). (After Larson, 1965, courtesy of the author.)|Eye diplostomiasis in Ictalurus melas. A. Cross-section of the eye showing herniation (h) of the lens (l) and the metacercariae inside (arrows). B. Damaged lens (l) with proliferated epithelium around a necrotic core (i); note vascularization of the vitreous humour (arrow). C, D. Cross-section of herniated lenses; empty spaces contained worms before processing. (After Larson, 1965, courtesy of the author.)
CopyrightIlan Paperna & Ronald Dzikowski
Eye diplostomiasis in Ictalurus melas. A. Cross-section of the eye showing herniation (h) of the lens (l) and the metacercariae inside (arrows). B. Damaged lens (l) with proliferated epithelium around a necrotic core (i); note vascularization of the vitreous humour (arrow). (After Larson, 1965, courtesy of the author.)|Eye diplostomiasis in Ictalurus melas. A. Cross-section of the eye showing herniation (h) of the lens (l) and the metacercariae inside (arrows). B. Damaged lens (l) with proliferated epithelium around a necrotic core (i); note vascularization of the vitreous humour (arrow). C, D. Cross-section of herniated lenses; empty spaces contained worms before processing. (After Larson, 1965, courtesy of the author.)
Eye diplostomiasisEye diplostomiasis in Ictalurus melas. A. Cross-section of the eye showing herniation (h) of the lens (l) and the metacercariae inside (arrows). B. Damaged lens (l) with proliferated epithelium around a necrotic core (i); note vascularization of the vitreous humour (arrow). (After Larson, 1965, courtesy of the author.)|Eye diplostomiasis in Ictalurus melas. A. Cross-section of the eye showing herniation (h) of the lens (l) and the metacercariae inside (arrows). B. Damaged lens (l) with proliferated epithelium around a necrotic core (i); note vascularization of the vitreous humour (arrow). C, D. Cross-section of herniated lenses; empty spaces contained worms before processing. (After Larson, 1965, courtesy of the author.)Ilan Paperna & Ronald Dzikowski
Eye diplostomiasis in Ictalurus melas. C, D. Cross-section of herniated lenses; empty spaces contained worms before processing. (After Larson, 1965, courtesy of the author.)
TitleEye diplostomiasis
CaptionEye diplostomiasis in Ictalurus melas. C, D. Cross-section of herniated lenses; empty spaces contained worms before processing. (After Larson, 1965, courtesy of the author.)
CopyrightIlan Paperna & Ronald Dzikowski
Eye diplostomiasis in Ictalurus melas. C, D. Cross-section of herniated lenses; empty spaces contained worms before processing. (After Larson, 1965, courtesy of the author.)
Eye diplostomiasisEye diplostomiasis in Ictalurus melas. C, D. Cross-section of herniated lenses; empty spaces contained worms before processing. (After Larson, 1965, courtesy of the author.)Ilan Paperna & Ronald Dzikowski
Clonorchis sinensis whole worm (original, x 6).
TitleClonorchis sinensis worm
CaptionClonorchis sinensis whole worm (original, x 6).
CopyrightRonald C. Ko
Clonorchis sinensis whole worm (original, x 6).
Clonorchis sinensis wormClonorchis sinensis whole worm (original, x 6).Ronald C. Ko
Egg of Clonorchis sinensis recovered from faeces (original).
TitleEgg of Clonorchis sinensis
CaptionEgg of Clonorchis sinensis recovered from faeces (original).
CopyrightRonald C. Ko
Egg of Clonorchis sinensis recovered from faeces (original).
Egg of Clonorchis sinensisEgg of Clonorchis sinensis recovered from faeces (original).Ronald C. Ko
Metacercariae of Clonorchis sinensis (original).
TitleMetacercariae of Clonorchis sinensis
CaptionMetacercariae of Clonorchis sinensis (original).
CopyrightRonald C. Ko
Metacercariae of Clonorchis sinensis (original).
Metacercariae of Clonorchis sinensisMetacercariae of Clonorchis sinensis (original).Ronald C. Ko
A carp pond in Hong Kong, showing an outside toilet over pond (original).
TitleCarp pond
CaptionA carp pond in Hong Kong, showing an outside toilet over pond (original).
CopyrightRonald C. Ko
A carp pond in Hong Kong, showing an outside toilet over pond (original).
Carp pondA carp pond in Hong Kong, showing an outside toilet over pond (original).Ronald C. Ko

Identity

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Preferred Scientific Name

  • Digenean infections of fish

Overview

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The Digenea (previously termed digenetic trematodes) are one of the three major taxa of parasitic Platyhelminthes, the other two being the Cestoda and the Monogenea. Digeneans are heteroxenous (i.e. they require more than one host to complete their life cycle), and their adult stage is parasitic in vertebrates. All major groups of vertebrates serve as hosts for adult digeneans. Apart from being hosts to adult digeneans, fish may also be infected by the metacercarial larval stage. With one exception, digeneans undergo part or all of their larval development in molluscs. Members of the genus Aporocotyle complete their larval development in polychaete annelids (Koie, 1982).

Very few adult-stage digeneans are known to cause significant harm to the fish host. Notable exceptions are the extraintestinal parasites, such as sanguinicoliid blood flukes, the cyst-forming didymozoids and the skin-inhabiting Transversotrema spp.

Metacercarial infection in fish is the main source of disease, with subsequent economic loss. Metacercariae may affect growth and survival, or disfigure fish so that they lose their market value as a food or ornamental product (Paperna, 1991, 1996). Some metacercariae in fisheries and aquaculture products (fish and shellfish) are a source of infections in humans and domestic animals (Ito, 1964; Deardorff and Overstreet, 1991). Metacercariae of Nanophyetus salmonis also transmit rickettsial infection (‘salmon poisoning’) to dogs (Philip, 1955).

The focus of this review is on parasites that cause disease in fish and the circumstances that promote the deterioration of the steady-state balance between the fish and its associated parasites. Recent practical aspects of the relationships between digeneans and their fish hosts are also included. The development in molecular methodologies did not bypass research on digeneans. A special section is included on molecular methodologies for the study and diagnosis of fish trematode infections. These methodologies have also been applied succesfully to explore digenean phylogenesis and, more importantly, to unveil life histories and consequently to provide a precise tool for specific identification (Cribb et al., 1998; Levy et al., 2002).

Digeneans that parasitize fish are numerous and diverse in their morphologies and life histories. Studies on adult digeneans and metacercariae require different approaches. The importance of digeneans to the fish culture has long been underestimated; also their risk to public health has not received adequate recognition. With the rapid development of warm-water aquaculture, as well as mariculture, and the spread of exotic culinary practices to Western societies risks to cultured fish and to consumers of infected fish are likely to become significant (Ko, 1995). Intra-intestinal adult digeneans are potentially pathogenic to maricultured fish: however, piscine digeneans are receiving less attention now than in the past. In the last decade, the burden due to massive infections by metacercariae has increasingly alarmed freshwater fish farmers. Some of the most troubling infections are of the gills by Centrocestus spp. and of the muscles and connective tissue by Bolbophorus spp. and by clinostomids (yellow grub), causing continuous losses or marketing problems in warm-water fish farms, notably of catfish in the USA and cichlids in Afro-Asia. To make matters worse, during the last decade, the snail vector, Melanoides tuberculata, of C. formosanus and subsequently the parasite have been introduced into Mexico and the southern USA.

Future Studies

In the last decade the transition in parasitology from an essentially zoological to a biochemical, immunological and molecular science has had only a marginal impact on fish digenean research. Future studies should include nutritional physiology, immunology and taxonomy that make use of DNA analysis. Temperature dependence of the immune response, combined with the peculiarities of the defence mechanisms against helminthic infections, offers an attractive and challenging research model. The emerging methodology of DNA taxonomy is potentially the best option for resolving taxonomic affinities and revealing life histories of digeneans by specific recognition of larval stages.

