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Myxozoan infections of fish

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Myxozoan infections of fish

Summary

  • Last modified
  • 05 September 2018
  • Datasheet Type(s)
  • Animal Disease
  • Preferred Scientific Name
  • Myxozoan infections of fish
  • Overview
  • Myxozoans are highly specialized metazoan parasites of aquatic hosts with a very wide host range. This diverse group of organisms is characterized by multicellular spores with polar capsules containing extrudable polar...

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Line drawings of representative myxospore genera showing key morphological features (not to scale). A. Ceratomyxa, B. Henneguya, C. Myxobilatus, D. Hoferellus, E. Myxobolus, F. Sphaerospora, G. Kudoa, H. Chloromyxum, I. Parvicapsula, J. Sphaeromyxa, K. Myxidium, L. Thelohanellus, M. Unicapsula.
TitleMorphological features of myxospore genera
CaptionLine drawings of representative myxospore genera showing key morphological features (not to scale). A. Ceratomyxa, B. Henneguya, C. Myxobilatus, D. Hoferellus, E. Myxobolus, F. Sphaerospora, G. Kudoa, H. Chloromyxum, I. Parvicapsula, J. Sphaeromyxa, K. Myxidium, L. Thelohanellus, M. Unicapsula.
CopyrightStephen W. Feist & Matt Longshaw
Line drawings of representative myxospore genera showing key morphological features (not to scale). A. Ceratomyxa, B. Henneguya, C. Myxobilatus, D. Hoferellus, E. Myxobolus, F. Sphaerospora, G. Kudoa, H. Chloromyxum, I. Parvicapsula, J. Sphaeromyxa, K. Myxidium, L. Thelohanellus, M. Unicapsula.
Morphological features of myxospore generaLine drawings of representative myxospore genera showing key morphological features (not to scale). A. Ceratomyxa, B. Henneguya, C. Myxobilatus, D. Hoferellus, E. Myxobolus, F. Sphaerospora, G. Kudoa, H. Chloromyxum, I. Parvicapsula, J. Sphaeromyxa, K. Myxidium, L. Thelohanellus, M. Unicapsula.Stephen W. Feist & Matt Longshaw
Line drawings of the actinospore collective groups showing key morphological features (not to scale). A. Endocapsa (after Hallett et al., 1999), B. Sphaeractinomyxon (after Hallett et al., 1999), C. Tetraspora (after Hallett and Lester, 1999), D. Tetractinomyxon, E. Aurantiactinomyxon, F. Neoactinomyxum (after Ormières and Frézil, 1969), G. Neoactinomyxum (after El-Mansy et al., 1998a), H. Guyenotia (after Naville, 1930), I. Echinactinomyxon, J. Raabeia, K. Triactinomyxon, L. Pseudotriactinomyxon (after Lowers and Bartholomew, 2003), M. Hexactinomyxon (after Janiszewska, 1957), N. Ormieractinomyxon (after Marques, 1984), O. Siedleckiella (after Janiszewska, 1955), inset, collection of eight spores forming characteristic net, P. Synactinomyxon (after McGeorge et al., 1997), inset, individual spore shown, Q. Synactinomyxon, R. Antonactinomyxon (after Janiszewska, 1955).
TitleMorphological features of actiospore collective groups
CaptionLine drawings of the actinospore collective groups showing key morphological features (not to scale). A. Endocapsa (after Hallett et al., 1999), B. Sphaeractinomyxon (after Hallett et al., 1999), C. Tetraspora (after Hallett and Lester, 1999), D. Tetractinomyxon, E. Aurantiactinomyxon, F. Neoactinomyxum (after Ormières and Frézil, 1969), G. Neoactinomyxum (after El-Mansy et al., 1998a), H. Guyenotia (after Naville, 1930), I. Echinactinomyxon, J. Raabeia, K. Triactinomyxon, L. Pseudotriactinomyxon (after Lowers and Bartholomew, 2003), M. Hexactinomyxon (after Janiszewska, 1957), N. Ormieractinomyxon (after Marques, 1984), O. Siedleckiella (after Janiszewska, 1955), inset, collection of eight spores forming characteristic net, P. Synactinomyxon (after McGeorge et al., 1997), inset, individual spore shown, Q. Synactinomyxon, R. Antonactinomyxon (after Janiszewska, 1955).
CopyrightStephen W. Feist & Matt Longshaw
Line drawings of the actinospore collective groups showing key morphological features (not to scale). A. Endocapsa (after Hallett et al., 1999), B. Sphaeractinomyxon (after Hallett et al., 1999), C. Tetraspora (after Hallett and Lester, 1999), D. Tetractinomyxon, E. Aurantiactinomyxon, F. Neoactinomyxum (after Ormières and Frézil, 1969), G. Neoactinomyxum (after El-Mansy et al., 1998a), H. Guyenotia (after Naville, 1930), I. Echinactinomyxon, J. Raabeia, K. Triactinomyxon, L. Pseudotriactinomyxon (after Lowers and Bartholomew, 2003), M. Hexactinomyxon (after Janiszewska, 1957), N. Ormieractinomyxon (after Marques, 1984), O. Siedleckiella (after Janiszewska, 1955), inset, collection of eight spores forming characteristic net, P. Synactinomyxon (after McGeorge et al., 1997), inset, individual spore shown, Q. Synactinomyxon, R. Antonactinomyxon (after Janiszewska, 1955).
Morphological features of actiospore collective groupsLine drawings of the actinospore collective groups showing key morphological features (not to scale). A. Endocapsa (after Hallett et al., 1999), B. Sphaeractinomyxon (after Hallett et al., 1999), C. Tetraspora (after Hallett and Lester, 1999), D. Tetractinomyxon, E. Aurantiactinomyxon, F. Neoactinomyxum (after Ormières and Frézil, 1969), G. Neoactinomyxum (after El-Mansy et al., 1998a), H. Guyenotia (after Naville, 1930), I. Echinactinomyxon, J. Raabeia, K. Triactinomyxon, L. Pseudotriactinomyxon (after Lowers and Bartholomew, 2003), M. Hexactinomyxon (after Janiszewska, 1957), N. Ormieractinomyxon (after Marques, 1984), O. Siedleckiella (after Janiszewska, 1955), inset, collection of eight spores forming characteristic net, P. Synactinomyxon (after McGeorge et al., 1997), inset, individual spore shown, Q. Synactinomyxon, R. Antonactinomyxon (after Janiszewska, 1955).Stephen W. Feist & Matt Longshaw
Photomicrographs of representative myxospore genera. A. Henneguya zschokkei from coho salmon (Oncorhynchus kisutch), B. Henneguya psorospermica from pike (Esox lucius) gills, C. Thelohanellus pyriformis from gill of tench (Tinca tinca), D. Chloromyxum sp. from P. phoxinus gall bladder, E. Leptotheca sp. from gall bladder of tadpole fish (Raniceps raninus), F. Sphaerospora elegans from stickleback (Gasterosteus aculeatus) kidney, G. Parvicapsula assymetrica from Cyclopterus lumpus urinary bladder, H. Kudoa thyrsites from scabbardfish (Lepidopus caudatus) muscle, I. Myxoproteus ambiguus from the urinary bladder of anglerfish (Lophius piscatorius).
TitlePhotomicrographs of representative myxospore genera
CaptionPhotomicrographs of representative myxospore genera. A. Henneguya zschokkei from coho salmon (Oncorhynchus kisutch), B. Henneguya psorospermica from pike (Esox lucius) gills, C. Thelohanellus pyriformis from gill of tench (Tinca tinca), D. Chloromyxum sp. from P. phoxinus gall bladder, E. Leptotheca sp. from gall bladder of tadpole fish (Raniceps raninus), F. Sphaerospora elegans from stickleback (Gasterosteus aculeatus) kidney, G. Parvicapsula assymetrica from Cyclopterus lumpus urinary bladder, H. Kudoa thyrsites from scabbardfish (Lepidopus caudatus) muscle, I. Myxoproteus ambiguus from the urinary bladder of anglerfish (Lophius piscatorius).
CopyrightStephen W. Feist & Matt Longshaw
Photomicrographs of representative myxospore genera. A. Henneguya zschokkei from coho salmon (Oncorhynchus kisutch), B. Henneguya psorospermica from pike (Esox lucius) gills, C. Thelohanellus pyriformis from gill of tench (Tinca tinca), D. Chloromyxum sp. from P. phoxinus gall bladder, E. Leptotheca sp. from gall bladder of tadpole fish (Raniceps raninus), F. Sphaerospora elegans from stickleback (Gasterosteus aculeatus) kidney, G. Parvicapsula assymetrica from Cyclopterus lumpus urinary bladder, H. Kudoa thyrsites from scabbardfish (Lepidopus caudatus) muscle, I. Myxoproteus ambiguus from the urinary bladder of anglerfish (Lophius piscatorius).
Photomicrographs of representative myxospore generaPhotomicrographs of representative myxospore genera. A. Henneguya zschokkei from coho salmon (Oncorhynchus kisutch), B. Henneguya psorospermica from pike (Esox lucius) gills, C. Thelohanellus pyriformis from gill of tench (Tinca tinca), D. Chloromyxum sp. from P. phoxinus gall bladder, E. Leptotheca sp. from gall bladder of tadpole fish (Raniceps raninus), F. Sphaerospora elegans from stickleback (Gasterosteus aculeatus) kidney, G. Parvicapsula assymetrica from Cyclopterus lumpus urinary bladder, H. Kudoa thyrsites from scabbardfish (Lepidopus caudatus) muscle, I. Myxoproteus ambiguus from the urinary bladder of anglerfish (Lophius piscatorius).Stephen W. Feist & Matt Longshaw
Photomicrographs of representative myxospore genera. J. Myxobilatus gasterostei from the kidney of stickleback Gasterosteus aculeatus, K. Myxidium sp. from the gall bladder of rudd (Scardinius erythrophthalmus), L. Ceratomyxa sp. from common goby (Pomatoschistus microps), M. Myxidium gadi from the gall bladder of whiting (Merlangius merlangus), N. Myxobolus sp. from dace (Leuciscus leuciscus) buccal cavity cyst, O. Sphaeromyxa sp. from two-spot goby (Gobiosculus flavescens) gall bladder.
TitlePhotomicrographs of representative myxospore genera
CaptionPhotomicrographs of representative myxospore genera. J. Myxobilatus gasterostei from the kidney of stickleback Gasterosteus aculeatus, K. Myxidium sp. from the gall bladder of rudd (Scardinius erythrophthalmus), L. Ceratomyxa sp. from common goby (Pomatoschistus microps), M. Myxidium gadi from the gall bladder of whiting (Merlangius merlangus), N. Myxobolus sp. from dace (Leuciscus leuciscus) buccal cavity cyst, O. Sphaeromyxa sp. from two-spot goby (Gobiosculus flavescens) gall bladder.
CopyrightStephen W. Feist & Matt Longshaw
Photomicrographs of representative myxospore genera. J. Myxobilatus gasterostei from the kidney of stickleback Gasterosteus aculeatus, K. Myxidium sp. from the gall bladder of rudd (Scardinius erythrophthalmus), L. Ceratomyxa sp. from common goby (Pomatoschistus microps), M. Myxidium gadi from the gall bladder of whiting (Merlangius merlangus), N. Myxobolus sp. from dace (Leuciscus leuciscus) buccal cavity cyst, O. Sphaeromyxa sp. from two-spot goby (Gobiosculus flavescens) gall bladder.
Photomicrographs of representative myxospore generaPhotomicrographs of representative myxospore genera. J. Myxobilatus gasterostei from the kidney of stickleback Gasterosteus aculeatus, K. Myxidium sp. from the gall bladder of rudd (Scardinius erythrophthalmus), L. Ceratomyxa sp. from common goby (Pomatoschistus microps), M. Myxidium gadi from the gall bladder of whiting (Merlangius merlangus), N. Myxobolus sp. from dace (Leuciscus leuciscus) buccal cavity cyst, O. Sphaeromyxa sp. from two-spot goby (Gobiosculus flavescens) gall bladder.Stephen W. Feist & Matt Longshaw
Myxosporean extrasporogonic and plasmodial stages. Fresh preparations unless otherwise stated. A. Myxobolus artus plasmodium encysted in the renal tissue of koi carp (Cyprinus carpio). B. Gill cysts of Myxobolus macrocapsularis in juvenile chub, Leuciscus cephalus. C. Sporogonic plasmodium of Zschokkella sp. from the gall bladder of three-bearded rockling (Gaidropsaurus vulgaris). D. May-Grünwald Giemsa-stained smear of a Myxidium incurvatum plasmodium from the gall bladder of flounder (Platichthys flesus). E. Sphaeromyxa sp. plasmodia contained within the gall bladder of two-spot goby (Gobiusculus flavescens). F. Sphaerospora truttae pseudoplasmodia within renal tubule of brown trout (Salmo trutta). G. Plasmodium of Myxobilatus gasterostei containing two mature spores. From the kidney of the three-spined stickleback (Gasterosteus aculeatus). H. Extrasporogonic stage from the rete mirabile in the eye of G. aculeatus. I. Giemsa-stained section of the rete mirabile with the parasites loc
TitleMyxosporean extrasporogonic and plasmodial stages
CaptionMyxosporean extrasporogonic and plasmodial stages. Fresh preparations unless otherwise stated. A. Myxobolus artus plasmodium encysted in the renal tissue of koi carp (Cyprinus carpio). B. Gill cysts of Myxobolus macrocapsularis in juvenile chub, Leuciscus cephalus. C. Sporogonic plasmodium of Zschokkella sp. from the gall bladder of three-bearded rockling (Gaidropsaurus vulgaris). D. May-Grünwald Giemsa-stained smear of a Myxidium incurvatum plasmodium from the gall bladder of flounder (Platichthys flesus). E. Sphaeromyxa sp. plasmodia contained within the gall bladder of two-spot goby (Gobiusculus flavescens). F. Sphaerospora truttae pseudoplasmodia within renal tubule of brown trout (Salmo trutta). G. Plasmodium of Myxobilatus gasterostei containing two mature spores. From the kidney of the three-spined stickleback (Gasterosteus aculeatus). H. Extrasporogonic stage from the rete mirabile in the eye of G. aculeatus. I. Giemsa-stained section of the rete mirabile with the parasites loc
CopyrightStephen W. Feist & Matt Longshaw
Myxosporean extrasporogonic and plasmodial stages. Fresh preparations unless otherwise stated. A. Myxobolus artus plasmodium encysted in the renal tissue of koi carp (Cyprinus carpio). B. Gill cysts of Myxobolus macrocapsularis in juvenile chub, Leuciscus cephalus. C. Sporogonic plasmodium of Zschokkella sp. from the gall bladder of three-bearded rockling (Gaidropsaurus vulgaris). D. May-Grünwald Giemsa-stained smear of a Myxidium incurvatum plasmodium from the gall bladder of flounder (Platichthys flesus). E. Sphaeromyxa sp. plasmodia contained within the gall bladder of two-spot goby (Gobiusculus flavescens). F. Sphaerospora truttae pseudoplasmodia within renal tubule of brown trout (Salmo trutta). G. Plasmodium of Myxobilatus gasterostei containing two mature spores. From the kidney of the three-spined stickleback (Gasterosteus aculeatus). H. Extrasporogonic stage from the rete mirabile in the eye of G. aculeatus. I. Giemsa-stained section of the rete mirabile with the parasites loc
Myxosporean extrasporogonic and plasmodial stagesMyxosporean extrasporogonic and plasmodial stages. Fresh preparations unless otherwise stated. A. Myxobolus artus plasmodium encysted in the renal tissue of koi carp (Cyprinus carpio). B. Gill cysts of Myxobolus macrocapsularis in juvenile chub, Leuciscus cephalus. C. Sporogonic plasmodium of Zschokkella sp. from the gall bladder of three-bearded rockling (Gaidropsaurus vulgaris). D. May-Grünwald Giemsa-stained smear of a Myxidium incurvatum plasmodium from the gall bladder of flounder (Platichthys flesus). E. Sphaeromyxa sp. plasmodia contained within the gall bladder of two-spot goby (Gobiusculus flavescens). F. Sphaerospora truttae pseudoplasmodia within renal tubule of brown trout (Salmo trutta). G. Plasmodium of Myxobilatus gasterostei containing two mature spores. From the kidney of the three-spined stickleback (Gasterosteus aculeatus). H. Extrasporogonic stage from the rete mirabile in the eye of G. aculeatus. I. Giemsa-stained section of the rete mirabile with the parasites locStephen W. Feist & Matt Longshaw
Myxosporean extrasporogonic and plasmodial stages. J. Phase-contrast image of a plasmodium of Myxidium lieberkuehni from the urinary bladder of pike (Esox lucius), showing the characteristic villous projections on the surface of the plasmodium. K. Interference-contrast image of M. lieber-kuehni plasmodia showing the clear ectoplasmic layer. L. Cyst of Myxidium rhodei from the kidney of roach (Rutilus rutilus). M. Presporogonic stages of Sphaerospora elegans in Bowman's space of the glomerulus in the kidney of G. aculeatus. N. Numerous plasmodia attached to the epithelium of the urinary bladder of a juvenile dace (Leuciscus leuciscus).
TitleMyxosporean extrasporogonic and plasmodial stages