[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 and site specificity

According to Manter (1957), piscine digeneans are host specific, since the majority have been from a single host species or from hosts of the same or related genera. Adult-stage digeneans are primarily parasites of the digestive tract. However, several digenean families have become specialized to extra-intestinal habitats. These digeneans have special structural adaptations as well as functional specializations. Sanguinicolidae (e.g. Aporocotyle, Sanguinicola and Cardicola) are in the blood vessels (Smith, 1972). Transversotrematidae are ectoparasitic, attached under the scales of marine (Witenberg, 1944a), brackish-water (Velasquez, 1958; Angel, 1969) and freshwater fishes (Mills, 1979). Didymozoidae are in cavities and in tissues. Worms occur in the body cavity in M. cephalus (Skinner, 1975), in the orbits of Labeo spp. (Khalil, 1969) and encysted in the tissue near the body cavity, the mouth, the branchial cavity or the fins (see Yamaguti, 1958). Nemarobibothrioides histoldi lives in the skeletal muscles of the ocean sunfish (Mola mola) (Thulin, 1980), and Nemotobothrium texomensis in the ovaries of Ictiobus bubalus (Hoffman, 1967). Didymocystis palali and Didymocystis superpalati are in cysts in the palate of Neothunnus macropterus [Thunnus albacares]. Their late larval stages and adults are in the same fish host (Yamaguti, 1970). Species of Phyllodistomum (Phyllodistominae; Gorgoderiidae) are parasitic in the urinary bladder of fishes.

Other extra-intestinal digeneans belong to predominantly gut-inhabiting taxa. The bucephalid Paurorhynchus hiodontis occurs in the body cavity of Hiodon tergisus (Manter, 1957). The rhynchi, which are characteristic in other bucephalids, are degenerate in this species. One hemiurid, Gonocerca macroformis (Manter, 1957), infects cod (Gadus morhua) ovaries. Acetodextra ameiuri (Cryptogonimidae) infects the swim bladder, body cavity and ovaries of Ictalurus spp. (Hoffman, 1967).

Many digenean metacercariae in fishes have the ability to infect a wide range of hosts. D. spathaceum has been reported from more than 125 fish species (Hoglund, 1991). Only a few metacercariae have some degree of predilection for one or a few closely related piscine hosts. Metacercariae of C. tilapiae, Clinostomum cutaneum and B. levantinus occur only in cichlids, which mostly belong to Oreochromis and Sarotherodon (Paperna, 1964b; Ukoli, 1966; Britz et al., 1985; Yekutiel, 1985; Finkelman, 1988) Also, some metacercariae which have a wide range of hosts nevertheless have preferences for particular fishes. Heterophyes spp. in the eastern Mediterranean prefer grey mullet (Mugilidae) over other fishes and, among the grey mullets, Liza ramada (= Mugil capito) is the most preferred species (Paperna and Overstreet, 1981).

Distribution

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Host range and geographical distribution



The most important limiting factor for digenean dispersal is the molluscan hosts. Molluscs that serve as the first intermediate host belong to the Gastropoda and Pelecypoda (bivalves), but a few digeneans also develop in Scaphopoda (e.g. an elasmobranch parasite, Ptychogonimus megastoma, in Dentalium spp. (Palombi, 1941, Wright, 1971)). Marine and freshwater prosobranch snails are the most common intermediate hosts of fish digeneans. All known freshwater and marine Bucephalidae develop in bivalve molluscs. Bivalves are also the first intermediate hosts of Fellodistomatidae (Fellodistomum, Bacciger), Gorgoderidae (Phyllodistomum and Gorgodera) and Allocreadiidae (Allocreadium, Bunodera, Crepidostomum) (Yamaguti, 1958; Hoffman, 1967; Schell, 1970). Except Cyathocotylidae and Apatemon spp., which develop in freshwater prosobranch operculids, all other strigeids, as well as Clinostomidae and Plagiorchidae, are associated primarily with pulmonate snails (Yamaguti, 1958; Hoffman, 1960, 1967; Lo et al., 1982; Finkelman, 1988). However, Sanguinicolidae of marine and freshwater fishes diverge in their choice of intermediate hosts, and they include both pulmonates and operculids. The marine members can also be found in bivalves and in polychaetes (Smith, 1972). Specificity of individual digenean species to the molluscan host seems to be very restricted, usually to species, or even to subspecies. However, this has only been demonstrated in non-piscine digeneans, notably in human schistosomes (Paperna, 1968a;Wright, 1971). Some digeneans infecting widely distributed fish (such as the grey mullet, Mugil cephalus) seem to be less specific in their choice of molluscan host (Paperna and Overstreet, 1981).

Fig. 1. A. Pirenella conica (actual size 15 mm). B. Bulinus truncatus (actual size 6-11 mm).

Many digeneans of land vertebrates use fish for their metacercarial stages. Metacercariae are frequently the most common infections in fishes in the marine, estuarine and lacustrine littoral zones. These nutrient-rich waters are equally attractive to fish, molluscs and piscivorous birds; hence infections by skin, gill and visceral metacercariae are high. The high natural infections of young-of-the-year plaice with Cryptocotyle lingua and Stephanostomum baccatum in the north-east Atlantic littoral zones are well documented (MacKenzie, 1968; MacKenzie and Liversidge, 1975). There are also heavy heterophyid muscle infections in grey mullets and in juveniles of other fishes in lagoons and inshore marshes and estuaries in the eastern Mediterranean Sea. These areas are inhabited by high populations of the intermediate host Pirenella conica (Fig. 1A; Paperna, 1975; Paperna and Overstreet, 1981; Taraschewski and Paperna, 1981; Taraschewski, 1984). The Syrian-African rift is a major migratory route for birds between Europe and Africa. Water bodies from the Jordan to the East African Great Lakes have common fishes (cichlids, Clarias and Barbus), snails (Melanoides tuberculata (Fig. 2), Bulinus (Fig. 1B) and Lymnaea spp.) and metacercariae whose definitive hosts are herons (Ardeidae), cormorants (Phalacrocorax spp.) and pelicans (Pelecanus onochrotalus). Gill infection with the heterophyids Centrocestus spp. and Haplorchis spp., transmitted by M. tuberculata, occur in all young-of the-year cichlids that inhabit shallow waters (Farstey, 1986; Paperna, 1996). Bulinus and Lymnaea are more fastidious in their habitat demands, and demonstrate, particularly in the seasonal Mediterranean zone, greater seasonal and annual variations in abundance (Paperna, 1968b, c), and likewise the metacercariae they transmit (Neascus (black spot) and flesh and brain diplostomatids and clinostomids) (Khalil, 1963, 1969; Paperna, 1964a, 1996; Britz et al., 1985; Yekutiel, 1985; Finkelman, 1988; Dzikowski et al., 2003a). In the temperate inland waters of the northern hemisphere, infections by metacercariae of strigeoid digeneans are particularly common. The common infections in North America are 'white grub' (Posthodiplostomum minimum), encysting in the viscera (Hoffman, 1960; Grizzle and Goldsby, 1996), species of Bolbophorus, encysting in muscles (Overstreet et al., 2002), 'black grub' (pigmented skin metacercariae - Uvulifer ambloplitis and Crassiphiala bulboglossa), muscle metacercaria of Hysteromorpha triloba (Neascus musculicola (Hoffman, 1960; Huggins, 1972)) and 'yellow grub' (Clinostomum marginatum (Hunter and Hunter, 1934)). C. marginatum, together with 'diplostomulum' and 'neascus', has been reported to infect fish (Poecilia gillii) in dry-season pools fringing a stream in Costa Rica (Chandler et al., 1995). The diplostomatid eye infections are important and common in European and North American freshwater fishes. Dubois (1953) considered the American Diplostomum flexicaudum to be the European Diplostomum spathaceum. Both parasites develop to adult stage in gulls (Hoffman, 1960). D. spathaceum is in more than 125 fish species (Hoglund, 1991). Parasitic cataract, leading to blindness, has been reported in rainbow trout farmed in Patagonia, Argentina, where the local snail Chilina dombeiana is the molluscan host (Semenas, 1998). From five sites in the St Lawrence River, 12 fish species have been found infected with metacercariae of Diplostomum spp., with benthic fish (brown bullhead, Ictalurus nebulosus, and white sucker, Catastomus commersoni) having the heaviest infection. Diplostomum spp. metacercariae are in the vitreous humour of brown bullheads, and this is the same for the vast majority of diplostomatids in the other fishes (Marcogliese and Compagna, 1999). Wooten (1974) and Lester and Huizinga (1977) reported Diplostomum adamsi, Diplostomum gasterostei and Tylodelphis clavata from the humour or the retina of the eyes. Ocular diplostomiasis has been reported to occur in juvenile cichlids and Barbus spp. from East and South Africa (Thurston, 1965; Mashego, 1982; Paperna, 1996), while in Lake Kariba eye infection in Tilapia rendalli and Oreochromis mortimeri has been about 85% (Douellou, 1992).