CaptionMyxosporean extrasporogonic and plasmodial stages. J. Phase-contrast image of a plasmodium of Myxidium lieberkuehni from the urinary bladder of pike (Esox lucius), showing the characteristic villous projections on the surface of the plasmodium. K. Interference-contrast image of M. lieber-kuehni plasmodia showing the clear ectoplasmic layer. L. Cyst of Myxidium rhodei from the kidney of roach (Rutilus rutilus). M. Presporogonic stages of Sphaerospora elegans in Bowman's space of the glomerulus in the kidney of G. aculeatus. N. Numerous plasmodia attached to the epithelium of the urinary bladder of a juvenile dace (Leuciscus leuciscus).
CopyrightStephen W. Feist & Matt Longshaw
Myxosporean extrasporogonic and plasmodial stages. J. Phase-contrast image of a plasmodium of Myxidium lieberkuehni from the urinary bladder of pike (Esox lucius), showing the characteristic villous projections on the surface of the plasmodium. K. Interference-contrast image of M. lieber-kuehni plasmodia showing the clear ectoplasmic layer. L. Cyst of Myxidium rhodei from the kidney of roach (Rutilus rutilus). M. Presporogonic stages of Sphaerospora elegans in Bowman's space of the glomerulus in the kidney of G. aculeatus. N. Numerous plasmodia attached to the epithelium of the urinary bladder of a juvenile dace (Leuciscus leuciscus).
Myxosporean extrasporogonic and plasmodial stagesMyxosporean extrasporogonic and plasmodial stages. J. Phase-contrast image of a plasmodium of Myxidium lieberkuehni from the urinary bladder of pike (Esox lucius), showing the characteristic villous projections on the surface of the plasmodium. K. Interference-contrast image of M. lieber-kuehni plasmodia showing the clear ectoplasmic layer. L. Cyst of Myxidium rhodei from the kidney of roach (Rutilus rutilus). M. Presporogonic stages of Sphaerospora elegans in Bowman's space of the glomerulus in the kidney of G. aculeatus. N. Numerous plasmodia attached to the epithelium of the urinary bladder of a juvenile dace (Leuciscus leuciscus).Stephen W. Feist & Matt Longshaw
A-C. Deformed myxospores; D-I. Actinospores released from oligochaetes. A. Giemsa-stained smear of Myxidium giardi spores from eel Anguilla anguilla, one of which contains three polar capsules. B. Deformed Thelohanellus pyriformis spore from gill of tench (Tinca tinca). C. Triradiate spores of Ceratomyxa sp. from the gall bladder of common goby (Pomatoschistus microps). D. Aurantiactinomyxon-type actinospore. E. Echinactinomyxon-type actinospore. F. Neoactinomyxum-type actinospore. G. Triactinomyxon-type actinospore. Note presence of large style and sporoplasm and polar capsules at apex of spore. H. Collection of Synactinomyxon-type actinospores. I. Secondary cells released from the sporoplasm of a Triactinomyxon-type actinospore.
TitleDeformed myxospores; Actinospores released from oligochaetes
CaptionA-C. Deformed myxospores; D-I. Actinospores released from oligochaetes. A. Giemsa-stained smear of Myxidium giardi spores from eel Anguilla anguilla, one of which contains three polar capsules. B. Deformed Thelohanellus pyriformis spore from gill of tench (Tinca tinca). C. Triradiate spores of Ceratomyxa sp. from the gall bladder of common goby (Pomatoschistus microps). D. Aurantiactinomyxon-type actinospore. E. Echinactinomyxon-type actinospore. F. Neoactinomyxum-type actinospore. G. Triactinomyxon-type actinospore. Note presence of large style and sporoplasm and polar capsules at apex of spore. H. Collection of Synactinomyxon-type actinospores. I. Secondary cells released from the sporoplasm of a Triactinomyxon-type actinospore.
CopyrightStephen W. Feist & Matt Longshaw
A-C. Deformed myxospores; D-I. Actinospores released from oligochaetes. A. Giemsa-stained smear of Myxidium giardi spores from eel Anguilla anguilla, one of which contains three polar capsules. B. Deformed Thelohanellus pyriformis spore from gill of tench (Tinca tinca). C. Triradiate spores of Ceratomyxa sp. from the gall bladder of common goby (Pomatoschistus microps). D. Aurantiactinomyxon-type actinospore. E. Echinactinomyxon-type actinospore. F. Neoactinomyxum-type actinospore. G. Triactinomyxon-type actinospore. Note presence of large style and sporoplasm and polar capsules at apex of spore. H. Collection of Synactinomyxon-type actinospores. I. Secondary cells released from the sporoplasm of a Triactinomyxon-type actinospore.
Deformed myxospores; Actinospores released from oligochaetesA-C. Deformed myxospores; D-I. Actinospores released from oligochaetes. A. Giemsa-stained smear of Myxidium giardi spores from eel Anguilla anguilla, one of which contains three polar capsules. B. Deformed Thelohanellus pyriformis spore from gill of tench (Tinca tinca). C. Triradiate spores of Ceratomyxa sp. from the gall bladder of common goby (Pomatoschistus microps). D. Aurantiactinomyxon-type actinospore. E. Echinactinomyxon-type actinospore. F. Neoactinomyxum-type actinospore. G. Triactinomyxon-type actinospore. Note presence of large style and sporoplasm and polar capsules at apex of spore. H. Collection of Synactinomyxon-type actinospores. I. Secondary cells released from the sporoplasm of a Triactinomyxon-type actinospore.Stephen W. Feist & Matt Longshaw
Sporogonic stages of Tetracapsuloides bryosalmonae. A. Fresh spore of T. bryosalmonae showing the four polar capsules and two sporoplasm cells surrounded by valvogenic cells. B. Diagrams of T. bryosalmonae spores in three-dimensional view and apical view. Note presence of four capsular cells and eight valvular cells. C. Section through a complete spore showing the sporoplasm cells with each containing a secondary cell. A single polar capsule can also be seen. Inset: Characteristic sporoplasmosome within the cytoplasm of the sporoplasm cell, showing the typical bar-like invagination, also seen in the histozoic fish stage of the parasite. D. Sacs of T. bryosalmonae released from the bryozoan host. E. Section through the polar capsule showing the exit pore for the filament and characteristic reticulated cap. Note the gap between the valvogenic cells at the exit pore of the polar filament. F. As E, showing the nucleus of the capsulogenic cell and sections through the coiled polar filament.
TitleSporogonic stages of Tetracapsuloides bryosalmonae
CaptionSporogonic stages of Tetracapsuloides bryosalmonae. A. Fresh spore of T. bryosalmonae showing the four polar capsules and two sporoplasm cells surrounded by valvogenic cells. B. Diagrams of T. bryosalmonae spores in three-dimensional view and apical view. Note presence of four capsular cells and eight valvular cells. C. Section through a complete spore showing the sporoplasm cells with each containing a secondary cell. A single polar capsule can also be seen. Inset: Characteristic sporoplasmosome within the cytoplasm of the sporoplasm cell, showing the typical bar-like invagination, also seen in the histozoic fish stage of the parasite. D. Sacs of T. bryosalmonae released from the bryozoan host. E. Section through the polar capsule showing the exit pore for the filament and characteristic reticulated cap. Note the gap between the valvogenic cells at the exit pore of the polar filament. F. As E, showing the nucleus of the capsulogenic cell and sections through the coiled polar filament.
CopyrightStephen W. Feist & Matt Longshaw
Sporogonic stages of Tetracapsuloides bryosalmonae. A. Fresh spore of T. bryosalmonae showing the four polar capsules and two sporoplasm cells surrounded by valvogenic cells. B. Diagrams of T. bryosalmonae spores in three-dimensional view and apical view. Note presence of four capsular cells and eight valvular cells. C. Section through a complete spore showing the sporoplasm cells with each containing a secondary cell. A single polar capsule can also be seen. Inset: Characteristic sporoplasmosome within the cytoplasm of the sporoplasm cell, showing the typical bar-like invagination, also seen in the histozoic fish stage of the parasite. D. Sacs of T. bryosalmonae released from the bryozoan host. E. Section through the polar capsule showing the exit pore for the filament and characteristic reticulated cap. Note the gap between the valvogenic cells at the exit pore of the polar filament. F. As E, showing the nucleus of the capsulogenic cell and sections through the coiled polar filament.
Sporogonic stages of Tetracapsuloides bryosalmonaeSporogonic stages of Tetracapsuloides bryosalmonae. A. Fresh spore of T. bryosalmonae showing the four polar capsules and two sporoplasm cells surrounded by valvogenic cells. B. Diagrams of T. bryosalmonae spores in three-dimensional view and apical view. Note presence of four capsular cells and eight valvular cells. C. Section through a complete spore showing the sporoplasm cells with each containing a secondary cell. A single polar capsule can also be seen. Inset: Characteristic sporoplasmosome within the cytoplasm of the sporoplasm cell, showing the typical bar-like invagination, also seen in the histozoic fish stage of the parasite. D. Sacs of T. bryosalmonae released from the bryozoan host. E. Section through the polar capsule showing the exit pore for the filament and characteristic reticulated cap. Note the gap between the valvogenic cells at the exit pore of the polar filament. F. As E, showing the nucleus of the capsulogenic cell and sections through the coiled polar filament.Stephen W. Feist & Matt Longshaw
Intra-oligochaete myxozoan life cycle (actinospore phase), where heavy arrow = life cycle of M. cerebralis demonstrated experimentally by El-Matbouli and Hoffmann (1998) and light arrow = life cycle for other actinospores. A binucleate or uninucleate sporoplasm (1) released from a myxospore following ingestion by the oligochaete undergoes a series of nuclear divisions to produce a multinucleate cell. Following plasmotomy, the cells either undergo further nuclear divisions to produce more multinucleate stages (2) or produce a two- and then four-nuclei cell (3). Plasmotomy follows to produce four cells, two of which envelop the other two cells (4). For all actinospore types, the outer cells undergo two mitotic divisions to produce a pansporocyst wall containing eight cells. For M. cerebralis and other actinospores producing eight actinospores per pansporocyst, division of the inner cells produces 16 diploid cells via three mitotic divisions (5). Following one meiotic division, 16 haploid
TitleIntra-oligochaete myxozoan life cycle (actinospore phase)
CaptionIntra-oligochaete myxozoan life cycle (actinospore phase), where heavy arrow = life cycle of M. cerebralis demonstrated experimentally by El-Matbouli and Hoffmann (1998) and light arrow = life cycle for other actinospores. A binucleate or uninucleate sporoplasm (1) released from a myxospore following ingestion by the oligochaete undergoes a series of nuclear divisions to produce a multinucleate cell. Following plasmotomy, the cells either undergo further nuclear divisions to produce more multinucleate stages (2) or produce a two- and then four-nuclei cell (3). Plasmotomy follows to produce four cells, two of which envelop the other two cells (4). For all actinospore types, the outer cells undergo two mitotic divisions to produce a pansporocyst wall containing eight cells. For M. cerebralis and other actinospores producing eight actinospores per pansporocyst, division of the inner cells produces 16 diploid cells via three mitotic divisions (5). Following one meiotic division, 16 haploid
CopyrightStephen W. Feist & Matt Longshaw
Intra-oligochaete myxozoan life cycle (actinospore phase), where heavy arrow = life cycle of M. cerebralis demonstrated experimentally by El-Matbouli and Hoffmann (1998) and light arrow = life cycle for other actinospores. A binucleate or uninucleate sporoplasm (1) released from a myxospore following ingestion by the oligochaete undergoes a series of nuclear divisions to produce a multinucleate cell. Following plasmotomy, the cells either undergo further nuclear divisions to produce more multinucleate stages (2) or produce a two- and then four-nuclei cell (3). Plasmotomy follows to produce four cells, two of which envelop the other two cells (4). For all actinospore types, the outer cells undergo two mitotic divisions to produce a pansporocyst wall containing eight cells. For M. cerebralis and other actinospores producing eight actinospores per pansporocyst, division of the inner cells produces 16 diploid cells via three mitotic divisions (5). Following one meiotic division, 16 haploid
Intra-oligochaete myxozoan life cycle (actinospore phase)Intra-oligochaete myxozoan life cycle (actinospore phase), where heavy arrow = life cycle of M. cerebralis demonstrated experimentally by El-Matbouli and Hoffmann (1998) and light arrow = life cycle for other actinospores. A binucleate or uninucleate sporoplasm (1) released from a myxospore following ingestion by the oligochaete undergoes a series of nuclear divisions to produce a multinucleate cell. Following plasmotomy, the cells either undergo further nuclear divisions to produce more multinucleate stages (2) or produce a two- and then four-nuclei cell (3). Plasmotomy follows to produce four cells, two of which envelop the other two cells (4). For all actinospore types, the outer cells undergo two mitotic divisions to produce a pansporocyst wall containing eight cells. For M. cerebralis and other actinospores producing eight actinospores per pansporocyst, division of the inner cells produces 16 diploid cells via three mitotic divisions (5). Following one meiotic division, 16 haploidStephen W. Feist & Matt Longshaw
Intra-piscine myxozoan life cycle (myxospore phase). This applies to those myxozoans undergoing a two-host life cycle. Heavy arrow = life cycle of M. cerebralis, demonstrated experimentally by El-Matbouli et al. (1995); light arrow = postulated life cycle for other myxospores.
TitleIntra-piscine myxozoan life cycle (myxospore phase)
CaptionIntra-piscine myxozoan life cycle (myxospore phase). This applies to those myxozoans undergoing a two-host life cycle. Heavy arrow = life cycle of M. cerebralis, demonstrated experimentally by El-Matbouli et al. (1995); light arrow = postulated life cycle for other myxospores.
CopyrightStephen W. Feist & Matt Longshaw
Intra-piscine myxozoan life cycle (myxospore phase). This applies to those myxozoans undergoing a two-host life cycle. Heavy arrow = life cycle of M. cerebralis, demonstrated experimentally by El-Matbouli et al. (1995); light arrow = postulated life cycle for other myxospores.
Intra-piscine myxozoan life cycle (myxospore phase)Intra-piscine myxozoan life cycle (myxospore phase). This applies to those myxozoans undergoing a two-host life cycle. Heavy arrow = life cycle of M. cerebralis, demonstrated experimentally by El-Matbouli et al. (1995); light arrow = postulated life cycle for other myxospores.Stephen W. Feist & Matt Longshaw
Histological sections of gill, pseudobranch and fin infections. A. Giemsa-stained section of carp (Cyprinus carpio) gill infected with Sphaerospora molnari (inset: mature spore in sutural view). B. Large cyst of Myxobolus koi in the gill of koi carp (C. carpio) (inset: mature spore in valvular view). C. Multiple cysts of Henneguya psorospermica in the gill of pike (Esox lucius). D. Multiple cysts of Myxobolus macrocapsularis in the gill of chub (L. cephalus) (inset: mature spore in valvular view).
TitleHistological sections of gill, pseudobranch and fin infections
CaptionHistological sections of gill, pseudobranch and fin infections. A. Giemsa-stained section of carp (Cyprinus carpio) gill infected with Sphaerospora molnari (inset: mature spore in sutural view). B. Large cyst of Myxobolus koi in the gill of koi carp (C. carpio) (inset: mature spore in valvular view). C. Multiple cysts of Henneguya psorospermica in the gill of pike (Esox lucius). D. Multiple cysts of Myxobolus macrocapsularis in the gill of chub (L. cephalus) (inset: mature spore in valvular view).
CopyrightStephen W. Feist & Matt Longshaw
Histological sections of gill, pseudobranch and fin infections. A. Giemsa-stained section of carp (Cyprinus carpio) gill infected with Sphaerospora molnari (inset: mature spore in sutural view). B. Large cyst of Myxobolus koi in the gill of koi carp (C. carpio) (inset: mature spore in valvular view). C. Multiple cysts of Henneguya psorospermica in the gill of pike (Esox lucius). D. Multiple cysts of Myxobolus macrocapsularis in the gill of chub (L. cephalus) (inset: mature spore in valvular view).
Histological sections of gill, pseudobranch and fin infectionsHistological sections of gill, pseudobranch and fin infections. A. Giemsa-stained section of carp (Cyprinus carpio) gill infected with Sphaerospora molnari (inset: mature spore in sutural view). B. Large cyst of Myxobolus koi in the gill of koi carp (C. carpio) (inset: mature spore in valvular view). C. Multiple cysts of Henneguya psorospermica in the gill of pike (Esox lucius). D. Multiple cysts of Myxobolus macrocapsularis in the gill of chub (L. cephalus) (inset: mature spore in valvular view).Stephen W. Feist & Matt Longshaw
Histological sections of gill, pseudobranch and fin infections. E. Cysts of M. macrocapsularis in the pseudobranch of dace (L. leuciscus). F. Sporogonic plasmodium of an unidentified Myxobolus sp. in the cartilage of the caudal peduncle of L. cephalus. G. Longitudinal section through the fin of a juvenile roach (R. rutilus) showing Myxobolus sp. cysts within the epithelium. H. Myxobolus sp. cysts in connective tissue of the fin of barbel (Barbus barbus).
TitleHistological sections of gill, pseudobranch and fin infections
CaptionHistological sections of gill, pseudobranch and fin infections. E. Cysts of M. macrocapsularis in the pseudobranch of dace (L. leuciscus). F. Sporogonic plasmodium of an unidentified Myxobolus sp. in the cartilage of the caudal peduncle of L. cephalus. G. Longitudinal section through the fin of a juvenile roach (R. rutilus) showing Myxobolus sp. cysts within the epithelium. H. Myxobolus sp. cysts in connective tissue of the fin of barbel (Barbus barbus).
CopyrightStephen W. Feist & Matt Longshaw
Histological sections of gill, pseudobranch and fin infections. E. Cysts of M. macrocapsularis in the pseudobranch of dace (L. leuciscus). F. Sporogonic plasmodium of an unidentified Myxobolus sp. in the cartilage of the caudal peduncle of L. cephalus. G. Longitudinal section through the fin of a juvenile roach (R. rutilus) showing Myxobolus sp. cysts within the epithelium. H. Myxobolus sp. cysts in connective tissue of the fin of barbel (Barbus barbus).
Histological sections of gill, pseudobranch and fin infectionsHistological sections of gill, pseudobranch and fin infections. E. Cysts of M. macrocapsularis in the pseudobranch of dace (L. leuciscus). F. Sporogonic plasmodium of an unidentified Myxobolus sp. in the cartilage of the caudal peduncle of L. cephalus. G. Longitudinal section through the fin of a juvenile roach (R. rutilus) showing Myxobolus sp. cysts within the epithelium. H. Myxobolus sp. cysts in connective tissue of the fin of barbel (Barbus barbus).Stephen W. Feist & Matt Longshaw
Histological features of muscle-invading species. A. Sporogonic plasmodia of Myxobolus artus in the skeletal muscle of carp (C. carpio) (inset: mature spore in valvular view). B. Granulomatous host response to a plasmodium of M. artus within the renal tissue (same fish as in Fig. 8.10A). C. A plasmodium of Myxobolus pseudodispar within the muscle of a juvenile roach (Rutilus rutilus). D. Focal inflammatory response to M. pseudodispar spores and sporogonic stages from a ruptured plasmodium. E. Intra-myofibrillar Kudoa infection in cow-nosed ray (Rhinoptera bonasus). F. Multiple plasmodia of Kudoa sp. in the myofibrils of common goby (Pomatoschistus microps). G. Giemsa-stained section of Kudoa thyrsites infection in cod (Gadus morhua). H. Fibrous encapsulation of a Kudoa sp. plasmodium in viviparous blenny (Zoarces viviparus).
TitleHistological features of muscle-invading species
CaptionHistological features of muscle-invading species. A. Sporogonic plasmodia of Myxobolus artus in the skeletal muscle of carp (C. carpio) (inset: mature spore in valvular view). B. Granulomatous host response to a plasmodium of M. artus within the renal tissue (same fish as in Fig. 8.10A). C. A plasmodium of Myxobolus pseudodispar within the muscle of a juvenile roach (Rutilus rutilus). D. Focal inflammatory response to M. pseudodispar spores and sporogonic stages from a ruptured plasmodium. E. Intra-myofibrillar Kudoa infection in cow-nosed ray (Rhinoptera bonasus). F. Multiple plasmodia of Kudoa sp. in the myofibrils of common goby (Pomatoschistus microps). G. Giemsa-stained section of Kudoa thyrsites infection in cod (Gadus morhua). H. Fibrous encapsulation of a Kudoa sp. plasmodium in viviparous blenny (Zoarces viviparus).
CopyrightStephen W. Feist & Matt Longshaw
Histological features of muscle-invading species. A. Sporogonic plasmodia of Myxobolus artus in the skeletal muscle of carp (C. carpio) (inset: mature spore in valvular view). B. Granulomatous host response to a plasmodium of M. artus within the renal tissue (same fish as in Fig. 8.10A). C. A plasmodium of Myxobolus pseudodispar within the muscle of a juvenile roach (Rutilus rutilus). D. Focal inflammatory response to M. pseudodispar spores and sporogonic stages from a ruptured plasmodium. E. Intra-myofibrillar Kudoa infection in cow-nosed ray (Rhinoptera bonasus). F. Multiple plasmodia of Kudoa sp. in the myofibrils of common goby (Pomatoschistus microps). G. Giemsa-stained section of Kudoa thyrsites infection in cod (Gadus morhua). H. Fibrous encapsulation of a Kudoa sp. plasmodium in viviparous blenny (Zoarces viviparus).
Histological features of muscle-invading speciesHistological features of muscle-invading species. A. Sporogonic plasmodia of Myxobolus artus in the skeletal muscle of carp (C. carpio) (inset: mature spore in valvular view). B. Granulomatous host response to a plasmodium of M. artus within the renal tissue (same fish as in Fig. 8.10A). C. A plasmodium of Myxobolus pseudodispar within the muscle of a juvenile roach (Rutilus rutilus). D. Focal inflammatory response to M. pseudodispar spores and sporogonic stages from a ruptured plasmodium. E. Intra-myofibrillar Kudoa infection in cow-nosed ray (Rhinoptera bonasus). F. Multiple plasmodia of Kudoa sp. in the myofibrils of common goby (Pomatoschistus microps). G. Giemsa-stained section of Kudoa thyrsites infection in cod (Gadus morhua). H. Fibrous encapsulation of a Kudoa sp. plasmodium in viviparous blenny (Zoarces viviparus).Stephen W. Feist & Matt Longshaw
Pathological features of cartilage infections. A. Low-power view of head cartilage of rainbow trout (Oncorhynchus mykiss) infected with Myxobolus cerebralis, showing disintegration of cartilaginous elements. B. High-power view showing spores and developmental stages of M. cerebralis amongst destroyed cartilage (A and B are from Giemsa-stained sections). C and D. Vertebral deformation and complete fusion of vertebrae in chub (L. cephalus) caused by Myxobolus buckei (C. inset: mature spore of M. buckei in valvular view).
TitlePathological features of cartilage infections
CaptionPathological features of cartilage infections. A. Low-power view of head cartilage of rainbow trout (Oncorhynchus mykiss) infected with Myxobolus cerebralis, showing disintegration of cartilaginous elements. B. High-power view showing spores and developmental stages of M. cerebralis amongst destroyed cartilage (A and B are from Giemsa-stained sections). C and D. Vertebral deformation and complete fusion of vertebrae in chub (L. cephalus) caused by Myxobolus buckei (C. inset: mature spore of M. buckei in valvular view).
CopyrightStephen W. Feist & Matt Longshaw
Pathological features of cartilage infections. A. Low-power view of head cartilage of rainbow trout (Oncorhynchus mykiss) infected with Myxobolus cerebralis, showing disintegration of cartilaginous elements. B. High-power view showing spores and developmental stages of M. cerebralis amongst destroyed cartilage (A and B are from Giemsa-stained sections). C and D. Vertebral deformation and complete fusion of vertebrae in chub (L. cephalus) caused by Myxobolus buckei (C. inset: mature spore of M. buckei in valvular view).
Pathological features of cartilage infectionsPathological features of cartilage infections. A. Low-power view of head cartilage of rainbow trout (Oncorhynchus mykiss) infected with Myxobolus cerebralis, showing disintegration of cartilaginous elements. B. High-power view showing spores and developmental stages of M. cerebralis amongst destroyed cartilage (A and B are from Giemsa-stained sections). C and D. Vertebral deformation and complete fusion of vertebrae in chub (L. cephalus) caused by Myxobolus buckei (C. inset: mature spore of M. buckei in valvular view).Stephen W. Feist & Matt Longshaw
Pathological features of cartilage infections. E. Cysts of Myxobolus aeglifini in the scleral cartilage of the eye of poor cod (Trisopterus minutus) (inset: mature spore of M. aeglifini in valvular view). F. Section through two large cysts of M. aeglifini in the scleral cartilage of whiting (Merlangius merlangus). G. Myxobolus sp. in the cranial cartilage of common goby (P. microps). H. Cysts of Myxobolus sp. in the cranial cartilage of roach (R. rutilus).
TitlePathological features of cartilage infections
CaptionPathological features of cartilage infections. E. Cysts of Myxobolus aeglifini in the scleral cartilage of the eye of poor cod (Trisopterus minutus) (inset: mature spore of M. aeglifini in valvular view). F. Section through two large cysts of M. aeglifini in the scleral cartilage of whiting (Merlangius merlangus). G. Myxobolus sp. in the cranial cartilage of common goby (P. microps). H. Cysts of Myxobolus sp. in the cranial cartilage of roach (R. rutilus).
CopyrightStephen W. Feist & Matt Longshaw
Pathological features of cartilage infections. E. Cysts of Myxobolus aeglifini in the scleral cartilage of the eye of poor cod (Trisopterus minutus) (inset: mature spore of M. aeglifini in valvular view). F. Section through two large cysts of M. aeglifini in the scleral cartilage of whiting (Merlangius merlangus). G. Myxobolus sp. in the cranial cartilage of common goby (P. microps). H. Cysts of Myxobolus sp. in the cranial cartilage of roach (R. rutilus).
Pathological features of cartilage infectionsPathological features of cartilage infections. E. Cysts of Myxobolus aeglifini in the scleral cartilage of the eye of poor cod (Trisopterus minutus) (inset: mature spore of M. aeglifini in valvular view). F. Section through two large cysts of M. aeglifini in the scleral cartilage of whiting (Merlangius merlangus). G. Myxobolus sp. in the cranial cartilage of common goby (P. microps). H. Cysts of Myxobolus sp. in the cranial cartilage of roach (R. rutilus).Stephen W. Feist & Matt Longshaw
Proliferative kidney disease, pathology and morphology of the causative agent Tetracapsuloides bryosalmonae. A. Rainbow trout (Oncorhynchus mykiss) fingerling exhibiting renal hypertrophy. B. Giemsa stained renal impression smear showing extrasporogonic stage of T. bryosalmonae surrounded by host phagocytes. C. Diagrammatic representations of stages in the development of histozoic T. bryosalmonae from rainbow trout kidney (primary cell nucleus, N; secondary cell, S; tertiary cell, T). D. Electron micrograph of a renal interstitial stage from rainbow trout kidney. Primary cell contains a prominent nucleus (N) with two secondary cells (S), one of which contains two tertiary cells (T). E. Fresh preparation of T. bryosalmonae from trout kidney with characteristic cytoplasmic granules and secondary cells within the primary cell, which is itself surrounded by phagocytic cells. F. Low-power view of a section of infected renal tissue showing loss of excretory elements and proliferation of inte
TitleProliferative kidney disease, pathology and morphology
CaptionProliferative kidney disease, pathology and morphology of the causative agent Tetracapsuloides bryosalmonae. A. Rainbow trout (Oncorhynchus mykiss) fingerling exhibiting renal hypertrophy. B. Giemsa stained renal impression smear showing extrasporogonic stage of T. bryosalmonae surrounded by host phagocytes. C. Diagrammatic representations of stages in the development of histozoic T. bryosalmonae from rainbow trout kidney (primary cell nucleus, N; secondary cell, S; tertiary cell, T). D. Electron micrograph of a renal interstitial stage from rainbow trout kidney. Primary cell contains a prominent nucleus (N) with two secondary cells (S), one of which contains two tertiary cells (T). E. Fresh preparation of T. bryosalmonae from trout kidney with characteristic cytoplasmic granules and secondary cells within the primary cell, which is itself surrounded by phagocytic cells. F. Low-power view of a section of infected renal tissue showing loss of excretory elements and proliferation of inte
CopyrightStephen W. Feist & Matt Longshaw
Proliferative kidney disease, pathology and morphology of the causative agent Tetracapsuloides bryosalmonae. A. Rainbow trout (Oncorhynchus mykiss) fingerling exhibiting renal hypertrophy. B. Giemsa stained renal impression smear showing extrasporogonic stage of T. bryosalmonae surrounded by host phagocytes. C. Diagrammatic representations of stages in the development of histozoic T. bryosalmonae from rainbow trout kidney (primary cell nucleus, N; secondary cell, S; tertiary cell, T). D. Electron micrograph of a renal interstitial stage from rainbow trout kidney. Primary cell contains a prominent nucleus (N) with two secondary cells (S), one of which contains two tertiary cells (T). E. Fresh preparation of T. bryosalmonae from trout kidney with characteristic cytoplasmic granules and secondary cells within the primary cell, which is itself surrounded by phagocytic cells. F. Low-power view of a section of infected renal tissue showing loss of excretory elements and proliferation of inte
Proliferative kidney disease, pathology and morphologyProliferative kidney disease, pathology and morphology of the causative agent Tetracapsuloides bryosalmonae. A. Rainbow trout (Oncorhynchus mykiss) fingerling exhibiting renal hypertrophy. B. Giemsa stained renal impression smear showing extrasporogonic stage of T. bryosalmonae surrounded by host phagocytes. C. Diagrammatic representations of stages in the development of histozoic T. bryosalmonae from rainbow trout kidney (primary cell nucleus, N; secondary cell, S; tertiary cell, T). D. Electron micrograph of a renal interstitial stage from rainbow trout kidney. Primary cell contains a prominent nucleus (N) with two secondary cells (S), one of which contains two tertiary cells (T). E. Fresh preparation of T. bryosalmonae from trout kidney with characteristic cytoplasmic granules and secondary cells within the primary cell, which is itself surrounded by phagocytic cells. F. Low-power view of a section of infected renal tissue showing loss of excretory elements and proliferation of inteStephen W. Feist & Matt Longshaw
Pathology of renal infections. A. Intracellular and coelozoic stages of Hoferellus carassii in the renal tubule of goldfish (Carassius auratus). B. Three-spined stickleback (G. aculeatus) kidney showing distension of renal tubules caused by numerous sporogonic stages of Myxobilatus gasterostei. C. Large xenoma of extrasporogonic stages of Myxidium lieberkuehni, which has replaced the glomerular tissue in the kidney of pike (E. lucius). D. Glomerular and tubule infections of Myxobilatus platessae in the kidney of European flounder (Platichthys flesus). E. Sporogonic stages of Myxidium minteri in the renal tubule lumen of chinook salmon (Oncorhynchus tshawytscha). F. Parvicapsula sp. sporogonic stages within the renal.