Fig. 2. Melanoides tuberculata (actual size 20 mm).

The host range and geographical distribution of trematodes have also been altered through human intervention. Snails (carrying larval trematodes) have been translocated with fish culture stock or the ornamental fish trade from one geographical region to another (Scholtz and Salgado, 2000; Scholtz et al., 2001). This allowed the expansion of piscivorous bird trematodes, with metacercariae that are not piscine host specific. Some of these resulting infections, most notoriously Centrocestus formosanus and Haplorchis pumilio, dispersed through the introduction of the Afro-Asian snail Melanoides tuberculata, became health risks for both wild and farmed fish in many water bodies in the USA and Mexico (Mitchell et al., 2000; Brandt southwest.fws.gov/fishery/ trematode.pdf; Scholtz et al., 2001). Four aquacultured fish species (Ictalurus punctatus, hybrid Morone, Notemigonus crysoleucas and Pimephales promelas) became readily infected when exposed to C. formosanus cercariae (Mitchell et al., 2002); in Mexico, farmed carp were also infected. The only definitive host identified was the heron Butorides striatus (Scholtz et al., 2001).

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

Sea Areas

Atlantic, NortheastPresentSmith and Williams, 1967; MacKenzie, 1968; Shotter, 1973; MacKenzie and Liversidge, 1975
Atlantic, NorthwestPresentSindermann and Resenfield, 1954; Scott, 1969
Mediterranean and Black SeaPresentFares and Maillard, 1974; Paperna, 1975; Maillard et al., 1980; Paperna and Overstreet, 1981; Taraschewski and Paperna, 1981; Taraschewski, 1984

Asia

ChinaPresentZeng and Liao, 2000; Wang et al., 2001
IndiaPresentMadhavi, 1979; Madhavi and Rukmini, 1991; Madhavi and Rukmini, 1992; Mohan et al., 1999
IsraelPresentPaperna, 1964a; Paperna, 1964b; Taraschewski and Paperna, 1981; Finkelman, 1988; Paperna, 1996
JapanPresentYanohara and Kagei, 1983; Ogawa et al., 1989
MalaysiaPresentAnderson and Shaharom-Harrison, 1986
TaiwanPresentLiu, 1979; Lo et al., 1981; Ooi et al., 1999
ThailandPresentSithithaworn et al., 1997; Sukontason et al., 1999

Africa

Central AfricaPresentDe and De, 1952
Congo Democratic RepublicPresentKabunda and Sommerville, 1984
East AfricaPresentPaperna, 1996
GhanaPresentUkoli, 1966
MoroccoPresentDollfus, 1954
NigeriaPresentEzenwaji and Llozumba, 1992; Ezenwaji and Inyang, 1998
South AfricaPresentLombard, 1968; Mashego, 1982
SudanPresentKhalil, 1969
UgandaPresentThurston, 1965
ZimbabwePresentBeverly-Burton, 1963; Douellou, 1992

North America

CanadaPresentMarcogliese and Compagna, 1999
MexicoPresentGarcía et al., 1993; Velez-Hernandez et al., 1998; Scholz and Salgado-Maldonado, 2000; Scholtz et al., 2001
USAPresentWales, 1958; Rogers, 1972; Mitchell et al., 1982; Hoffman et al., 1985; Stables and Chappell, 1986; Lorio, 1989; Deardorff and Overstreet, 1991; Mitchell, 1995; Mitchell et al., 2000; Terhune et al., 2002; Dzikowski et al., 2004
-CaliforniaPresentMartin, 1955
-FloridaPresentSkinner, 1975
-HawaiiPresentMartin, 1969; Yamaguti, 1970
-MississippiPresentLevy et al., 2002; Terhune et al., 2002
-OregonPresentMillemann and Knapp, 1970
-South DakotaPresentHuggins, 1972

Central America and Caribbean

Costa RicaPresentChandler et al., 1995
Puerto RicoPresentCable, 1954

South America

ArgentinaPresentSemenas, 1998

Europe

FinlandPresentValtonen and Gibson, 1997
FrancePresentMaillard et al., 1980; Euzet and Raibaut, 1985
GermanyPresentHoffmann et al., 1990
Isle of Man (UK)PresentShotter, 1973
NetherlandsPresentBroek and Jong, 1979
SwitzerlandPresentMuller, 1995
UKPresentWootten, 1974; Evans, 1978; McKeown and Irwin, 1997
-ScotlandPresentStables and Chappell, 1986; Barber and Crompton, 1997

Oceania

AustraliaPresentAngel, 1969; Koie and Lester, 1985
New ZealandPresentMcArthur, 1978

Pathology

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Adult trematodes

Adult intestinal trematodes are normally considered not to cause disease even when their numbers are high. Extra-intestinal parasites are, however, potentially pathogenic. Blood flukes (sanguinicolids and aporocotyles) cause considerable damage to the gills and impair respiration. Adult worms and eggs can physically obstruct the passage of blood and cause thrombosis and subsequent tissue necrosis (Hoffman et al., 1985; Ogawa et al., 1989). Extensive rupture of the gill lining by emerging miracidia and tissue response around trapped eggs in capillaries and internal organs appeared to be the direct cause of mortality of bloodfluke-infected fish (Figs 7B-D, 14 and 15). This is the case in Sanguinicola fontinalis-infected brook trout (S. fontinalis) in Pennsylvania, USA (Hoffman et al., 1985). In flatfishes (Hippoglossoides platessoides, L. limanda and Pleuronectes platessa) infected with A. simplex, fragments of disintegrated worms in the efferent gill filament arteries cause the gill filament to become stunted and necrotic (Koie, 1982). The blood fluke Paradeontacylix sp. causes mass mortalities of cage-cultured Seriola purpurascens. Infections with up to 1000 worms per single gill filament have been found and they invoke gill hyperplasia. Eggs in the gills and the ventriculum are encapsulated and they induce papillate proliferation of the endothelium of afferent branchial arteries. However, there is necrosis in the gills or elsewhere (Ogawa et al., 1989). Proliferation of the arterial endothelium has also been reported in common carp infected with S. inermis (Prost, M., quoted in Lucky, 1964). Heavy blood loss occurs when miracidia escape from the gills and apparently this is the main cause of mortality in trout fingerlings infected with Sanguinicola davisi and Sanguinicola klamathensis (Wales, 1958; Davis et al., 1961). Loss of blood is evidenced by the pale colour of the gills and the decline in packed cell volumes and oxyhaemoglobin levels (Evans, 1974b). Heavily infected cultured carp apparently die of suffocation during transportation (Lucky, 1964; Smith, 1972). In chronic infections, adult worms disperse and become stranded in the heart, kidneys and caudal vessels. Some eggs become encapsulated, and may also become surrounded with focal granuloma. Nodular foci occur in the heart, head kidney and spleen of Oreochromis spp. (Fig. 7D), in the gills, heart and kidneys of carp (Scheuring, 1922) and in the liver of Paracardicola hawaiensis-infected Tetraodon hispidus (Martin, 1969). Eggs of S. armata are widespread in the viscera of grass carp (C. idella) and bighead (A. nobilis), but the tissue response is negligible (Anderson and Shaharom-Harrison, 1986). Eggs in the kidneys of S. klamathensis-infected cutthroat trout (S. clarki) cause hypertrophy or necrosis of the renal epithelium and renal calcifications (Evans, 1974a). In sanguinicolosis of cichlid fishes and in Pearsonellum corventum-infected serranids (Plectropomus leopardus), granulomas form around eggs in the viscera and these are accompanied by melanomacrophage aggregates (Fig. 15; Overstreet and Thulin, 1989). Dead A. simplex are encapsulated in the superficial parts of the liver (Koie, 1982). General effects on growth and condition are not always evident however, and they are correlated with the duration and severity of the infection. Encapsulated lesions around residues of Sanguinicola eggs in the heart have been found in a dying native population of Sarotherodon galilaeus in Lake Kinnereth, Israel (Paperna, 1996). Smith and Willams (1967) did not find any apparent effect on weight in hake (Merluccius merluccius) infected with Aporocotyle spinosicanalis.

Fig. 15. Degenerate eggs of the sanguinicolis Pearsonellum corventum in the heart of the fish Plectropomus leopardus. A. Encapsulated egg mass enclosed by infiltrated macrophages. B. Granuloma encased egg residue within a melanomacrophage. (From Overstreet and Thulin, 1989, courtesy of R. Overstreet.)