TitlePathology of renal infections
CaptionPathology of renal infections. A. Intracellular and coelozoic stages of Hoferellus carassii in the renal tubule of goldfish (Carassius auratus). B. Three-spined stickleback (G. aculeatus) kidney showing distension of renal tubules caused by numerous sporogonic stages of Myxobilatus gasterostei. C. Large xenoma of extrasporogonic stages of Myxidium lieberkuehni, which has replaced the glomerular tissue in the kidney of pike (E. lucius). D. Glomerular and tubule infections of Myxobilatus platessae in the kidney of European flounder (Platichthys flesus). E. Sporogonic stages of Myxidium minteri in the renal tubule lumen of chinook salmon (Oncorhynchus tshawytscha). F. Parvicapsula sp. sporogonic stages within the renal.
CopyrightStephen W. Feist & Matt Longshaw
Pathology of renal infections. A. Intracellular and coelozoic stages of Hoferellus carassii in the renal tubule of goldfish (Carassius auratus). B. Three-spined stickleback (G. aculeatus) kidney showing distension of renal tubules caused by numerous sporogonic stages of Myxobilatus gasterostei. C. Large xenoma of extrasporogonic stages of Myxidium lieberkuehni, which has replaced the glomerular tissue in the kidney of pike (E. lucius). D. Glomerular and tubule infections of Myxobilatus platessae in the kidney of European flounder (Platichthys flesus). E. Sporogonic stages of Myxidium minteri in the renal tubule lumen of chinook salmon (Oncorhynchus tshawytscha). F. Parvicapsula sp. sporogonic stages within the renal.
Pathology of renal infectionsPathology of renal infections. A. Intracellular and coelozoic stages of Hoferellus carassii in the renal tubule of goldfish (Carassius auratus). B. Three-spined stickleback (G. aculeatus) kidney showing distension of renal tubules caused by numerous sporogonic stages of Myxobilatus gasterostei. C. Large xenoma of extrasporogonic stages of Myxidium lieberkuehni, which has replaced the glomerular tissue in the kidney of pike (E. lucius). D. Glomerular and tubule infections of Myxobilatus platessae in the kidney of European flounder (Platichthys flesus). E. Sporogonic stages of Myxidium minteri in the renal tubule lumen of chinook salmon (Oncorhynchus tshawytscha). F. Parvicapsula sp. sporogonic stages within the renal.Stephen W. Feist & Matt Longshaw
Pathology of renal infections. A. Sphaerospora sp. in the kidney of dace (Leuciscus leuciscus) causing dilatation of renal tubule with reduction in tubule epithelial height. B. Atrophy of the glomerular tuft surrounded by a coelozoic plasmodium of Myxidium rhodei in the kidney of dace (Leuciscus leuciscus). C. Fibrous encapsulation of sporogonic plasmodia of M. rhodei in the renal interstitial tissue of R. rutilus. D. Intracellular extrasporogonic stage of an unidentified myxosporean in the renal tubule epithelium of dace (L. leuciscus).
TitlePathology of renal infections
CaptionPathology of renal infections. A. Sphaerospora sp. in the kidney of dace (Leuciscus leuciscus) causing dilatation of renal tubule with reduction in tubule epithelial height. B. Atrophy of the glomerular tuft surrounded by a coelozoic plasmodium of Myxidium rhodei in the kidney of dace (Leuciscus leuciscus). C. Fibrous encapsulation of sporogonic plasmodia of M. rhodei in the renal interstitial tissue of R. rutilus. D. Intracellular extrasporogonic stage of an unidentified myxosporean in the renal tubule epithelium of dace (L. leuciscus).
CopyrightStephen W. Feist & Matt Longshaw
Pathology of renal infections. A. Sphaerospora sp. in the kidney of dace (Leuciscus leuciscus) causing dilatation of renal tubule with reduction in tubule epithelial height. B. Atrophy of the glomerular tuft surrounded by a coelozoic plasmodium of Myxidium rhodei in the kidney of dace (Leuciscus leuciscus). C. Fibrous encapsulation of sporogonic plasmodia of M. rhodei in the renal interstitial tissue of R. rutilus. D. Intracellular extrasporogonic stage of an unidentified myxosporean in the renal tubule epithelium of dace (L. leuciscus).
Pathology of renal infectionsPathology of renal infections. A. Sphaerospora sp. in the kidney of dace (Leuciscus leuciscus) causing dilatation of renal tubule with reduction in tubule epithelial height. B. Atrophy of the glomerular tuft surrounded by a coelozoic plasmodium of Myxidium rhodei in the kidney of dace (Leuciscus leuciscus). C. Fibrous encapsulation of sporogonic plasmodia of M. rhodei in the renal interstitial tissue of R. rutilus. D. Intracellular extrasporogonic stage of an unidentified myxosporean in the renal tubule epithelium of dace (L. leuciscus).Stephen W. Feist & Matt Longshaw
Infections of the gall bladder, liver and neural tissues. A. Low-power view of papillomatous ingrowths of the gall-bladder epithelium of saithe (Pollachius virens) infected with Myxidium gadi. B. Semi-thin resin section from the previous specimen showing attachment of sporogonic M. gadi plasmodia. C. Hepatobiliary fibrosis associated with invasion of the bile ductules with elongate plasmodia of Myxidium truttae infecting brown trout (Salmo trutta). D. High-power view showing attenuation of gall-bladder epithelium and the presence of a small plasmodium of M. truttae within the hepatic parenchyma. E. Spores of Myxobolus cotti within the brain of bullhead (Cottus gobio).
TitleInfections of the gall bladder, liver and neural tissues
CaptionInfections of the gall bladder, liver and neural tissues. A. Low-power view of papillomatous ingrowths of the gall-bladder epithelium of saithe (Pollachius virens) infected with Myxidium gadi. B. Semi-thin resin section from the previous specimen showing attachment of sporogonic M. gadi plasmodia. C. Hepatobiliary fibrosis associated with invasion of the bile ductules with elongate plasmodia of Myxidium truttae infecting brown trout (Salmo trutta). D. High-power view showing attenuation of gall-bladder epithelium and the presence of a small plasmodium of M. truttae within the hepatic parenchyma. E. Spores of Myxobolus cotti within the brain of bullhead (Cottus gobio).
CopyrightStephen W. Feist & Matt Longshaw
Infections of the gall bladder, liver and neural tissues. A. Low-power view of papillomatous ingrowths of the gall-bladder epithelium of saithe (Pollachius virens) infected with Myxidium gadi. B. Semi-thin resin section from the previous specimen showing attachment of sporogonic M. gadi plasmodia. C. Hepatobiliary fibrosis associated with invasion of the bile ductules with elongate plasmodia of Myxidium truttae infecting brown trout (Salmo trutta). D. High-power view showing attenuation of gall-bladder epithelium and the presence of a small plasmodium of M. truttae within the hepatic parenchyma. E. Spores of Myxobolus cotti within the brain of bullhead (Cottus gobio).
Infections of the gall bladder, liver and neural tissuesInfections of the gall bladder, liver and neural tissues. A. Low-power view of papillomatous ingrowths of the gall-bladder epithelium of saithe (Pollachius virens) infected with Myxidium gadi. B. Semi-thin resin section from the previous specimen showing attachment of sporogonic M. gadi plasmodia. C. Hepatobiliary fibrosis associated with invasion of the bile ductules with elongate plasmodia of Myxidium truttae infecting brown trout (Salmo trutta). D. High-power view showing attenuation of gall-bladder epithelium and the presence of a small plasmodium of M. truttae within the hepatic parenchyma. E. Spores of Myxobolus cotti within the brain of bullhead (Cottus gobio).Stephen W. Feist & Matt Longshaw
Infections of the intestine and associated tissues. A. Destruction of the intestinal mucosa of gilthead seabream (Sparus aurata) caused by Enteromyxum leei. B. Intraepithelial stages of E. leei in the intestinal mucosa of S. aurata. C. Section of pyloric caecae with large numbers of plasmodia and spores of Ceratomyxa shasta (arrows) in the underlying connective tissue. Inset: developing spores of C. Shasta. D. Intraepithelial stages of C. shasta in the mucosal epithelium and submucosal tissues (arrow). E and F. Cysts of Myxobolus sp. in the thin connective-tissue layer underlying the intestinal epithelium of minnow (Phoxinus phoxinus) and roach (R. rutilus), respectively.
TitleInfections of the intestine and associated tissues
CaptionInfections of the intestine and associated tissues. A. Destruction of the intestinal mucosa of gilthead seabream (Sparus aurata) caused by Enteromyxum leei. B. Intraepithelial stages of E. leei in the intestinal mucosa of S. aurata. C. Section of pyloric caecae with large numbers of plasmodia and spores of Ceratomyxa shasta (arrows) in the underlying connective tissue. Inset: developing spores of C. Shasta. D. Intraepithelial stages of C. shasta in the mucosal epithelium and submucosal tissues (arrow). E and F. Cysts of Myxobolus sp. in the thin connective-tissue layer underlying the intestinal epithelium of minnow (Phoxinus phoxinus) and roach (R. rutilus), respectively.
CopyrightStephen W. Feist & Matt Longshaw
Infections of the intestine and associated tissues. A. Destruction of the intestinal mucosa of gilthead seabream (Sparus aurata) caused by Enteromyxum leei. B. Intraepithelial stages of E. leei in the intestinal mucosa of S. aurata. C. Section of pyloric caecae with large numbers of plasmodia and spores of Ceratomyxa shasta (arrows) in the underlying connective tissue. Inset: developing spores of C. Shasta. D. Intraepithelial stages of C. shasta in the mucosal epithelium and submucosal tissues (arrow). E and F. Cysts of Myxobolus sp. in the thin connective-tissue layer underlying the intestinal epithelium of minnow (Phoxinus phoxinus) and roach (R. rutilus), respectively.
Infections of the intestine and associated tissuesInfections of the intestine and associated tissues. A. Destruction of the intestinal mucosa of gilthead seabream (Sparus aurata) caused by Enteromyxum leei. B. Intraepithelial stages of E. leei in the intestinal mucosa of S. aurata. C. Section of pyloric caecae with large numbers of plasmodia and spores of Ceratomyxa shasta (arrows) in the underlying connective tissue. Inset: developing spores of C. Shasta. D. Intraepithelial stages of C. shasta in the mucosal epithelium and submucosal tissues (arrow). E and F. Cysts of Myxobolus sp. in the thin connective-tissue layer underlying the intestinal epithelium of minnow (Phoxinus phoxinus) and roach (R. rutilus), respectively.Stephen W. Feist & Matt Longshaw
(Left panel) SSU rRNA gene primer map for C. shasta. Predicted amplicon sizes for different combinations of the primers (Cs1, Cs2, R, Cs3, Cs4, Cs5, 18SUNIF and 18SUNIR are shown in base pairs. Right panel shows specificity of different primer pairs for C. shasta DNA. Amplicons of the predicted size were obtained with DNA from C. shasta spores (lanes a, d, g, j and m), but not rainbow trout (lanes b, e, h, k and n) or Henneguya salminicola (c, f, i, l and o). Primer pairs were Cs1-Cs5 (lanes a-c); Cs1-Cs3 (lanes d-f); Cs2-Cs5 (lanes g-i); R-Cs4 (lanes j-l); and r-Cs5 (lanes m-o). (From Palenzuela et al., 1999.) with permission from Diseases of Aquatic Organisms.)
TitlePrimers
Caption(Left panel) SSU rRNA gene primer map for C. shasta. Predicted amplicon sizes for different combinations of the primers (Cs1, Cs2, R, Cs3, Cs4, Cs5, 18SUNIF and 18SUNIR are shown in base pairs. Right panel shows specificity of different primer pairs for C. shasta DNA. Amplicons of the predicted size were obtained with DNA from C. shasta spores (lanes a, d, g, j and m), but not rainbow trout (lanes b, e, h, k and n) or Henneguya salminicola (c, f, i, l and o). Primer pairs were Cs1-Cs5 (lanes a-c); Cs1-Cs3 (lanes d-f); Cs2-Cs5 (lanes g-i); R-Cs4 (lanes j-l); and r-Cs5 (lanes m-o). (From Palenzuela et al., 1999.) with permission from Diseases of Aquatic Organisms.)
CopyrightTed G. Clark
(Left panel) SSU rRNA gene primer map for C. shasta. Predicted amplicon sizes for different combinations of the primers (Cs1, Cs2, R, Cs3, Cs4, Cs5, 18SUNIF and 18SUNIR are shown in base pairs. Right panel shows specificity of different primer pairs for C. shasta DNA. Amplicons of the predicted size were obtained with DNA from C. shasta spores (lanes a, d, g, j and m), but not rainbow trout (lanes b, e, h, k and n) or Henneguya salminicola (c, f, i, l and o). Primer pairs were Cs1-Cs5 (lanes a-c); Cs1-Cs3 (lanes d-f); Cs2-Cs5 (lanes g-i); R-Cs4 (lanes j-l); and r-Cs5 (lanes m-o). (From Palenzuela et al., 1999.) with permission from Diseases of Aquatic Organisms.)
Primers(Left panel) SSU rRNA gene primer map for C. shasta. Predicted amplicon sizes for different combinations of the primers (Cs1, Cs2, R, Cs3, Cs4, Cs5, 18SUNIF and 18SUNIR are shown in base pairs. Right panel shows specificity of different primer pairs for C. shasta DNA. Amplicons of the predicted size were obtained with DNA from C. shasta spores (lanes a, d, g, j and m), but not rainbow trout (lanes b, e, h, k and n) or Henneguya salminicola (c, f, i, l and o). Primer pairs were Cs1-Cs5 (lanes a-c); Cs1-Cs3 (lanes d-f); Cs2-Cs5 (lanes g-i); R-Cs4 (lanes j-l); and r-Cs5 (lanes m-o). (From Palenzuela et al., 1999.) with permission from Diseases of Aquatic Organisms.)Ted G. Clark
Localization of M. cerebralis in host tissues by in situ hybridization. Panels (A) and (B) show sections of cranial cartilage from rainbow trout infected with M. cerebralis and processed for in situ hybridization 3 months post-exposure. Oligonucleotide probes were labelled at their 3' ends with dUTP-digoxigenin. Preincubation of sections with excess unlabelled probe eliminated tissue staining (panel A). Strong staining of developmental stages of the parasite was visible in host tissues using the labelled probe by itself (arrows, panel B). (From Antonio et al., 1998.) with permission from Journal of Aquatic Animal Health.)
TitleLocalization of M. cerebralis
CaptionLocalization of M. cerebralis in host tissues by in situ hybridization. Panels (A) and (B) show sections of cranial cartilage from rainbow trout infected with M. cerebralis and processed for in situ hybridization 3 months post-exposure. Oligonucleotide probes were labelled at their 3' ends with dUTP-digoxigenin. Preincubation of sections with excess unlabelled probe eliminated tissue staining (panel A). Strong staining of developmental stages of the parasite was visible in host tissues using the labelled probe by itself (arrows, panel B). (From Antonio et al., 1998.) with permission from Journal of Aquatic Animal Health.)
CopyrightTed G. Clark
Localization of M. cerebralis in host tissues by in situ hybridization. Panels (A) and (B) show sections of cranial cartilage from rainbow trout infected with M. cerebralis and processed for in situ hybridization 3 months post-exposure. Oligonucleotide probes were labelled at their 3' ends with dUTP-digoxigenin. Preincubation of sections with excess unlabelled probe eliminated tissue staining (panel A). Strong staining of developmental stages of the parasite was visible in host tissues using the labelled probe by itself (arrows, panel B). (From Antonio et al., 1998.) with permission from Journal of Aquatic Animal Health.)
Localization of M. cerebralisLocalization of M. cerebralis in host tissues by in situ hybridization. Panels (A) and (B) show sections of cranial cartilage from rainbow trout infected with M. cerebralis and processed for in situ hybridization 3 months post-exposure. Oligonucleotide probes were labelled at their 3' ends with dUTP-digoxigenin. Preincubation of sections with excess unlabelled probe eliminated tissue staining (panel A). Strong staining of developmental stages of the parasite was visible in host tissues using the labelled probe by itself (arrows, panel B). (From Antonio et al., 1998.) with permission from Journal of Aquatic Animal Health.)Ted G. Clark

Identity

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Preferred Scientific Name

  • Myxozoan infections of fish

Overview

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Myxozoans are highly specialized metazoan parasites of aquatic hosts with a very wide host range. This diverse group of organisms is characterized by multicellular spores with polar capsules containing extrudable polar filaments. Interest in the group has intensified along with the development of aquaculture since many species cause serious disease outbreaks in farmed fish species, in both freshwater and marine environments. Myxobolus cerebralis (Hofer, 1903), Tetracapsuloides bryosalmonae (Canning, Curry, Feist, Longshaw and Okamura, 1999) and Ceratomyxa shasta Noble, 1950, in salmonids are examples. The economic impact of such parasites can be severe, especially where prevalence rates are high. They can also have a severe impact on wild fish stocks. Infections with multivalvulid myxozoans such as Kudoa spp. within the musculature of several marine fish species severely reduce the flesh quality and in some cases cause extensive myoliquefaction, rendering the product unmarketable. In freshwater environments, M. cerebralis has been shown to be a significant factor in population declines of wild stocks of Pacific salmonids.

Recognition of the requirement for oligochaete and bryozoan obligatory hosts in the life cycle of several freshwater species has resulted in an increased interest in the biology of the group. Many new species have been described and significant advances have been made in the understanding of the transmission biology of several species. The development of specific diagnostic methods has facilitated studies on the pathogenesis and the use of molecular phylogenetic techniques has provided fundamental advances in the taxonomy of the Myxozoa. In particular, insights into the evolution of Myxozoa have been provided by morphological and molecular studies of the class Malacosporea. Current research continues to be focused on the occurrence of disease outbreaks, pathogenesis and phylogeny of these parasites. However, for several important and wellcharacterized diseases, including ceratomyxosis, whirling disease and proliferative kidney disease (PKD), the emphasis is on host immunity, prevention and control. This section provides a summary of current knowledge of the biology of the phylum Myxozoa, with particular attention given to those parasites as agents of disease in fish.

During the past decade there have been several significant advances in the knowledge of myxozoan parasites and the diseases they cause. Many new species have been identified and the increasing use of rDNA sequencing has and will continue to play a pivotal role in the confident assignment of these, particularly since many 'species' exhibit spore pleomorphism, both within individual plasmodia and in those occurring in multiple sites within hosts. Although most species cause little harm, a few have become recognized as serious pathogens, especially in aquaculture situations. Detailed studies on host-parasite interactions have provided valuable information on the pathogenesis of these, and the cellular aspects of the host response to myxozoan infections is reasonably well understood. However, this is true only for a small percentage of known species. For most, the mechanisms of migration and replication within the host remain a mystery. Host immunology during myxozoan infections remains little studied, apart from recent investigations into diseases such as PKD and ceratomyxosis in salmonids. Increasing knowledge in this area will offer insights into the molecular aspects of host-parasite responses, which may offer potential for the development of DNA-based vaccines. However, it currently seems unlikely that these will be developed in the near future. The application of genomic, proteomic and metabolomic methodology offers powerful tools to characterize the changes occurring within the hosts during the disease process and may reveal molecular targets for diagnosis and treatment. The elucidation of the life cycle for several myxosporeans and the malacosporean T. bryosalmonae offers the most immediate opportunity for amelioration of the diseases caused by these organisms. With this knowledge, management strategies aimed at avoidance of the infectious stage or deliberate exposure aimed at eliciting an effective host response without the manifestation of clinical disease will continue to be refined according to local situations until proprietary treatments become available.

Significant advances regarding the phylogeny of the Myxozoa have been made in recent years, in particular, associated with the suppression of the class Actinosporea and the erection of the class Malacosporea following the recognition of bryozoans as hosts. The recognition of the enigmatic parasite Buddenbrockia plumatellae as a myxozoan and subsequent morphological and molecular studies have provided compelling evidence that myxozoans are highly specialized triploblast animals, possibly related to the Nematoda. There are likely to be several interesting developments in this area as new invertebrate hosts for similar parasites become recognized. Fundamental understanding of the evolutionary aspects of myxozoan biology will also be refined. Interest in the life cycles of marine species and their anticipated invertebrate hosts will surely provide many fascinating discoveries. Practical applications for infections in mariculture species can be anticipated. In addition, the discovery of myxozoan parasites in mammals and birds indicates the possibility that many other hosts are yet to be identified.

Most investigations consider myxozoan pathogenicity at the individual level, with comment on the extent of mortalities in captive populations. However, with the notable exception of whirling disease, there have been few investigations on the effect of myxozoan parasites on wild populations. That the pathogenicity of myxozoan parasites and other pathogens is dependent on a wide range of factors, both internal and external to the host, is accepted. Usually, descriptions of the 'diseases' caused indicate the more severe manifestations and may comment on individual survivability in the short term. However, most myxozoans, even if they cause acute disease, do not kill their host unless secondary stressors and/or reduced immunocompetence has a significant effect. Consequently, myxozoan parasitism tends to be chronic, with the host able to survive for several years, depending on the species. There are very few data on the effects of such chronic parasitism in wild fish populations and only in the case of whirling disease have population declines been associated with the infection. An important area of research will be to understand the long-term impacts of these infections at the population level. Knowledge of the population biology of target species and of invertebrate hosts will be essential in this regard.

With the continuing increase in aquaculture and the introduction of new species, it is likely that myxozoan parasites will also continue to pose health challenges to the industry, both from previously undescribed species and from others able to exploit a new niche in high-density host populations. Parasites, such as Enteromyxum leei, with direct transmission capabilities will pose the most serious threats. Epidemiological approaches and the application of risk analysis for assessment of potential disease spread have begun to be used for fish diseases, particularly those listed by the Office International des Epizooties (OIE). Similar approaches applied to the study of myxozoan infections in wild populations will be useful for understanding temporal and spatial changes in infection rates, especially where environmental data are also considered.

[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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Myxozoans are common parasites of teleost fish and invertebrates, with a few species infecting elasmobranchs (Arthur and Lom, 1985; Stoffregen and Anderson, 1990; Heupel and Bennett, 1996), amphibians and reptiles and a single case of infection in a digenean (Overstreet, 1976), and a Kudoa sp. has been reported causing myoliquefaction in octopus (Yokoyama and Masuda, 2001). However, recent studies have revealed a much wider host range amongst animal phyla than had previously been imagined. A myxozoan-like parasite in European mole, Talpa europaea (Friedrich et al., 2000) and the presence of myxozoans in inflammatory lesions of the bile ducts and hepatic parenchyma of anatid ducks held in an enclosure at a zoological garden have been reported (Lowenstine et al., 2002). In addition, there have been several reports of myxozoan spores in human stool samples. In each case, the patients were suffering from abdominal pain or diarrhoea that were related to other enteric pathogens. These are likely to have arisen through the ingestion of infected fish material (McClelland et al., 1997; Boreham et al., 1998; Lebbad and Wilcox, 1998; Moncada et al., 2001).

There is no clear pattern regarding host specificity within the Myxozoa. Some species appear to be highly specific to their host, whilst others may be specific at the family level (e.g. Myxobolus cerebralis and Tetracapsuloides bryosalmonae in salmonid hosts). Others possess a wide host specificity (e.g. Myxobolus aeglefiniAuerbach, 1906, in both gadoid and pleuronectid hosts) (Nielsen et al., 2002).

Distribution

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Myxozoans appear to be distributed throughout marine, estuarine and freshwater environments worldwide. Phylogenetic studies by Kent et al. (2001) have shown that, at the genus level, marine taxa branch separately from genera that usually infect freshwater fish.

There are discontinuities in the prevalence of myxozoans in fish hosts with differences in prevalence being found across the range of the hosts. However, some myxozoans have become established or at least have the potential to become established in other geographical regions via the anthropogenic translocation of the hosts. Myxobolus cerebralis is thought to have been transferred to the USA, New Zealand, South Africa and Australia through the movement of either live fish or dead, chilled fish destined for human consumption (Bartholomew and Reno, 2002, Whipps et al., 2004b). Lowers and Bartholomew (2003) demonstrated that the importation of oligochaetes into the USA carried a risk of translocating Myxozoa from Europe to the USA since actinospore stages (see later section) were found within oligochaetes destined for the pet trade. Myxobolus artus Akhmerov, 1960 in the musculature of carp was originally restricted to the former USSR, but through the international trade in the host species it has now been recorded in Japan, Taiwan and Indonesia (Ogawa et al., 1992) and in the UK (M. Longshaw and S.W. Feist, unpublished data). Similarly, Myxobolus koi Kudo, 1919 has been reported outside its normal geographical range in the UK (Crawshaw and Sweeting, 1986) and a parasite morphologically identical to the European Myxobolus pseudodispar Gorbunova, 1936 has been reported in peamouths in the USA (Kent et al., 1996). The myxozoan Sphaeromyxa sevastopoli Naidenova, 1970, a parasite originally recorded in gobies from the Black Sea area of the former USSR, has been reported in two recently established European goby species in Michigan, USA (Pronin et al., 1997). Sphaerospora molnari Lom, Dyková, Pavlásková and Grupcheva, 1983 and Sphaerospora cf. chinensis (Lee and Nie, 1965) have been reported in Germany and the USA, respectively, outside their normal range (Hedrick et al., 1990; Kaup et al., 1995). However, the paucity of data on the diversity of myxozoans in many hosts and the geographical range for most species makes it difficult to come to firm conclusions regarding possible translocation of these parasites.

The discontinuity in parasite prevalence and distribution has been used to suggest their use as biological tags for a number of hosts. M. aeglefini has been used as a biological tag in studies of stock structure of Pleuronectes platessa and to examine migration routes of Gadus morhua hiemalis (van Banning et al., 1978; Timofeeva and Marasaeva, 1984). Hemmingsen et al. (1991) suggested that Myxidium spp. could be useful tags to discriminate stocks of Gadus morhua in northern Norway and, in particular Larsen et al. (1997) showed that Myxidium oviforme Parisi, 1912 and Zschokkella hildae Auerbach, 1910 were suitable tags in this case. Kabata (1963) discriminated stocks of Trisopterus luscus using Ceratomyxa arcuata Thélohan, 1892 and Myxidium sphaericum Thélohan, 1895 and stocks of Melanogrammus aeglefinus using Myxidium sp., Leptotheca sp. and Sphaeromyxa hellandi Auerbach, 1909.

Two species of Myxobolus in the central nervous system of salmonids, Myxobolus neurobius Schuberg and Schröder, 1905 and Myxobolus arcticus Pugachev and Khokhlov, 1979, have been used as biological tags to study the migration and stock structure of salmonids. Awakura et al. (1995) used both M. arcticus and M. neurobius as a biological tag to demonstrate that Oncorhynchus masou caught in Japanese coastal waters originated from a number of different river systems and that there was migration of the fish from the Sea of Okhotsk to the Sea of Japan. Furthermore, Urawa and Nagasawa (1995) and Urawa et al. (1998) have shown marked differences in the prevalence of M. arcticus both in different geographical regions and in different Oncorhynchus spp., which means that M. arcticus shows promise as a biological tag. Margolis (1993) reported a court case in Canada in which the prevalence of M. arcticus in sockeye salmon was used to demonstrate that the fish were caught outside the commercial fishing season.

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

Arctic SeaPresentTimofeeva and Marasaeva, 1984
Atlantic, NortheastPresentBanning et al., 1978; Feist and Bucke, 1992
Atlantic, NorthwestPresentMorrison et al., 1996
Indian Ocean, WesternPresentDiamant and Paperna, 1992
Mediterranean and Black SeaPresentSitjà-Bobadilla and Alvarez-Pellitero, 1993
Pacific, NortheastPresentArthur and Lom, 1985; Urawa and Nagasawa, 1995; Urawa et al., 1998
Pacific, NorthwestPresentYasutake and Wood, 1957; Urawa and Nagasawa, 1995; Urawa et al., 1998; Yokoyama and Wakabayashi, 2000
Pacific, SouthwestPresentHeupel and Bennett, 1996; Whipps et al., 2003

Asia

ChinaPresentPresent based on regional distribution.
-ZhejiangPresentWu et al., 1975
IndonesiaPresentOgawa et al., 1992
JapanPresentOgawa et al., 1992; Urawa and Awakura, 1994; Awakura et al., 1995; Székely et al., 2003
Korea, Republic ofPresentCho and Kim, 2003
TaiwanPresentOgawa et al., 1992; Chen et al., 2001

Africa

BeninPresentGbankoto et al., 2001
EgyptPresentEl-Mansy, 2002
TunisiaPresentBahri and Marques, 1996

North America

CanadaPresentMargolis, 1993
-British ColumbiaPresentKent et al., 1997; Jones et al., 2003; Jones et al., 2004
-Newfoundland and LabradorPresentMaloney et al., 1991
-Nova ScotiaPresentDzulinksy et al., 1994
-OntarioPresentXiao and Desser, 1998a; Xiao and Desser, 1998b; Xiao and Desser, 2000a; Xiao and Desser, 2000c; Xiao and Desser, 1997
USAPresentHedrick et al., 1990; Kent et al., 1996
-CaliforniaPresentHendrickson et al., 1989; Modin, 1998
-ColoradoPresentAllen and Bergersen, 2002
-FloridaPresentLandsberg, 1993
-IowaPresentMitchell et al., 1980
-MichiganPresentPronin et al., 1997
-MinnesotaPresentCone et al., 1997
-MontanaPresentMitchell, 1989
-OregonPresentRatliff, 1983
-WashingtonPresentJones et al., 2004

South America

BrazilPresentMartins and Souza, 1997; Martins et al., 1997

Europe

FinlandPresentBrummer-Korvenkontio et al., 1991; Feist and Rintamäki, 1994
Former USSRPresentSchulman, 1966
GermanyPresentKaup et al., 1995; Oumouna et al., 2003
HungaryPresentEl-Mansy et al., 1998a; El-Mansy et al., 1998b; Molnár, 2002a; Molnár, 1998; Molnár and Székely, 1999
IrelandPresentFrasca et al., 1998; Frasca et al., 1999
NorwayPresentHemmingsen et al., 1991; Larsen et al., 1997; Karlsbakk et al., 2002; Sterud et al., 2003
SpainPresentAlvarez-Pellitero et al., 1982; Gonzalez-Lanza and Alvarez-Pellitero, 1984; Gonzalez-Lanza and Alvarez-Pellitero, 1985; Alvarez-Pellitero, 1989; Palenzuela et al., 1997; Székely et al., 2000
UKPresentMcGeorge et al., 1996a; Andrews, 1979; Seagrave et al., 1981; Davies, 1985; Crawshaw and Sweeting, 1986; Lom et al., 1991; Feist et al., 2002; Ozer et al., 2002; Longshaw et al., 2003

Oceania

AustraliaPresentLangdon, 1987; Langdon, 1990; Roubal, 1994; Rothwell et al., 1997
New ZealandPresentMeglitsch, 1960

Pathology

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Diseases According to Host Organ

Gills

Infections of the gill can compromise the respiratory capability if present in sufficient numbers, and disruption of the gill epithelium by the release of myxospores can provide a route of entry for secondary infections. Pathology associated with myxozoan infections in the gills includes fusion of lamellae, inflammation, hyperplasia, pressure atrophy and cellular necrosis. Molnár (2002b) proposed a system to describe the specific site locations of myxozoans in gills. This distinguishes between interlamellar epithelial and intralamellar vascular types for plasmodia in the gill lamellae, with either chondroidal, vascular or epithelial intrafilamental types developing in the gill filaments.