Encapsulation of didymozoid worms induces no more than limited local reactions (Fig. 8; Lester, 1980; Perera, 1992a, b), although at times there is also haemorrhaging of peripheral capillaries in the buccal dermis of Platycephalus fuscus (Lester, 1980). Nematobothrium spinneri, in the muscles of Acanthocybium solandri, is enclosed in a thick-walled capsule. Connective tissue and blood capillaries are interdigitated with worm coils, which facilitates worm feeding on blood (Lester, 1980). Only natural infections have been recorded, and these are likely to be moderate. It is not known whether these infections are harmful to maricultured fish. T. patialense, an ectoparasite on the Brachydanio rerio integument, leaves pressure and feeding indentations on the body surface. Tissue regeneration occurs soon after the worm changes position (Mills, 1979). Kidney damage induced by heavy infection of the urethra with Phyllodistomum umblae (=conostomum) adversely affects survival of charr migrating from fresh water to sea water (Berland, 1987). A. kubanica are pathogenic to R. rutilus heckeli in the Sea of Azov when they infect the kidneys rather than the intestine (Bauer, 1958).

Metacercariae

Clinical effects of infection are often not obvious. Metacercariae in supposedly sensitive organs, such as the brain, cranial nerves or spinal cord, e.g. B. gracilescens in cod, G. morhua (Matthews, 1974), Diplostomum mashonense and Diplostomum tregenna in Clarias spp. (Beverly-Burton, 1963; Khalil, 1963) and Ornithodiplostomum ptychocheilus in P. promelas (Hoffman, 1958; So and Wittrock, 1982), do not necessarily have obvious debilitating effects on fish even when the number of parasites is relatively high and despite visible structural damage to the organs.

A sudden massive outbreak of infection is often fatal. Exposure to massive numbers of cercariae will kill fry within a few hours (Sommerville, 1982), but such exposures do not normally occur in nature. Cercariae penetrate and encyst deeper in the tissues of small fishes and the relatively larger cysts may interfere with organ function (Fig. 16). The effects of cercarial infestation are most severe in 0-year-class plaice and minimal in the 1+-year class (Sommerville, 1981). Hoffmann et al. (1990) reported a massive Bucephalus polymorphus infestation and mortality of cyprinid fishes after the water temperature was suddenly increased from 12-14 to 20°C. Massive metacercariae infections in juvenile (0-class) fish have been incriminated as an important cause of natural mortalities, e.g. C. lingua infections of plaice, P. platessa (MacKenzie, 1968), Bolbophorus levantinus in cichlid fishes (Fig. 17B; Yekutiel, 1985; Paperna, 1991). Population studies and field observations suggest that fish heavily infected with metacercariae are lost from the host population (Chubb, 1979; Lemly and Esch 1984a, b). Lemly and Esch (1984b) confirmed that heavily U. ambloplitisinfected young-of-the-year bluegill sunfish (50 per fish) did not survive the winter because of depleted reserves.

Fig. 16. Metacercariae in fish tissues. A. Cartilaginous cyst around Centrocestus sp. metacercariae in gills of Oreochromis aurea, Lake Kinnereth. B. Ascocotyle coelostoma (live) in the truncus arteriosus of Liza ramada. C. Encysted Phagicola nana in the gut wall of Micropterus salmoides. D. Pigmented Neascus ('black spot') encysted on fins of young Oreochromis hybrids.

Entry into tissue and encystment

A pronounced inflammatory response often accompanies penetration and early migration. This is particularly obvious following heavy exposure. Fish may even die from the penetration wounds caused by D. spathaceum (Ferguson and Hayford, 1941) and from tissue damage (focal haemorrhages) caused during H. pumilio migration in muscles (Sommerville, 1982) and in the brain caused by D. adamsi (Lester and Huizinga, 1977). Cercariae of C. lingua and S. baccatus induce temporary epidermal lesions. They may also cause some disruption of connective tissues, inflammatory cell proliferation, sometimes myofibrillar necrosis (associated with bacteria) and reactive swelling of the intermuscular septa (McQueen et al., 1973; Sommerville, 1981). The inflammatory reaction is particularly intense around non-encysting metacercariae (e.g. Rhipidocotyle johnstonei and Prosorhynchus crucibulum in plaice and turbot) (Matthews, 1973a, 1974) and precedes the eventual enclosure of the parasite in a fibrous capsule (Yekutiel, 1985; Fig. 18A). The fibrous capsule produced by the host is superimposed on the acellular wall secreted by the encysting cercaria (Fig. 18B). The peripheral inflammatory cellular infiltrate that precedes fibrous capsule formation consists mainly of macrophages (Sommerville, 1981). Inflammatory cells become stratified into an epitheloid (Fig. 18B), while the innermost layer gradually degenerates. This early capsule, which has the form of a chronic granuloma (and also contains giant cells) is gradually replaced by the fibrous capsule (Fig. 16C; Sommerville, 1981; Yekutiel, 195). At 5°C, cyst formation is much slower than at 15°C (McQueen et al., 1973). Centrocestus metacercariae on the gills become encysted in cartilaginous capsules (Fig. 16A). The 'sand-grain' grub of yellow perch is caused by metacercarial cysts of Apophallus brevis. It is composed of bone, which becomes clustered around the parasite within the peripheral blood vessels. Such cysts have also been identified in ancient fossil perch. The bony structure has two opposite escape canals for the parasites and has lines interpreted as the growth rings of the bone (Sinclair, 1972). Metacercariae of B. levantinus migrate in tissues before encysting in the muscles (Paperna and Lengy, 1963). The inflammatory response and macrophage proliferation are limited to host tissues damaged in the earlier stages of the encystment (Yekutiel, 1985). Cellular damage around Heterophyes spp. infecting grey mullet is minimal even in heavy infections (6000 metacercariae per gram muscle) (Paperna, 1975). Under certain circumstances, however, the same metacercariae can induce more extensive cellular changes, particularly during the late stages of infection when damaged tissues and dead parasites have accumulated (Font et al., 1984). In muscles of grey mullet infected with Stellantchasmus falcatus, myofibrils degenerating near the parasitic cysts are replaced by large fat cells (Lee and Cheng, 1970). Cellular and deposited ground substances may vary with species, but may also be affected by the host tissues. Collagen may be deposited in the liver (Mitchell, 1974) and in the spleen (Font et al., 1984), but it is usually absent. When crowded on the serosal surface of the pyloric caeca (Fig. 16C), metacercariae of Phagicola nana evoke intense proliferation of the connective tissues, with a considerable amount of collagen. When the parasite is in the submocosa, the proliferation includes smooth muscle (Font et al., 1984).

Fig. 17. Diplostomatid metacercariae: A. Diplostomum sp. removed from a visceral cyst in Clarias gariepinus, Uganda. B. Heavy Bolbophorus levantinum metacercaria in muscles of young Oreochromis aurea from Lake Kinnereth. C. Early-stage B. levantinum metacercaria with expanded posterior half. D. Later-stage B. levantinum metacercaria with emptied posterior end (actual size 0.6-0.8 mm).

Gill infection due to Centrocestus spp.

Natural infections by Centrocestus sp. are common in cichlids (as well as in introduced Gambussia affinis) throughout the distribution range of its snail host M. tuberculata along the Syrio-African rift valley and nearby fish farms (Farstey, 1986; Paperna, 1996). Centrocestus is also in carp fry in India (Mohan et al., 1999). In the Far East, a different species, C. formosanus, infects farmed eel (Yanohara and Kagei, 1983) and grass carp (C. idella) (Zeng and Liao, 2000). Introduction of M. tuberculata to Mexico and to the USA extended C. formosanus infection to farmed (carp) and native fishes (Mitchell et al., 2000; Scholtz et al., 2001). Metacercariae encysted on the gill filament cartilage are enclosed by a cartilagenous cell capsule (Fig. 12A). Cercariae penetrate the gill filaments and encyst in the connective tissue adjacent to the cartilage layer. Chondroblasts proliferate from the ray perichondrium and, following deposition of ground substance, themetacercaria becomes encapsulated (by day 7 postinfection) within a cartilaginous extension of the filament ray (Fig. 16A; Farstey, 1986; Paperna, 1996). Infection induces epithelial hyperplasia; infected gill filaments are thickened and often shortened and distorted when infection is heavy, and sometimes haemorrhagic. The proliferated cartilage causes filaments to expand several times the normal diameter, and this leads to fusion of filaments and disruption of the normal gill morphology. Heavy gill infection appears to lower respiratory efficiency. Heavily infected fish seem to be selectively dying out from the natural and farmed populations, particularly under stressful circumstances (e.g. overwintering and extreme oxygen conditions). Heavily infected fish do not survive well during transportation. This has been demonstrated under simulated experimental conditions: all young cichlids (S. galilaeus) heavily infected with Centrocestus (116 ± 48 per fish) succumbed, while all same-size lightly infected fish (with 15 ± 15 per fish) survived during a 3 h journey at ambient temperature of 24°C (Farstey, 1986). Infections in common carp fry in India similarly cause both acute and chronic mortalities (Mohan et al., 1999).