Gill sphaerosporosis of carp, caused by Sphaerospora molnari, has been reported in Cyprinus carpio and Carassius carassius in Europe (Dyková and Lom, 1988a; Fig. 9). Additionally, Hedrick et al. (1990) reported the presence of Sphaerospora cf. chinensis in the gills of Carassius auratus. In both infections, moderate to severe gill hyperplasia results and a large proportion of the respiratory epithelium can be replaced by sporogonic stages. In mild S. molnari infections, local circulatory disorders and dystrophic changes also occur. S. molnari spores measure 10.5 µm x 10.3 µm and those of S. chinensis measure 7.4 µm x 7 µm. Both the parasites form monosporic pseudoplasmodia.

Proliferative gill disease (PGD, hamburger gill disease), caused by the extrasporogonic stage of Henneguya ictaluri Pote, Hanson and Shivaji, 2000, is a major disease of channel catfish (Ictalurus punctatus). and can result in high mortalities amongst juvenile farmed catfish (Pote et al., 2000). The parasite elicits a strong granulomatous inflammatory response in the gills. Styer et al. (1991) demonstrated experimentally that the life cycle alternated between the fish host and the oligochaete Dero digitata. Both stages in the life cycle have been confirmed using molecular techniques (Pote et al., 2000; Hanson et al., 2001). The spore body of the myxospore measures 24 µm x 6 µm and the total spore length is 48-80 µm.

Another pathogenic Henneguya sp. in the gills of I. punctatus is Henneguya exilis Kudo, 1929. The presence of plasmodia at the base of the secondary lamellae (epithelial filamental type of Molnár, 2002b) elicits a severe tissue proliferation in this atypical site. Known as 'lamellar disease', it can lead to high mortalities of fingerlings. The formation of cysts in other sites is not pathogenic. The alternate stage is an Aurantiactinomyxon type in the oligochaete Dero digitata (Lin et al., 1999). Spores measure 17.6 µm x 4.9 µm, and total length is 52.3 µm.

Fig. 9. Histological sections of gill, pseudobranch and fin infections. A. Giemsa-stained section of carp (Cyprinus carpio) gill infected with Sphaerospora molnari (inset: mature spore in sutural view). B. Large cyst of Myxobolus koi in the gill of koi carp (C. carpio) (inset: mature spore in valvular view). C. Multiple cysts of Henneguya psorospermica in the gill of pike (Esox lucius). D. Multiple cysts of Myxobolus macrocapsularis in the gill of chub (L. cephalus) (inset: mature spore in valvular view). E. Cysts of M. macrocapsularis in the pseudobranch of dace (L. leuciscus). F. Sporogonic plasmodium of an unidentified Myxobolus sp. in the cartilage of the caudal peduncle of L. cephalus. G. Longitudinal section through the fin of a juvenile roach (R. rutilus) showing Myxobolus sp. cysts within the epithelium. H. Myxobolus sp. cysts in connective tissue of the fin of barbel (Barbus barbus).

The pathology associated with Henneguya creplini (Gurley, 1894) on the gills of Stizostedion lucioperca in Lake Balaton was described by Molnár (1998). Plasmodia on the gills are large, reaching 900 µm x 750 µm in size, with an average of 30-60 plasmodia per fish. During cyst development, host responses are minimal. Following sporogenesis and rupture of plasmodia, an intense host response, consisting of epithelial proliferation, overgrowth of the damaged plasmodium and cell necrosis, becomes apparent. Spores are 14.3 µm x 5.5 µm, with a total length of 43 µm including caudal appendages. Infections in pike (Esox lucius) and perch (Perca fluviatilis) caused by Henneguya psorospermica Thélohan, 1895 also give rise to multiple large plasmodia displacing the gill tissue (Fig. 9).

Henneguya piaractus Martins and de Souza, 1997 in the gills of captive Piaractus mesopotamicus can cause mortalities (Martins and de Souza, 1997; Martins et al., 1997). Clinical signs of infection include decreased feeding, lethargy, erratic swimming and loss of equilibrium. Histologically, haemorrhaging and severe inflammation occur in the gill epithelium and larger cysts cause pressure atrophy on adjacent lamellae. Spores measure 11.3 µm x 3.2 µm, total length, including caudal process 48.4 µm.

Infections with Myxobolus basilamellaris Lom and Molnár, 1983 cause a localized pathological response to the parasite cysts within the gills of its host (Kovács-Gayer and Molnár, 1983). The cysts develop in a basifilamental position on the gill. Growth of the cyst lifts the basal part of the gill filament and deforms adjoining lamellae, reducing the respiratory surface. Those cysts developing in the gill arch restrict nerves and blood vessels passing through the gill arch, leading to local blockages.

Myxobolus koi forms large and small-type cysts in the gills of C. carpio (Fig. 9). The smaller cysts develop in an interlamellar position within the gills and host responses are minimal. In contrast, large cysts produce extensive pathological changes in the gill, including hypertrophy of the branchial epithelium and clubbing of the gill filaments (Yokoyama et al., 1997). Spores measure 12-15 µm x 5-9 µm and contain two equal-sized polar capsules measuring 6-7.4 µm x 1.6-2.7 µm.

Myxobolus pavlovskii infects the gills of bighead (Aristichthys nobilis) and silver carp (Hypophthalmichthys molitrix) and in young cultured fish can cause pathological damage to the gill structure due to the development of plasmodia affecting respiratory function (El-Matbouli et al., 1992a). Parasite plasmodia develop in an interlamellar and basifilamental position on the gills (Molnár, 2002b). The parasite has been shown to undergo a two-host life cycle, producing Hexactinomyxon-type actinospores in the oligochaete Tubifex tubifex (El-Matbouli and Hoffmann, 1991a). Myxospores measure 7-9.5 µm x 5-8 µm.

Serious infections of the pseudobranch are uncommon, although infections of this organ have been recorded (Molnár, 2002b). Mortality of farmed Salmo salar associated with Parvicapsula pseudobranchicola Karlsbakk, Sæther, Høstlund, Fjellsøy and Nylund, 2002 has been reported. The Parvicapsula species reported by Sterud et al. (2003) is likely to be P. pseudobranchicola, based on tissue location, pathogenesis and geographical location, although there are some minor differences in the spore dimensions. Clinical signs include increased surface swimming, lethargy, lack of feeding, exophthalmia and mortality. Petechial haemorrhaging of infected pseudobranchs is observed. It has been suggested that the infection ultimately leads to blindness of the host due to loss of pseudobranch function which supplies blood to the eye (Karlsbakk et al., 2002; Sterud et al., 2003). The spores measure 12 µm in length and 6.2 µm in thickness and possess two slightly unequal polar capsules measuring 2.4 and 2.3 µm in diameter (Karlsbakk et al., 2002).

Skin, scales, fins and subcutaneous tissues

Infections of the skin, scales or subcutaneous tissues are generally not considered serious (Fig. 9). However, in fish destined for the aquarium trade or for human consumption, they can be considered problematic as they reduce the aesthetic appeal of the fish and their value. Typical of this group is Myxobolus diversus Nie and Li, 1973 in the fins of goldfish. Whilst serious pathological changes are generally absent, the obvious presence of myxozoan cysts on the fins reduces the value of the fish (Molnár and Székely, 2003).

Molnár (2002a) described the characteristic types and development of myxozoan plasmodia in fins. Plasmodia developing within the skin between the fin-rays and inside the lumen of the fin-rays produced limited host responses. However, plasmodia developing on the outer surfaces of the fin-rays, as typified by Thelohanellus nikolskii, markedly deform the cartilaginous elements of the fin-rays of carp (Molnár, 1982; Moshu and Molnár, 1997). T. nikolskii has also been reported on the scales of C. carpio carpio (common carp), where plasmodial development is associated with calcified collagen but with low pathogenicity (Moshu and Molnár, 1997). C. c. carpio, C. auratus (goldfish) and C. carpio haematopterus (koi carp) possess differential susceptibility to infections by T. nikolskii with C. c. haematopterus being less susceptible compared with C. c. carpio, and C. auratus being resistant (Molnár, 2002c). Székely et al. (1998) demonstrated that T. nikolskii alternates through T. tubifex, producing an Aurantiactinomyxon type with a spore body measuring 21 µm in diameter and possessing caudal processes measuring 13 µm x 9 µm. The myxospore measures 16.5 µm x 10 µm and contains a single polar capsule measuring 6.5 µm x 5.6 µm.

Thelohanellus hovorkai, a parasite of the connective tissue, causes haemorrhagic thelohanellosis in koi carp. The disease is manifested by haemorrhaging and oedema in skin lesions, with destruction of the capillary network caused by the release of mature spores into the surrounding tissues, and chronic mortalities (Yokoyama et al., 1998). The parasite alternates through the oligochaete Branchiura sowerbyi, producing an Aurantiactinomyxon-type actinospore with a spore body 19 µm in diameter and possessing caudal processes measuring 29 µm x 9 µm (Székely et al., 1998). Actinospores enter the fish host via the gills (Yokoyama and Urawa, 1997). Myxospores measure 20 µm x 10 µm and possess a single polar capsule measuring 9 µm x 8 µm.

Thelohanellus wuhanensis Xiao and Chen, 1993 is a serious disease of Carassius auratus gibelio that causes large swellings on the skin of affected fish and leads to mortality of infected fry (Wang et al., 2001). Myxospores measure 23 µm x 14 µm and contain a single polar capsule measuring 10.5 µm x 8 µm.

Several Myxobolus species in the skin and subcutaneous tissues have been reported as pathogens. Myxobolus drjagini (Akhmerov, 1954) forms small plasmodia in the skin of the operculum, head and buccal cavity of Hypophthalmichthys molitrix and causes twist disease in China (Wu et al., 1975). El-Mansy and Molnár (1997a) did not consider that it caused losses in Hungary. The parasite alternates through T. tubifex and produces a triactinomyxon-type actinospore. Myxospores measure 13 µm x 7 µm and contain two unequal-sized polar capsules measuring 5.8 µm x 3.7 µm and 3.4 µm x 2.5 µm. The market value of C. auratus auratus is reduced as a result of infections with Myxobolus rotundus (Nemezek, 1911), that causes large, visible cysts on the body surface (Lu et al., 2003).

Myxobolus episquamalis Egusa, Maeno and Sorimachi, 1990 infects the scales of Mugil cephalus. Small, pale, raised lesions are present on infected fish and can lead to erosion of the scales and attenuation of the dermis. Whilst the infection is of little pathological consequence, Rothwell et al. (1997) considered that up to 6% of the catch were unsaleable due to the loss of aesthetic quality of the fish.

Muscle

There are many reports of myxozoan infections in muscle of both marine and freshwater fishes worldwide. They range from innocuous infections with minimal host response to intense infections leading to mortality or spoilage of the musculature through enzymatic degradation on the death of the host. Typically, host responses to the parasite are usually not initiated until sporogenesis of the myxospores is complete. Whilst the majority of infections are reported from somatic muscle, any organ containing muscle tissue, including the heart, intestine and gills, can be affected. In addition, the presence of myxozoan development stages in muscle, which normally occur in other host tissues, can lead to pathological changes in the musculature (Fernández-de-Luco et al., 1997).

The synonymy of Henneguya zschokkei Doflein, 1901 from whitefishes (Coregonus spp.) and Henneguya salminicola Ward, 1919 from salmonids (Oncorhynchus and Salmo spp.) is problematic. Whilst some authors consider H. salminicola as a junior synonym of H. zschokkei, Eiras (2002) included them as separate species in a synopsis of the genus. Recent molecular data have provided further evidence that they are distinct species (Kent et al., 2001). For both, heavy infections within the somatic muscle can render the flesh unmarketable as the production of fillets can cut through these cysts to produce a milky exudate (Awakura and Kimura, 1977). The spore body of H. salminicola measures 12 µm x 8 µm and total spore length is 43-52 µm. The spore body of H. zschokkei measures 10 µm x 7 µm and total spore length is 55 µm (Fig. 3).

In heavy infections with M. artus in C. carpio, the body surface of the host is 'irregular and uneven' due to the presence of numerous cysts in the muscle (Ogawa et al., 1992). Two types of cysts are found - a larger, interfibrillar type and a smaller intrafibrillar type. Following sporogenesis, spores are subjected to a host response and the rupture of pseudocysts leads to the lysis of adjacent muscle fibres. Spores are phagocytosed and transported to other organs in the body via the blood (Fig. 10). Yokoyama et al. (1996) showed that carp release up to 3 x 105M. artus spores per fish per day throughout their lifetime, possibly via the intestine and skin. Spores measure 8 µm x 11 µm in size.

Cyprinid fishes are parasitized by a number of Myxobolus spp. in the musculature (Fig. 10). Some workers have considered that M. pseudodispar from the musculature of roach is a junior synonym of Myxobolus cyprini Doflein, 1898 due to the pleomorphic nature of the spore. Molnár et al. (2002) used DNA sequence data to demonstrate that M. pseudodispar Gorbunova, 1936 from a number of hosts, Myxobolus musculi Keysselitz, 1908 from barbel (Barbus barbus) and M. cyprini from C. carpio were distinct species. In addition, barbel are hosts to Myxobolus pfeifferi Thélohan, 1895 in the muscle, which has caused epizootics of 'barbel boil disease'. M. Longshaw (unpublished data) found M. pfeifferi in the musculature of juvenile Leuciscus cephalus, Leuciscus leuciscus and Rutilus rutilus without significant host response. For all four Myxobolus spp. the pathogenesis of the disease is similar to that reported for M. artus, whereby spores develop within cysts in the myofibrils. Growth of cysts results in marked enlargement of infected muscle fibres and pressure atrophy of adjacent myofibrils. On completion of sporogenesis, spores are subjected to a vigorous granulomatous response involving epitheloidtype cells and phagocytic cells. Infected myofibrils are destroyed and localized blockage of vessels and necrosis of the surrounding tissue can occur (Fig. 10). Spores are transported around the body via the blood and lymphatic system to various sites, including the thymus, liver, kidney, spleen, gills and intestine. The majority of infections in carp by M. cyprini are considered to cause only subclinical damage to the muscle and local necroses in other organs but can, in some cases, cause hydropic degeneration and dropsy (Molnár and Kovács-Gayer, 1985). In roach infected with M. pseudodispar, similar effects have not been reported (Baska, 1987; Athanassopoulou and Sommerville, 1993b). Spore measurements are as follows: M. cyprini 10-16 µm x 8-12 µm (polar capsules equal in size, measuring 5-7 µm in length); M. musculi 9-13 µm x 8-11 µm (polar capsules unequal in size, measuring 4.5-7 µm x 3-4 µm and 4-6 µm x 2-3.5 µm); M. pseudodispar 10-12 µm x 7-9.5 µm (polar capsules unequal in size, measuring 4.5-5.5 µm x 3 µm and 4 µm x 3 µm); M. pfeifferi 10-13 µm x 9-12 µm (polar capsules equal in size, measuring 5-6 µm x 2.5-3.5 µm).

Fig. 10. Histological features of muscle-invading species. A. Sporogonic plasmodia of Myxobolus artus in the skeletal muscle of carp (C. carpio) (inset: mature spore in valvular view). B. Granulomatous host response to a plasmodium of M. artus within the renal tissue (same fish as in Fig. 10A). C. A plasmodium of Myxobolus pseudodispar within the muscle of a juvenile roach (Rutilus rutilus). D. Focal inflammatory response to M. pseudodispar spores and sporogonic stages from a ruptured plasmodium. E. Intra-myofibrillar Kudoa infection in cow-nosed ray (Rhinoptera bonasus). F. Multiple plasmodia of Kudoa sp. in the myofibrils of common goby (Pomatoschistus microps). G. Giemsa-stained section of Kudoa thyrsites infection in cod (Gadus morhua). H. Fibrous encapsulation of a Kudoa sp. plasmodium in viviparous blenny (Zoarces viviparus).

Myxobolus lentisuturalis Dyková, Fiala and Nie, 2002 from Carassius gibelio induces severe regressive changes in the dorsal somatic musculature. Externally, the disease can manifest itself as a large swelling anterior to the dorsal fin, caused by the developing parasite cyst. In heavily infected individuals, necrotic changes occur in infected myofibrils. Spores measure 11.8 µm x 7.6 µm, with equal-sized polar capsules measuring 4.2 µm x 2.5 µm.

Intense infections with a variant of Myxobolus procercus (Kudo, 1934) were reported in the musculature of Percopsis omiscomaycus by Cone et al. (1997). Host responses to sporogonic forms of the parasites were in approximately 5% of the fish examined; otherwise fish appeared healthy. Cone et al. (1997) considered that the extremely large numbers of myxozoan cysts in the muscle may be due to an over abundance of oligochaetes and the depressed immune system of the fish host. Spores measure 14 µm x 6 µm, and polar capsules are often dissimilar in size measuring approximately 7.2 µm x 2.2 µm.

Most muscle infections in freshwater fishes are caused by bivalvulid myxospores and the host response is usually towards mature spores. In marine fish, however, myxozoan infections in the muscle are frequently caused by multivalvulid myxospores, which in a few cases result in the enzymatic destruction of the muscle tissue following the death of the host. Typical of this group are members of the genus Kudoa, reviewed by Moran et al. (1999b) (Fig. 10). Fish can be unmarketable due either to the presence of large, macroscopic, melanized cysts within the musculature or to post-mortem myoliquefaction. At least 50 species of Kudoa have been described from marine fishes worldwide. Kudoa thyrsites is known to infect more than 20 species of fish from a number of different families worldwide, including wild and farmed fish (Kent, 2000). Myoliquefaction has also been reported in Seriola holandi infected with Unicapsula seriolae Lester, 1982 and Kudoa (= Hexacapsula) neothunni (Arai and Matsumoto, 1953).

Ulcerative lesions in young-of-the-year Brevoortia tyrannus have been attributed to a number of aetiologies, including bacteria, fungi and harmful algal blooms. Reimschuessel et al. (2003) demonstrated that ulcers and chronic inflammatory infiltrates in this host were associated with invasive extrasporogonic plasmodia of Kudoa clupeidae (Hahn, 1917). Spores and plasmodia were present within the musculature of these fish and, whilst the authors were unable to unequivocally demonstrate that the plasmodia causing ulceration were early stages of K. clupeidae, strong circumstantial evidence was provided to suggest that these invasive extrasporogonic stages were responsible for the ulcerations.

Cartilage and ossified elements

Infections of cartilage and bone are relatively rare, and parasites infecting the host prior to ossification often become trapped within the ossifying or fully ossified elements. Infections of the cartilage can result in hypertrophy of cartilage and inflammation of surrounding tissues.

Myxobolus cerebralis is the causative agent of salmonid whirling disease. Clinical signs of the disease can include melanization of the tail region and skeletal deformities. The characteristic 'whirling' motion and tail chasing of infected fish are thought to be a consequence of constriction and pressure on the spinal cord (Rose et al., 2000). M. cerebralis was first recorded infecting brown trout from Central Europe and northeast Asia, which were transferred to a number of sites worldwide through the movement of live fish and fish products (Bartholomew and Reno, 2002; Bartholomew and Wilson, 2002). Evidence from the internal transcribed spacer (ITS) region of the ribosomal DNA (rDNA) confirmed that M. cerebralis was introduced to several countries from a limited geographical source (Andree et al., 1999a; Whipps et al., 2004b). It now occurs in Europe, the USA, South Africa and New Zealand. Movements of live infected fish from hatcheries and fish farms possibly increased the geographical spread of the pathogen within individual countries. Whilst there is strong evidence from other studies in some regions of the USA that M. cerebralis is responsible for declines of wild rainbow trout (Hedrick et al., 1998), Modin (1998) considered that 'there is little clear evidence of significant impacts on wild salmonids in California'.

Wolf and Markiw (1984) demonstrated that the parasite required an oligochaete host in its life cycle and subsequent studies have shown that the parasite is extremely specific to T. tubifex (Granath and Gilbert, 2002) and possibly even to distinct genetic lineages of T. tubifex (Beauchamp et al., 2001). Like numerous other myxozoans, it has been presumed that myxospores from fish were only available to the oligochaete host upon the death and decay of the fish host. However, Nehring et al. (2002) have demonstrated experimentally that live brown trout expel viable M. cerebralis myxospores, which are infective to T. tubifex. Sporoplasms from waterborne triactinomyxon actinospores enter the host via a number of entry points, including the mucus cells of the epithelium, gills and buccal cavity (Markiw, 1989; El-Matbouli et al., 1995, 1999b). Entry and establishment of the disease in fish is determined by the degree of ossification and, whilst adult, ossified fish can be infected, they are asymptomatic for the disease (Markiw, 1992). Following penetration of the epithelium by the triactinomyxon, sporoplasms reach the peripheral nerves, then the central nervous system and eventually the cartilaginous elements. During migration through the nervous tissues, the parasite undergoes a series of proliferative stages and appears to be unrecognized during this stage by the host immune system.Within the cartilaginous elements, phagocytosis of chondrocytes by plasmodia has been reported and the lysis of cartilage by the parasite can induce a vigorous host response (Fig. 11). Inflammatory cells involved in the host response include mononuclear leucocytes, and the combination of cartilage lysis with a vigorous host response can lead to severe disruption of the cartilage and result in skeletal deformities. M. cerebralis spores are highly variable in morphology, but in general are oval to circular in valvular view and measure 7-10 µm x 7-10 µm in size. Polar capsules are equal in size, measuring 4-6 µm x 3-3.5 µm, and contain five to six turns of the polar filament. The advent of molecular tools such as PCR and in situ hybridization has assisted in the identification and phylogeny of the parasite in the oligochaete and fish hosts and provided further insights into its biology (Andree et al., 1997, 1999b; Antonio et al., 1998).

Infections with Myxobolus buckei Longshaw, Frear and Feist, 2003, originally reported by Bucke and Andrews (1985) as Myxobolus ellipsoides Thélohan, 1892, cause serious disease in juvenile cyprinids in the UK, resulting in marked spinal compression in infected fish. Normal and burst swimming of infected fish is compromised and few juvenile fish survive to adulthood (Longshaw et al., 2003). Histological lesions in infected fish range from mild hypertrophy of the zygapophyseal elements to complete fusion of the vertebrae (Fig. 11). Hypertrophy of the vertebral elements and expansion of the intervertebral membrane to accommodate the developing plasmodia cause pressure on the spinal cord and adjacent blood vessels. Fish are infected prior to ossification and the parasite develops initially within the remnants of the embryonic notochord. A vigorous inflammatory response to the developing plasmodia and spores leads to the radial expansion and fusion of the vertebral centra during the cartilaginous phase of host development. Following ossification, parasites remain trapped within bony elements. Spores measure 14 µm x 11.5 µm in size.

Myxobolus aeglefini has been reported from a number of marine fishes in the North Atlantic, North Sea and Baltic Sea (Lom and Dyková, 1992). It has a wide host specificity, having been reported in flatfish and gadoids (Nielsen et al., 2002). The report by Yokoyama and Wakabayashi (2000) of M. aeglefini in cartilaginous tissues of the skeletal musculature of Japanese poroushead eelpout may represent a morphologically similar but genetically different species of Myxobolus. Whilst M. aeglefini is normally considered as a parasite with a specificity for head cartilage, it can be found in the eyes of gadoid hosts (Fig. 11). As with other cartilage-infecting myxozoans, damage to the host is caused by a combination of the host response and lysis of the cartilage by the action of the parasite. Spores measure 11-13 µm x 10-12 µm in size.

Fig. 11. Pathological features of cartilage infections. A. Low-power view of head cartilage of rainbow trout (Oncorhynchus mykiss) infected with Myxobolus cerebralis, showing disintegration of cartilaginous elements. B. High-power view showing spores and developmental stages of M. cerebralis amongst destroyed cartilage (A and B are from Giemsa-stained sections). C and D. Vertebral deformation and complete fusion of vertebrae in chub (L. cephalus) caused by Myxobolus buckei (C. inset: mature spore of M. buckei in valvular view). E. Cysts of Myxobolus aeglifini in the scleral cartilage of the eye of poor cod (Trisopterus minutus) (inset: mature spore of M. aeglifini in valvular view). F. Section through two large cysts of M. aeglifini in the scleral cartilage of whiting (Merlangius merlangus). G. Myxobolus sp. in the cranial cartilage of common goby (Pomatoschistus microps). H. Cysts of Myxobolus sp. in the cranial cartilage of roach (R. rutilus).

Reproductive tissues

Myxozoan infections of gonadal tissue are relatively rare and appear to be sex specific. Infections of the ovaries occur either within oocytes or within the interstitial or connective tissues, with those species occurring within the oocytes being more serious. In both male and female fish, infections can lead to reductions in fecundity by castration or gonadal regression. Negative impacts at the host population level may be possible but have not been reported to date.

Sphaerospora testicularis Sitja-Bobadilla and Alvarez-Pellitero, 1990 infects the seminiferous tubules in the testis of Dicentrarchus labrax. Infections lead to testicular hyalinization and hypertrophy, occlusion of the tubular lumens, ascites in the abdominal cavity and damage to the germ and Sertoli cells. Disporic pseudoplasmodia contain spores measuring 12 µm x 15 µm in size. Agarella gracilis Dunkerley, 1915 has been reported in the testis of Lepidosiren paradoxa in Brazil, with no data on the impact of the parasite. Spores measure 28-35 µm x 13-16.5 µm in size. Testicular tissue of Girardinus januarius is destroyed by spores of Myxobolus lutzi Aragão, 1919. Spores measure 10 µm x 7 µm in size.

Henneguya oviperda (Cohn, 1895) Labbé, 1899 has been reported from the ovaries of pike E. lucius in Russia and other European countries. In heavy infections, it causes complete degeneration of ovarian tissue and oocytes. Parasite cysts are found in the connective tissue and follicle epithelium of the ovary, whilst spores are found in the oocytes. Spores measure 27-42 µm x 5-10 µm, including caudal appendages.

Sphaerospora ovophila Xiao and Desser, 1997 spores in the ovary of Lepomis gibbosus produce visible cysts measuring up to 500 µm in diameter. Spores measure 8.2 µm x 6.2 µm in size. Myxobolus algonquinensis Xiao and Desser 1997 from Notemigonus crysoleucas produces cysts in the connective tissue of ovaries or interstitial tissue among oocytes measuring up to 800 µm in diameter. Spores measure 14.7 µm x 10.9 µm in size. No pathology was reported with these two forms (Xiao and Desser, 1997).

Kudoa ovivora Swearer and Robertson, 1999 occurs in the ovaries of Thalassoma bifasciatum and other labroid fishes. White-coloured plasmodia are restricted to the inner wall of the oocyte membrane and spores are only found in late developmental stage oocytes undergoing or completing vitellogenesis. Eggs are non-viable and infected females show reduced growth, fecundity and/or spawning activity. Spores measure 6.5 µm x 7.7 µm in size.

Myxobolus dahmeyensis (Siau, 1977) was reported in the ovaries of Tilapia zilli and Sarotherodon melanotheron melanotheron by Gbankoto et al. (2001). No parasite cysts were produced and spores were only present in mature oocytes, which were destroyed by the parasite, leading to castration of the host.

Henneguya amazonica Rocha, Matos and Azevedo, 1992, originally reported from the gills of Crenicichla lepidota, is found mainly within the ovarian stroma but also within the follicles of Hoplosternum littorale. Whilst Torres et al. (1994) reported that the morphology of infected oocytes was not significantly affected, they could not discard the possibility that ovaries may be destroyed by the parasite. Spore body measures 13.9 µm x 5.7 µm; total spore length, including caudal appendages, is 59.3 µm.

Central nervous system

Over 20 myxosporean species have been reported infecting the brain and spinal cord of teleosts (Hoffmannn et al., 1991; Frasca et al., 1999; Cho and Kim, 2003). In addition, unidentified myxozoan-like parasites have been described infecting the spinal cord of the toad (Bufo bufo) (Stensaas et al., 1967) and more recently from the brain of the mole (Talpa europea) (Friedrich et al., 2000). In these cases, only plasmodial stages containing secondary (occasionally tertiary cells in the mole parasite) were observed and no significant host response was recorded.

In general, myxosporeans infecting nervous tissue do not elicit a significant host response. Examples include Myxobolus galaxii Szidat, 1953 from the spinal cord of Galaxias olidus, a salmoniform freshwater fish from Australia (Langdon, 1990), Myxobolus neurophilus (Guilford, 1963) in yellow perch (Perca flavescens) (Dzulinksy et al., 1994) and Myxobolus inaequus Kent and Hoffmann, 1984 in South American knife fish (Eigenmannia virescens) (Kent and Hoffmann, 1984). However, encephalitis associated with presporogonic and sporogonic stages occurs with some species (Dyková and Lom, 1988a; Frasca et al., 1998). In others, a significant proportion of the nervous tissue can be replaced with parasitic cysts. Infections with Myxobolus hendricksoni Mitchell, Symour and Gamble, 1985 in fathead minnows (Pimephales promelas) result in macroscopic cysts up to 1.5 mm in diameter throughout the brain, in particular the optic lobes and corpus cerebrelli (Mitchell et al., 1985). Even with extensive infections, behavioural abnormalities have rarely been reported. Fish infected with M. arcticus have reduced mean swimming speed compared with uninfected fish (Moles and Heifetz, 1998), and infections with Myxobolus encephalicus (Mulsow, 1911) in carp fry can result in loss of equilibrium and circling motion. Plasmodia of M. encephalicus grow within the blood vessels of the brain and cause congestion of the vessels and focal oedema. Sporogonic stages invoke a severe granulomatous response (Dyková and Lom, 1988a). Neurological symptoms, including uncoordinated darting and rolling movements, are associated with infections of Myxobolus balantiocheirli Levsen, Alvik and Grotmol, 2004 in tricolour sharkminnow (Balantiocheilos melanopterus). In this case, parasite-induced inflammation is largely absent (Levsen et al., 2004).

There have been several reports of skeletal deformities associated with myxosporidiosis of neural tissues. Triangula percae Langdon, 1987 was described from redfin perch (Perca fluviatilis) from Victoria, Australia, where juveniles appear to be more susceptible to the infection and develop marked curvature (lordodis) of the spine. The parasites are located in the white matter of the mesencephalon, the diencephalon and the medulla oblongata (Langdon, 1987). In Scotland, Myxobolus sandrae Reuss, 1906 also causes lordosis and vertebral compression of the spine in the same host. Atrophy and demyelinization of the white matter of the spinal cord is the principal host response to the presence of sporogonic plasmodia (Lom et al., 1991). Egusa (1985) described infections with Myxobolus buri Egusa, 1985 causing severe scoliosis in cultured Seriola quinqueradiata in Japan. Several regions of the brain were affected, including the olfactory and optic lobes, cerebellum and medulla oblongata. Parasite stages are surrounded by a fibrous capsule and pericystic inflammation. Spinal curvature in mullet (M. cephalus) from Japan is caused by Myxobolus spinacurvatura Maeno, Sorimachi, Ogawa and Egusa, 1990. Cysts approximately 1 mm in diameter occupy several regions in the brain, including the olfactory and optic lobes, and also infect mesenteric tissue and the spleen (Maeno et al., 1990).

Bullheads (Cottus gobio) from the Czech Republic, southern England and Germany are hosts to Myxobolus cotti El-Matbouli, Fischer-Scherl and Hoffmannn, 1990, where plasmodia infect several regions of the brain and are also found within axons (Lom et al., 1989b; El-Matbouli et al., 1990; see Fig. 15). Intra-axonal myxosporean plasmodia have also been described from the spinal cord and brainstem of the common shiner (Notropis cornutus) (Ferguson et al., 1985).