Fig. 18. Histopathology of Bolbophorus levantinum infection in young Oreochromis aurea. A. Macrophage infiltration around young unencysted metacercaria. B. Maturing metacercaria with 'reserve cells' (r) enclosed in a parasite-derived wall (w) and foci of macrophage proliferations in the surrounding muscles (arrows).

Dermal encystment

Cysts formed around certain dermal metacercariae may contain cells heavily loaded with melanin (and, exceptionally, other pigments). These are commonly called 'black spot', and are formed by Neascus metacercariae from the genera Crassophiala, Ornithodiplostomum and Uvulifer (Hoffman, 1960; Wittrock et al., 1991; (Figs 16D and 19A, B) and heterophyids of the genera Cryptocotyle and Haplorchis. Cutaneous melanosis and cloudy eye have been reported in flounders heavily infected with pigmented C. lingua (Mawdesley-Thomas and Young, 1967).Metacercariae of a species of Clinostomum ('C. cutaneum'; Paperna, 1964a, b) encyst, sometimes in large numbers, beneath the scales of cichlid fish in Lake Kinnereth, Israel (Fig. 19C).

Heart infections

Metacercariae of some Ascocotyle and Phagicola have a predilection for the heart or truncus arteriosus (Sogandares-Bernal and Lumsden, 1964), but in heavy infections they may spread to other tissues. Cardiac pathology, resulting from thickening due to fibrogranulomatosis of the epicardium, is associated with infection of the pericardium with the strigeid A. gracilis (Watson et al., 1992). Tort et al. (1987) demonstrated that the in vitro pumping performance of hearts in infected fish was reduced by as much as 50%. Cardiac infections are also commonly caused by heterophyids of the genera Phagicola and Ascocotyle (Fig. 16B; Sogandares-Bernal and Lumsden, 1964; Stein and Lumsden, 1971). Ascocotyle pachycystis encysts in the lumen of the bulbus arteriosus, infection accumulates with age (up to 6800 per fish) and heavy infections reduce growth and overwintering survival (Coleman and Travis, 1998).

Fig. 19. Metacercarial skin infection in Lake Kinnereth cichlids. A, B. Low and heavy 'black spot' (Neascus) infection in young pond-reared Oreochromis aurea × nilotica. C. Clinostomum 'cutaneus'-infected Tristramella simonis (top) and Tilapia zilli (bottom).

Visceral infections

Metacercariae of Haplorchis are found throughout the integumental connective tissues (skin, fins and gills) (Paperna, 1964a; Sommerville, 1982; Yekutiel, 1985). Heavy visceral infection of P. minimum ('white grub'; 500-2300 metacercariae in 50-65 mm long fish) in fathead minnow causes abdominal dropsy (Fig. 20) with milky ascitic fluid that contains up to 92% leucocytes and 8% erythrocytes. Granulomas develop around degenerating parasites and parasitic debris (Mitchell et al., 1982). Milder or no clinical impacts have been reported in P. minimum centrarchi (white grub) infections of largemouth bass (M. salmoides) (Grizzle and Goldsby, 1996). Massive invasion by the heterophyid P. cheni in Taiwan of Japanese eels caused severe morbidity. Their eyes were cloudy white and swollen with oedema and hemorrhages. Metacercariae, encysted in the muscle tissue, were numerous around the eyeball. Once transmission was interrupted, recovery occurred within 2 weeks (Ooi et al., 1999). Stephanostomum tenue in the pericardial cavity of elvers of Anguilla rostrata impedes swimming (Oliveira and Campbell, 1998). Bolbophorus sp. invades as a protodiplostomum and then encapsulates in the muscles (Figs 17C, D and 18). Heavy infections of B. levantinus in Israel caused mortalities among young-of-the-year cichlids in Lake Kinneret and in farmed stock. Parasites invaded the major part of the somatic muscles and disfigured the fish (Fig. 17B); Yekutiel, 1985; Paperna, 1996). In the USA, B. damnificus and another, yet to be identified, Bolbophorus similarly invade and cause muscle damage to channel catfish farmed in southern states (Overstreet et al., 2002; Terhune et al., 2002).

Several species of clinostomiids occur in fish. C. tilapiae encysts in the connective tissues of the ventral articulation of gill arches, and sometimes also on the surface of the kidneys. C. complanatum becomes established in the muscles, with a preference in Tor (Barbus) canis for red muscles (92%) and a predilection for the branchial and mandibular muscles (62%) (Finkelman, 1988). C. cutaneum settles beneath the scales (Paperna, 1964a, b; Finkelman, 1988). Natural infections are sometimes exceptionally heavy; none the less, a full-sized fish can tolerate a remarkably high number of worms. A Tristramella sacra in Lake Kinneret, Israel, was found virtually covered with C. cutaneum; 130 Nephrocephalus sp. were recovered from a Heterotis niloticus, 70 Clinostomum sp. from Synodontis membranaceus (Ukoli, 1966) and the same number of C. complanatum from T. (B.) canis (Finkelman, 1988). Farmed juvenile cichlids, Oreochromis crossbreeds in Israel and Oreochromis mossambicus in Africa succumbed to infections as low as fewer than ten, or even three to five, of either C. tilapiae or Euclinostomum heterostomum; worms invaded the kidneys and the branchial cavity. Mortalities have also been reported among Clarias spp. due to E. clarias (Ezenwaji and Llozumba, 1992). Experimentally infected O. mossambicus fingerlings (30-35 mm long) died 62 days after being experimentally infected by 75-81 E. heterostomus (Donges, 1974). C. complanatum infections are regarded as a serious problem in Taiwan, causing death among farmed P. altivelis (ayu). The body wall of infected fish is profusely encysted with metacercariae, leading to its perforation (Lo et al., 1981). Heavy loads of Clinostomum van der horsti metacercariae often congest the body cavity of their fish host, the mormyrid Gnathonemus macrolepidotus in southern Africa (Ortlepp, 1935). In the USA, metacercariae of C. marginatum (yellow grub) encyst in the flesh of farmraised channel catfish. Apart from direct pathological damage, infected fish may not be marketable (Lorio, 1989). There are two reports of Clinostomum infections in humans, causing acute laryngitis, one in Israel and the other in Japan (Witenberg, 1944b; Kamo et al., 1962). The worms were adults and the infections were probably due to consumption of metacercaria-infected fish.

Fig. 20. Abdominal dropsy in Postdiplostomum minimum-infected Pimephales promelas (43 mm long) (after Mitchell et al., 1982, courtesy of G.L. Hoffman).

Eye damage in metacercarial infections

Damage to eyes may either be caused by metacercariae with a predilection for the organ (e.g. D. spathaceum) or be a nonspecific side effect (e.g. corneal infection by integument-encysting metacercariae). Impairment of vision is aggravated when metacercarial cysts are accompanied by melanin pigment (black spot). Severe infection leads to exophthalmos, cataracts and even complete collapse of the eye. Blindness can be uni- or bilateral (Sindermann and Resenfield, 1954; Mawdesley-Thomas and Young, 1967). D. spathaceum is a specific pathogen of the eye. The non-encysted metacercariae invade the lens, the vitreous humour and the retina. Detailed pathological and histopathological descriptions of diplostomiasis in catfish (Ictalurus melas) are in Larson (1965) (Fig. 21) and in rainbow trout in Shariff et al. (1980). Farmed rainbow trout yearlings with infection limited to a cataract (small white opacity in the lens with parasites visible beneath the lens) feed normally and remain in good condition. However, in 2-year-old trout, the cataract is more prominent, with the lens dislocated into the anterior chamber or absent. Fish at this stage are dark, apparently blind and emaciated and feed only from the bottom of the tank. Parasites are most frequently in the cortical region of the lens, within a cavity with fine debris, which results from liquefaction of the cortical fibres. Degeneration of the lens is evident. In the chronic condition, the lens becomes irregular and wrinkled from adherences with the inner components of the eye and is often ruptured. Thickening of the lens capsule leads to exfoliation, with proliferation of the epithelial layer. The iris is greatly distorted and is in contact with the lens. In catfishes (I. melas and I. nebulosus), the lens herniates near the point of attachment to the dorsal ligament, with metacercariae spilling into the protrusion and leaving the remainder of the lens clear for visual function (Fig. 21; Larson, 1965). Entry of parasites to the subretinal level results in retinal detachment. A cellular response usually becomes evident only in later-stage infections. Macrophages aggregate around the debris from ruptured lenses, in the damaged periphery of the iris, in the choroid and around necrotic foci in the optic nerve. The combined effects of cataracts and retinal detachment cause blindness (Shariff et al., 1980).

Fig. 21. Eye diplostomiasis in Ictalurus melas. A. Cross-section of the eye showing herniation (h) of the lens (l) and the metacercariae inside (arrows). B. Damaged lens (l) with proliferated epithelium around a necrotic core (i); note vascularization of the vitreous humour (arrow). C, D. Cross-section of herniated lenses; empty spaces contained worms before processing. (After Larson, 1965, courtesy of the author.)

Crowden and Broom (1980) described loss of efficiency and increase in time devoted to feeding. As the infection progresses, fish spend more time in the surface layer of the water. This would increase their risk of predation by gulls. When D. adamsi cercariae enter (in groups of 20 or more) the retina of Perca flavescens and remain unencysted, they form a cavity between the photoreceptor layer and the pigment epithelium. Diplostomum schudderi becomes established in a similar manner in the retina of Gasterosteus aculeatus. In both fish hosts, cytological damage and alterations occur in both layers of the retina. This suggests loss of vision in the foci of infection in the retina, but an overall minimal visual loss due to the peripheral location of the parasites (Lester and Huizinga, 1977). Metacercariae of Diplostomum compactum infected Oreochromis aureus and O. mossambicus introduced into Lake Amela, Mexico. The metacercariae enter the aqueous and vitreous humours of the eye, causing corneal oedema, conjunctivitis, neuritis of the optical nerve, eosinophilic iridoclitis and uveitis. They also invaded the brain, causing multifocal gliosis, eosinophilic meningitis, brain oedema and spongiosis (Garcia-Luis et al., 1993). Metacercariae of Posthodiplostomum brevicaudatum encysted only in the vitreous humour and in the retina and caused less obvious eye damage (Donges, 1969).

Estimating the extent of mortality in affected fish populations

Estimation of metacercaria-induced host mortality in natural fish populations has been attempted by extrapolating quantitative data on frequency distribution of infection. The decline in variance was considered to result from the loss of heavily parasitized fish. Hence a measure of overdispersion, in comparison with negative binomial distribution or by calculating the ratio of variance to mean, was advocated as an indirect method of estimating fish mortality (Lester, 1977, 1984; Anderson and Gordon, 1982; Gordon and Rau, 1982; Kennedy, 1984). Seasonal changes in the degree of overdispersion were related to suggested parasite-caused mortality in bluegill during the winter (Lemly and Esch, 1984b). However, extrapolations from field data have met with a variable degree of success and at times have produced ambiguous results (Kennedy, 1984; Lester, 1984).

Immune responses

In contrast to the wealth of information that has accumulated on the immunology of digeneans in higher vertebrates (Butterworth, 1984), data on fish-infecting digeneans are more preliminary in nature. The main difference between immune responses of fish and those of mammals and birds is that the ambient temperature affects antibody production in fish (Avtalion et al., 1973; van Muiswinkel, 1995). Eosinophilic granulocytes, which play an important role in protective immunity in mammalian digenean (schistosome) infections (Butterworth, 1984), do not seem to be as important in piscine infections (Hoglund and Thuvander, 1990). However, it is not certain whether piscine eosinophils are homologous with those of in mammals (Ellis, 1977). Aaltonen et al. (1997) demonstrated humoral immune response of Rhipidocotyle fennica, it was detected in wild R. rutilus in lakes where fish are infected by the parasite. Infection by R. fennica increases towards late summer: antibody levels are higher in sera collected in September than in those collected in June. Antibodies were produced against homogenized cercariae in experimentally immunized R. rutilus; however, the response was more pronounced after exposure to live cercariae. Western blots showed that responses in fish injected with homogenized cercariae were different from those exposed to live cercariae. Fish injected with the homogenate were protected on challenge.

Trout repeatedly exposed to cercariae of Diplostomum over a 12-week period developed protective immunity, but specific antibodies could not be detected in the blood (Hoglund and Thuvander, 1990). The protection seemed to be due to cell-mediated immunity or to a non-specific mechanism of protection against the migrating diplostomulae. In mammals, induction of cellular immunity is required for protection against helminth infections (Butterworth, 1984). Migrating diplostomulae induce non-specific cellular responses, which involve infiltration of neutrophils and monocytes (Ratanart- Brockelmann, 1974). Speed and Pauly (1985) showed that the survival time of immunized, challenged fish increased by 4 months over that of non-immunized controls. An immunocytochemical study demonstrated putative neurotransmitters in metacercariae of Diplostomum sp. and Cotylurus erraticus from rainbow trout. Cholinergic and serotoninergic staining was found primarily in the central nervous system (CNS) and in cell bodies associated with the ventral and dorsal nerve cords. Peptidergic immunoreactivity was localized in the CNS and the peripheral nervous system, revealing an extensive innervation within the holdfast organ and around the oral and ventral suckers (Barton et al., 1993).

Precipitating antibodies against antigen from the intestinal digenean Telogaster opisthorchis were demonstrated in the sera and gut mucus of eels, Anguilla australis schmidtii and Anguilla dieffenbachii (McArthur, 1978). Wood and Matthews (1987) reported induction of humoral antibodies, sensitized pronephric leucocytes and cytotoxic serum factors in Chelon labrosus exposed to C. lingua cercariae. Precipitating antibodies were detected, using the Ouchterlony technique, in plaice (P. platessa) infected with C. lingua and R. johnstonei. The immune sera had an immobilizing effect on the cercariae. Antibodies reacted with the tegument and with secretory glands in cercariae of both parasite species. Antibody production was temperature-dependent and did not occur at 5°C (Cottrell, 1977). The antibodies were macroglobulins, and they resemble mammalian immunoglobulin M (IgM). Bortz et al. (1984) detected humoral antibodies in trout with natural infections of D. spathaceum and trout injected with sonicated metacercariae using an enzyme-linked immunosorbent assay (ELISA). In trout immunized with antigens prepared from cercariae and diplostomulae, anti-cercarial and anti-diplostomulae circulating antibodies were demonstrated using ELISA and immunofluorescence. The two cross-reacted, although the reaction between anticercarial sera and diplostomatid antigen was weaker. Fluorescence studies indicated the tail region of the cercariae to be strongly antigenic (Whyte et al., 1987).

List of Symptoms/Signs

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SignLife StagesType
Finfish / 'Dropsy' - distended abdomen, 'pot belly' appearance - Body Aquatic:Adult Sign
Finfish / Hyperplasia - Gills Aquatic:Adult Sign
Finfish / Mortalities -Miscellaneous Aquatic:Adult Sign
Finfish / Nodules - Gills Aquatic:Adult Sign
Finfish / Paleness - Gills Aquatic:Adult Sign
Finfish / Parasites - Gills Aquatic:Adult Sign
Finfish / Pop-eye - Eyes Aquatic:Adult Sign

Epidemiology

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Seasonal distribution of adult stages

The seasonal occurrences of digeneans in freshwater fishes have been extensively reviewed (Chubb, 1979). However, a similar review on marine fish is not available. In some digenean infections, fish acquire their parasites by direct cercariae penetration (Sanguinicolidae and Transversotrematidae). Sanguinicola inermis, which infects the common carp (Cyprinus carpio) in Europe, is transmitted during the summer (Lucky, 1964). Sanguinicola sp. infecting cichlid fishes in Lake Kinnereth and the nearby fish ponds in Israel appears by spring and later by autumn (Paperna, 1996). Cercariae of Aporocotyle simplex have been found in December, and flatfishes are infected in 6ºC water (Koie, 1982). The tube-dwelling polychaete Lancides vayssieri, host of larval stage aporocotyles, occurs in Antarctic waters with temperatures not exceeding 1.6ºC (Martin, 1952). The ratio of juvenile to adult digeneans in a fish enables a determination of time of infection. In general, transmission usually takes place during the summer or early autumn (Chubb, 1979).

Infection transmitted via the predation of metacercariae-infected intermediate hosts can be only partially correlated with climatic fluctuations (Scott, 1969; Shotter, 1973). In the North Atlantic, Lecithobothrys bothryophorum infections in juvenile Argentina silus peak in May (Scott, 1969). The same has been reported for whiting (Odontagatus merlangus) infected with Derogenes varicus, Hemiurus communis, Stephanostomum pristis and Lecithaster gibbosus (Shotter, 1973). In the North Atlantic, infection by Derogenes and Hemiurus may be linked to peak abundance of zooplankton in the summer (Shotter, 1973). It becomes more complicated when there are several optional or consecutive intermediate hosts (e.g. D. varicus; see Koie, 1979a, 1985). European sea bass (Dicentrarchus labrax) become infected with Bucephalus haimeanus upon entry into estuarine habitats, where both molluscan hosts (Cardium edule) and metacercariae-infected gobies occur (Matthews, 1973b). Feeding on plankton has been correlated to Bunodera luciopercae infection of Perca fluviatilis in fresh water (Chubb, 1979). There is seasonal transmission of Allocreadium fasciatus in tropical waters in India and this has been assumed to be related to blooms of copepod populations following the monsoon season (Madhavi, 1979).

Seasonal distribution of metacercariae infections



It is difficult to extrapolate seasonal variations from direct counts of metacercariae on fish, since such infections normally persist for over 12 months (Donges, 1969). Under these circumstances, infection accumulates with fish age (presumably with size) (Karlsbakk, 2001). Marcogliese and Compagna (1999) suggested that, in the St Lawrence River, fish become infected with metacercariae in their first year of life, and that benthic fish are more prone to infections. In northern Finland, where eye diplostomiasis is widespread in numerous fish species (habitats ranging from freshwater lakes to brackish marine), infestation is stable despite the extremely narrow transmission window between the snail and the fish. Extended longevity and continuous accumulation precluded detection of any seasonal pattern (Valtonen and Gibson, 1997). This is similar in brain infections of Phoxinus phoxinus by Diplostomum phoxini in two habitats in Scotland (Barber and Crompton, 1997). In wild Rutilus rutilus, in the UK, the abundance of eye diplostomiasis peaks in late June and mid-September each year, while in farmed fish abundance is continuous. This has been explained by mortality among infected wild fish (McKeown and Irwin, 1997). In the Jiangkou reservoir, China, metacercariae of the bucephalid Dollfustrema vaneyi infect Pseudobagrus fulvidraco in late spring and summer, and the metacercaria has a predilection for the eyes (Wang et al., 2001). Presence of young metacercariae may indicate active transmission; however, differentiation of metacercariae by age is feasible only when they transform with time (e.g. Bolbophorus levantinus: Paperna and Lengy, 1963; Centrocestus sp.: Farstey, 1986). Seasonal fluctuations in infection can be detected among the young-of-the-year fish or in transitory fish after they enter a new habitat (Lemly and Esch, 1984a). Active shedding of cercariae and infection of fish commonly take place during the warmer times of the year in fresh water (Chubb, 1979; Lemly and Esch, 1984a; Stables and Chappell, 1986) and in marine and estuarine habitats (Matthews, 1973b; Koie, 1975; Cottrell, 1977). In some habitats, infection in snails only peaks in the autumn (McDaniel and Coggins, 1972). Transmission during the winter is rare since snails do not shed their cercariae below 10ºC (Stables and Chappell, 1986). There is, however, a report on invasion of Oncorhynchus mykiss by Diplostomum opataceum at low winter temperatures. This infection had apparently been acquired by predation on Lymnaea containing precocious (progenetic) metacercariae (Becker and Brunson, 1966). Another report involves Oncorhynchus kisutch in Oregon, which hatch in March and become infected with Nanophyetus salmincola by mid-April (Millemann and Knapp, 1970).

D. phoxini infections in P. phoxinus in a Swiss lowland alpine lake and in a Scottish highland lake peak by summer (Muller, 1995; Barber and Crompton, 1997). The favourable ambient temperatures in boreal or alpine habitats are between 10 and 17°C (Bauer, 1959; Wooten, 1974). This represents the low winter temperatures at which activity of pulmonate snails, such as Bulinus truncatus and Lymnaea (= Stagnicola) palustris, is interrupted in the south-eastern Mediterranean (Yekutiel, 1985; Farstey, 1986). The latter snails live in aquatic habitats fringing Lake Kinneret. M. tuberculata, which inhabits the lake proper, may also be found in deeper waters in the winter. During winter, some snails retain sporocysts that contain xiphidiocercariae, and some rediae may have pleurolophocercous cercariae (Farstey, 1986). Shedding of Bolbophorus damnificus cercariae by its snail, Planorbella trivolis, is also temperature dependent (Terhune et al., 2002). The pulmonates inhabiting the flood pools fringing the lake are more susceptible to annual flood-drought transitions in their habitat. Successive years of drought and flooding resulted in the elimination or reduction of pulmonate snails and the disappearance of metacercariae of Neascus, Bolbophorus levantinus, Clinostomum tilapiae and Euclinostomum heterostomum from the lake-dwelling cichlids for several years (Dzikowski et al., 2003a). However, infections transmitted by lake-inhabiting snails (M. tuberculata transmitting Centrocestus and Haplorchis spp., Melanopsis costata transmitting Pygidiopsis genata and Lymnasa (= Radix) auricularia transmitting Clinostomum complanatum) are not affected (Paperna, 1964a; Yekutiel, 1985; Farstey, 1986; Finkelman, 1988; Dzikowski et al., 2003a). In north-east Thailand, the seasonal pattern of Opistorchis viverini occurrence in cyprinid fish fluctuates between high abundance during the rainy season and a low during the dry season ('winter'). The number of metacercariae in fish is often positively associated with infection levels in humans (Sithithaworn et al., 1997). Recruitment of Haplorchis taichui in north Thailand is highest during the dry season (Sukontason et al., 1999).

P. conica in marine lagoons fringing the south-eastern Mediterranean and the northern gulfs of the Red Sea continues to shed cercariae (of Heterophyes and others) throughout the winter months, when water temperatures of fringing and landlocked sites may drop to below 10ºC (Taraschewski and Paperna, 1981, 1982). Year-round infection by larval digeneans has been reported in Cerithidea californica from mudflats in southern California (Martin, 1955). In the perennial habitats of the East African lakes, the M. tuberculata-transmitted metacercariae (Centrocestus and Haplorchis) and a variety of pigmented skin metacercariae (Bulinustransmitted) accumulate uninterruptedly until the young cichlids migrate to deeper waters (Paperna, 1996). A similar year-round recruitment occurs in India of C. formosanus and of Postdiplostomum grayii in Apocheilus panchax (Madhavi and Rukmini, 1991, 1992) and in China of C. formosanus in grass carp (Zeng and Liao, 2000).

Infections in farmed fishes

Extensive fish ponds and dam reservoirs, with usually a low fish biomass and established hydrophilic vegetation, provide a stable habitat for freshwater snails, which include vectors of Sanguinicola (blood flukes) and metacercariae harmful to fish, notably eye flukes. These habitats are also attractive to piscivorous birds. In warm latitudes, as in Israel, intensive-culture earth ponds, cultivating mainly carp, with their heavy organic and nitrogenous load and muddy (eutrophic) bottom are considered unfavourable habitats for all snails. Omnivorous fish, such as carp and siluroid catfishes, eat thin-shelled snails and snail eggs. Metacercarial infections are episodal only when ponds become obstructed from their routine intensive cultivation (Paperna, 1996). Diversification of the fish culture routine - introduction of new species with growing emphasis on cichlid fish ('tilapia') culture - will result in gradual adjustments of the water holding facilities. Smaller fish ponds with a firmer substrate (earth or gravel), often plastic-sheltered during winter, will provide an excellent environment for snails (predominantly M. tuberculata) to proliferate. These snails are good intermediate hosts for a whole range of heterophyid parasites, in particular Centrocestus spp. The new structures do not prevent entry of piscivorous birds that are definitive hosts of these trematodes (egrets, night herons and cormorants). Further intensification has led to the employment of circulation systems - earth and concrete raceways and circular ponds - which additionally favour proliferation of M. tuberculata, B. truncatus and sometimes species of Lymnaea (Paperna, 1996).

In the USA, circulation systems such as raceways and hatcheries similarly become heavily populated with snails (Stables and Chappell, 1986), but transmission is often limited to sanguinicolids (Wales, 1958; Hoffman et al., 1985). Metacercarial infections are usually prevented when piscivorous birds are excluded because of an efficient net system.

Blood flukes (Sanguinicola spp.) were implicated in massive mortalities of hatchery rainbow, cut-throat (Salmo clarki) and brook trout (Salvelinus fontinalis) after their snail hosts Oxytrema spp., Flumincola spp. and Leptoxis (Mudalia) spp. became established in the culture system (Davis et al., 1961; Evans, 1974b; Hoffman et al., 1985). S. inermis (transmitted by Lymnaea spp.) has been reported in pond-reared common carp in Eastern Europe (Lucky, 1964). Anderson and Shaharom-Harrison (1986) reported the introduction of Sanguinicola armata with infected bighead carp (Aristichthys nobilis) and grass carp (Ctenopharyngodon idellus) into fish farms in tropical Malaysia. An unidentified blood fluke became established in cichlids reared in circulation systems in Israel (Paperna, 1996).

Massive metacercarial infections of the gills by Centrocestus sp. and of subcutaneous tissues by Haplorchis are transmitted by M. tuberculata (Sommerville, 1982; Paperna, 1996). Gill infections by C. formosanus have resulted in mass mortality in farmed Japanese eels (Anguilla joponica) in Japan (Yanohara and Kagei, 1983). Common carp fry are also affected by Centrocestus sp. in Indian fish farms (Mohan et al., 1999). Infections have been reported from grass carp (Ctenopharyngodon idella) in China (Zeng and Liao, 2000) and from Puntius spp. and other cyprinids in north Thailand, where the infection coincided with H. pumilio, H. taichui and the human bile fluke O. viverini (Sukontason et al., 1999). Another troublesome heterophyid, Phagicola longa, accumulates in the truncus arteriosus in farmed cichlids in Israel. Its snail intermediate host is unknown. Skin Neascus ('black spot'), muscle infection with B. levantinus (Paperna, 1996), and visceral infections of C. tilapiae and E. heterostomum (Lombard, 1968; Britz et al., 1985; Paperna, 1996), transmitted by B. truncatus (Donges, 1974; Finkelman, 1988), interfere with growth and compromise survival of farmed cichlid fishes in Israel and in tropical and southern Africa. Visceral infections by Euclinostomum clarias are abundant in farmed and wild Clarias spp. in Nigeria (Ezenwaji and Inyang, 1998).

Lymnaea (Radix)-transmitted C. complanatum are responsible for heavy infection in farmed loach (Misgurnus anguillicaudatus) and ayu (Plecoglossus altivelis) in Taiwan, and these contribute to growth retardation and reduce survival (Liu, 1979; Lo et al., 1981). Procerovum cheni metacercariae cause severe muscle and connectivetissue infections in farmed eels in Taiwan (Ooi et al., 1999). In the USA, Mitchell et al. (1982) reported heavy mortality of commercially farmed fathead minnows (P. promelas), in Missouri from intensive visceral infection by P. minimum minimum. The snail involved is Physa.

Trout in farms in Europe are often troubled by eye infections with D. spathaceum, which cause blindness (Shariff et al., 1980; Stables and Chappell, 1986). The snails, Lymnaea spp., thrive in both earth ponds and raceways. Ocular diplostomiasis also affects farmed channel catfish (I. punctatus) in the southern USA (Rogers, 1972). Diplostomatid eye infections have also been reported from pond-reared largemouth bass (Micropterus salmoides) and rainbow trout in South Africa (Lombard, 1968). Ocular and brain infections of diplostomatid metacercariae in introduced cichlids (Oreochromis spp.) have been reported from Mexico (Garcia Luis et al., 1993).

The snail Planorbella, vector of Bolbophorus spp. and C. marginatum (Lorio, 1989), often occur in channel catfish (I. punctatus) ponds (Overstreet et al., 2002; Terhune et al., 2002). Metacercariae of two Bolbophorus spp. cause severe losses among pond-reared young channel catfish in the southern USA and infected fish are often rejected by commercial processors (Terhune et al., 2002). B. damnificus is vectored by the snail P. trivolis, and its definitive host is the American white pelican (Pelecanus erythrorhynchos). The life history of the second species, in pelicans and causing protodiplostomulum-stage infections in catfish, awaits further studies (Overstreet et al., 2002). Metacercariae of C. marginatum in the flesh of farmed channel catfish in the southern USA pose a potentially similar marketing problem for fish farmers (Lorio, 1989). Infection is also found in other farmed fish, such as hybrid bass (Mitchell, 1995). There is also a report of clinostomiasis (reported as C. complanatum, apparently C. marginatum (Dzikowski et al., 2004a) in introduced Oreochromis spp. from Mexico (Garcia Luis et al., 1993).

Metacercarial exotic infections, most notoriously C. formosanus and H. pumilio, have been dispersed through the introduction of the Afro-Asian snail M. tuberculata and have become a health risk for farmed fish in many water bodies in the USA and Mexico (Mitchell et al., 2000; Scholtz et al., 2001; Brandt southwest.fws.gov/fishery/ trematode.pdf). Four aquacultured fish species (I. punctatus, hybrid Morone, N. crysoleucas and P. promelas) are readily infected when exposed to C. formosanus cercariae (Mitchell et al., 2002). In Mexico infections have spread to the common carp (Velez Hernandez et al., 1998).

In the marine environment, in the eastern Mediterranean, extensively used seawater ponds often support dense populations of P. conica (Taraschewski and Paperna, 1981) and in the western Mediterranean Hydrobia spp. (Maillard et al., 1980). P. conica is first intermediate host of a number of heterophyids, including Heterophyes heterophyes, which primarily infect grey mullets (Mugilidae), but also cichlids and D. labrax (Paperna and Overstreet, 1981). Sparus auratus postlarvae, in a culture system in southern France, succumbed to massive infection with Acanthostomum imbutiforme metacercariae transmitted via Hydrobia acuta (Maillard et al., 1980; Euzet and Raibaut, 1985). Less information is available on metacercarial infection in cage-cultured fish: Lysne et al. (1994) report Cryptocotyle spp. infections in caged Atlantic cod.

Zoonoses and Food Safety

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Marketing food fish aspects

Fish containing conspicuously visible (large or coloured) free or encysted worms are rejected by consumers (Kabunda and Sommerville, 1984; Lorio, 1989; Paperna, 1996; Overstreet et al., 2002). Fish with badly damaged eyes (trout affected by eye diplostomiasis) are likely to be sold at depreciated prices. The mere suspicion of infection affects demand and depreciates market prices of the entire consignment of fish. Examples are the large yellow clinostomids, which also tend to encyst and crawl around after the death of the fish (Fig. 19C), metacercariae of diplostomatids in muscles (Fig. 17B); (Paperna, 1996), 'black spot' (Fig. 19A, B) and fish containing large didymozoids in skin and gills or in the muscles (Lester, 1980; Fig. 8). (For risk to public health, see Library Document "Fish-borne parasitic zoonoses".)

Fig. 8. Didymozoidae. A. Palate of Platycephalus fuscus infected with Neometadidymozoon helicis (in life, bright yellow). B. Lobatozoum multidacculatum on gills of Katsumonus pelamis, New Zealand. C. Larval stages of didymozoids encysted on the surface of the intestine of Favonigobius exquisitus. D. Section of Nematobothrium spinneri in the body wall muscle of Acanthocybium solndri, Queensland. E. N. helicis capsule in the body wall of P. fuscus.

Fig. 17. Diplostomatid metacercariae: A. Diplostomum sp. removed from a visceral cyst in Clarias gariepinus, Uganda. B. Heavy Bolbophorus levantinum metacercaria in muscles of young Oreochromis aurea from Lake Kinnereth. C. Early-stage B. levantinum metacercaria with expanded posterior half. D. Later-stage B. levantinum metacercaria with emptied posterior end (actual size 0.6-0.8 mm).

Fig. 19. Metacercarial skin infection in Lake Kinnereth cichlids. A, B. Low and heavy 'black spot' (Neascus) infection in young pond-reared Oreochromis aurea × nilotica. C. Clinostomum 'cutaneus'-infected Tristramella simonis (top) and Tilapia zilli (bottom).

References

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