Salmonids are hosts to several braininfecting myxosporeans. M. neurobius infects brown trout (Salmo trutta), Atlantic salmon (S. salar) and grayling (Thymallus thymallus) in Europe and Asia and Pacific salmonids in North America (Gonzales- Lanza and Alvarez-Pellitero, 1984; Maloney et al., 1991). Plasmodia are up to 400 µm in diameter. Spores are ovoid, 10-14 µm x 8-9.2 µm in size. Although generally regarded as non-pathogenic, intense infections can induce pressure atrophy of surrounding tissues. A similar Myxobolus sp. causes 'sleeping disease' in cultured masu and amago salmon in western Japan (Murakami, 1979, cited by Urawa and Awakura, 1994). In Pacific salmonids M. arcticus can be found at prevalences approaching 100% (Kent et al., 1993; Awakura et al., 1995). Spores are pyriform, measuring approximately 15 µm x 7.5 µm. M. arcticus is currently the only myxosporean parasite with a tropism for neural tissue for which the life cycle has been elucidated. Transformation of M. arcticus myxospores occurs in the oligochaete Styrodrilus heringianus, with triactinomyxon spores released from that host being infective to sockeye salmon. Mature myxospores develop in the neural tissue 3-4 months after exposure (Kent et al., 1993).

Neurotropic presporogonic stages of M. cerebralis, identified using small subunit rDNA sequence data and in situ hybridization, have been linked with neurological disease and mass mortality of Atlantic salmon smolts at a marine culture facility in Ireland (Frasca et al., 1998, 1999). Clinical signs are absent. Plasmodia are present between axons, most frequently in the optic tectum of the mesencephalon. Pathology ranged from mild to severe focal encephalitis, involving glial cells, macrophages and lymphocytes. It was hypothesized that infective stages originated from nearby riverine inputs to the farm location (Frasca et al., 1999).

Multivalvulid myxosporeans of the genus Kudoa have been described in the brain of cultured and wild marine fish. Kudoa paralichthys Cho and Kim, 2003 infects olive flounder in Korea (Cho and Kim, 2003), whilst Kudoa tetraspora Narasimhamurti and Kalavati, 1979 and Kudoa cerebralis Paperna and Zwerna, 1974 infect mullet (M. cephalus) and striped bass (Morone saxatilis), respectively (Paperna and Zwerna, 1974Narasimhamurti and Kalavati, 1979). In all of these infections, the host response is minimal. Whereas all neurotropic myxosporeans appear to sporulate in the central nervous system (CNS), a Kudoa sp. infecting Thunnus maccoyii from Western Australia usually sporulates in peripheral nerves (Langdon, 1990). Kudoa (= Pentacapsula) neurophila (Grossel, Dykova, Handlinger, and Munday, 2003) causes severe granulomatous meningoencephalomyelitis amongst cultured juvenile striped trumpeter (Latris lineata) in Tasmania (Grossel et al., 2003) and Kudoa (= Septemcapsula) yasunagai (Hsieh and Chen, 1984) forms large parasitic cysts in cultured Lateolabrax japonicus and Oplegnathus fasciatus in China and Japan (Hsieh and Chen, 1984).

Infections of the kidney and urinary tract

The kidney is the main target organ for a large number of myxosporean species. The various metabolic functions of the organ related to excretion, osmoregulation and haematopoiesis, as well as the presence of endocrine tissue, such as the corpus of Stannius, and inter-renal tissue, are reflected in its complex structure. Coelozoic species reside in all cavities, from Bowman's capsule of the glomerulus and the various compartments of the nephron to the renal collecting ducts and the urinary bladder. Histozoic species (e.g. Myxobolus spp.) are commonly found as small accumulations of loose or phagocytosed spores or contained in larger groups within macrophage aggregates, where they may have been transported from other organs for elimination from the host. In addition, they can occur as encysted plasmodia within the haematopoietic interstitial tissues (e.g. Myxidium rhodeiLéger, 1905). Histozoic and intracellular proliferative extrasporogonic stages of some myxosporean and malacosporean species induce a significant pathological host response, resulting in diseases such as PKD of salmonids, caused by T. bryosalmonae, and sphaerosporosis in carp, caused by Sphaerospora renicola. Renal infections of endocrine tissues have not been reported thus far. The principal histopathological features of renal myxosporidiosis range from acute to chronic inflammation to extensive tissue necrosis and fibrosis and are dependent on a variety of factors, including the immune status of the host, the pathogenicity of the parasite and the stage of development of the infection. Renal damage can be extensive, resulting in functional impairment. However, the host is often able to survive unless placed under additional environmental stress and/or secondary challenge from other pathogens.

Infections with the malacosporean T. bryosalmonae (formerly PKX) are responsible for serious economic loss in salmonid culture throughout Europe and North America, although the impacts on wild fish populations are less certain (Feist et al., 2002; Wahli et al., 2002). Infection of fish hosts is initiated when infective spores released from bryozoans enter the fish through the skin and gill epithelium, either between epithelial cells or intracellularly via mucus cells (Morris et al., 2000b; Longshaw et al., 2002). The spore has characteristic soft shell valves, comprising eight cells surrounding four small subspherical polar capsules, and two sporoplasm cells, which may each contain a single secondary cell. The sporoplasm cells contain characteristic sporoplasmosomes, identical to those seen in the extrasporogonic histozoic fish stage (Fig. 12).

Fig. 12. Proliferative kidney disease, pathology and morphology of the causative agent Tetracapsuloides bryosalmonae. A. Rainbow trout (Oncorhynchus mykiss) fingerling exhibiting renal hypertrophy. B. Giemsa stained renal impression smear showing extrasporogonic stage of T. bryosalmonae surrounded by host phagocytes. C. Diagrammatic representations of stages in the development of histozoic T. bryosalmonae from rainbow trout kidney (primary cell nucleus, N; secondary cell, S; tertiary cell, T). D. Electron micrograph of a renal interstitial stage from rainbow trout kidney. Primary cell contains a prominent nucleus (N) with two secondary cells (S), one of which contains two tertiary cells (T). E. Fresh preparation of T. bryosalmonae from trout kidney with characteristic cytoplasmic granules and secondary cells within the primary cell, which is itself surrounded by phagocytic cells. F. Low-power view of a section of infected renal tissue showing loss of excretory elements and proliferation of interstitial haemopoietic tissue (upper left). G. Chronic granulomatous response in pike (E. lucius) spleen in response to T. bryosalmonae stages.

Most reports of PKD involve rainbow or steelhead trout (Oncorhynchus mykiss), although outbreaks in brown trout (S. trutta) and Atlantic salmon (S. salar) in Europe and in chinook salmon (Oncorhynchus tshawytscha) and coho salmon (Oncorhynchus kisutch) in North America are common (Hedrick et al., 1993). In Europe, Arctic charr (Salvelinus alpinus) and grayling (T. thymallus) are also known to be hosts for the parasite, and may exhibit signs of the disease (Feist and Bucke, 1993). Pike (E. lucius) is the only non-salmonid fish known to be susceptible (Seagrave et al., 1981; Morris et al., 2000a). Brook trout (Salmo fontinalis) can become infected but do not exhibit clinical signs of PKD (Feist and Bucke, 1993). Juvenile fish are most at risk from the disease but naïve fish of any age are susceptible. Frequently 100% of susceptible stocks become infected. Outbreaks of PKD are seasonal in nature, occurring in late spring, with clinical signs being seen typically between June and September when temperatures reach 15°C and above (Clifton-Hadley et al., 1986). Parasites are usually seen within renal tissue about 4 weeks post-exposure, with lesions developing shortly thereafter. The histozoic stage consists of a primary cell containing one or more secondary cells and occasional tertiary cells (Seagrave et al., 1980a, b). The cytoplasm of the primary cell contains numerous characteristic sporoplasmosomes, morphologically identical to those in the sporoplasm cell of the bryozoan stage (Feist, 1997; Kent et al., 1998). Pericyte formation has been observed (Feist and Bucke, 1987). The main presenting features of the disease are abdominal distension, bilateral exophthalmia and pale gills. Internally, marked renal and splenic hypertrophy is typical and the kidney may have a mottled appearance.

The extrasporogonic stage of the parasite proliferates rapidly within the host, reaching almost every organ via the bloodstream and eliciting focal or multifocal inflammation within the tissues. In the skeletal musculature, severe granulomatous myositis can result (Fernández-de-Luco et al., 1997). In the kidney and spleen, both the proliferation of the parasite and the host response are more vigorous. Interstitial haematopoietic cell hyperplasia, with degenerative changes in the excretory elements, and vascular pathology, including haemoglobin crystallization, occur (Clifton-Hadley et al., 1987; Fig. 12). In chronic infections, granulomatous interstitial lesions, with phagocytic cells surrounding the parasites, are commonly seen (MacConnell et al., 1989). Eventually the parasite is eliminated and the lesions resolve. Some parasites traverse the renal tubule epithelium, becoming coelozoic within the tubules, where sporogony is initiated. This generally occurs at a low level, with only a few, apparently immature, ovoid spores being produced. An exception is in Arctic charr, where large numbers of spores are formed without an extensive host reaction to the extrasporogonic stages (Kent et al., 2000). The spores are formed within monosporous pseudoplasmodia and measure 12 µm in length and 7 µm in width. They contain two subspherical polar capsules and are surrounded by indistinct valvogenic cells. Hedrick et al. (2004) described more advanced stages in the urine of fish recovering from PKD. These were subsperical with a width of 16 µm and a height of 14 µm, with two spherical polar capsules. Transmission studies have so far failed to produce infections of bryozoans by these stages (Tops et al., 2004).

Management strategies aimed at reducing exposure to the infective stage are the most effective means to control the disease. Reduced water temperature suppresses the effects of the disease (Clifton-Hadley et al., 1986) and fish exposed late in the season, before ambient temperatures fall, survive the infection and appear less susceptible to the disease the following year. The infective stage is present throughout the year (Gay et al., 2001; Tops and Okamura, 2003). Practical measures to reduce the numbers and distribution of the bryozoan hosts in and around aquaculture facilities have yet to be evaluated. Treatment regimes are covered in the section on 'Prevention and Control'.

Two species of Parvicapsula infecting salmonid renal tissue have been reported. Parvicapsula minibicornis, Kent, Whitaker and Dawe, 1997 is known from several Pacific salmonids from Canada (Jones et al., 2003, 2004). Affected fish exhibit slight renal swelling. Trophozoites reside within Bowman's space and capillaries of the glomeruli and occasionally within the renal tubule epithelium. Sporogonic stages occur within the lumen of renal tubules. Pyriform spores contain two equalsized pyriform polar capsules and measure 11 µm in length and 7.5 µm in width (Kent et al., 1997). The parasite has been associated with prespawn mortality among Fraser River stocks of sockeye salmon (Oncorhynchus nerka) in British Columbia (St-Hilaire et al., 2002). Prevalence rates amongst Pacific salmon species range from 47 to 100% (Jones et al., 2003). A second unidentified Parvicapsula species infects marine-cultured coho salmon in North America (Hoffmann, 1984). Other Pacific salmonids and Atlantic salmon are also susceptible. The disease has not been reported from Europe. Infection occurs 8 to 10 weeks post-introduction to an infected site, with up to 50% losses being reported (Johnstone, 1984). The kidney is the primary site of infection and appears hypertrophied and haemorrhagic. Renal tubules become occluded with trophozoites and developing spores (Fig. 13). Tubule epithelium may also harbour sporulating trophozoites. Histologically, the tubule epithelium becomes atrophied. The release of parasite stages to tubule epithelium and lumen of coho salmon (O. kisutch). the renal interstitium following rupture of affected tubules elicits only a mild host response. Glomerular infections have not been reported with this species. Pseudobranch infections, similar to those occurring in cultured Atlantic salmon infected with Parvicapsula pseudobranchiola have also been reported (Yasutake and Elliott, 2003). Relationships between Parvicapsula sp., P. pseudobranchiola and P. minibicornis require further analysis.

Fig. 13. Pathology of renal infections. A. Intracellular and coelozoic stages of Hoferellus carassii in the renal tubule of goldfish (Carassius auratus). B. Three-spined stickleback (G. aculeatus) kidney showing distension of renal tubules caused by numerous sporogonic stages of Myxobilatus gasterostei. C. Large xenoma of extrasporogonic stages of Myxidium lieberkuehni, which has replaced the glomerular tissue in the kidney of pike (E. lucius). D. Glomerular and tubule infections of Myxobilatus platessae in the kidney of European flounder (Platichthys flesus). E. Sporogonic stages of Myxidium minteri in the renal tubule lumen of chinook salmon (Oncorhynchus tshawytscha). F. Parvicapsula sp. sporogonic stages within the renal tubule epithelium and lumen of coho salmon (O. kisutch).

Marked renal hypertrophy associated with Parvicapsula renalis Landsberg, 1993 infections have been reported in red drum (Sciaenops ocellatus) from the Atlantic Ocean off Florida (Landsberg, 1993). The parasite infects proximal tubules of the posterior kidney with both extrasporogonic and sporogonic stages in the lumen of the tubules. Epithelial changes include vacuolation and the presence of hyaline droplets, sometimes with the presence of rodlet cells. Interstitial renal tissue is not affected.

M. rhodei is a common parasite of the roach (Rutilus rutilus) and other cyprinids in Europe (Dyková et al., 1987; Alvarez-Pellitero, 1989; Athanassapoulou and Sommerville, 1993a). Plasmodia are frequently found in Bowman's space within the glomerulus, resulting in atrophy of the glomerular tuft (Fig. 14). Development within the haemopoietic interstitial tissues results in an intense granulomatous inflammatory reaction, which is often effective in destroying the parasite. Mortalities have not been associated with M. rhodei infections (Dyková et al., 1987). Other species of Myxidium have been associated with significant disease. Myxidium minteri Yasutake and Wood, 1957 is restricted to wild and hatchery stocks of salmonids of the Pacific north-west of the USA. The parasite is coelozoic in renal tubules (Fig. 13), with occasional spores present in interstitial tissues in heavy infections (Yasutake and Wood, 1957). Spores are fusiform, 9.3 to 12.6 µm in length and 6 to 7 µm in width, with equal sized pyriform polar capsules. Coho salmon are most susceptible to infection (up to 44%), with prevalence rates in chinook salmon and rainbow trout generally less than 10% (Sanders and Fryer, 1970). However, in these cases, the parasite was located in the gall bladder, with rare cysts found in the liver. Myxidiosis caused by M. giardi in eels, Anguilla anguilla, affects many tissues, including the kidney, where significant damage to the excretory elements occurs. Granulomatous inflammation and mortality amongst elvers can result.

Fig. 14. Pathology of renal infections. A. Sphaerospora sp. in the kidney of dace (Leuciscus leuciscus) causing dilatation of renal tubule with reduction in tubule epithelial height. B. Atrophy of the glomerular tuft surrounded by a coelozoic plasmodium of Myxidium rhodei in the kidney of dace (Leuciscus leuciscus). C. Fibrous encapsulation of sporogonic plasmodia of M. rhodei in the renal interstitial tissue of R. rutilus. D. Intracellular extrasporogonic stage of an unidentified myxosporean in the renal tubule epithelium of dace (L. leuciscus).

In pike (E. lucius), extrasporogonic stages of Myxidium lieberkuehni Buetschli, 1882 invade the endothelial cells of the glomerular tuft, resulting in grossly enlarged cells containing numerous parasites and a hugely hypertrophied nucleus, the whole structure constituting a parasitic xenoma. The cells contained within the xenoma frequently contain secondary and tertiary cells, which, on release from the xenoma, pass down the proximal tubule and develop into small plasmodia (Lom et al., 1989a; Feist, 1997). Many infected glomeruli become infiltrated by host phagocytes and granulation tissue and are destroyed by the host. Large polysporic plasmodia are commonly found attached to the epithelium of the renal collecting ducts, and in the urinary bladder they impart an orange tinge due to the pigment granules contained within the plasmodia (Fig. 13). A second extrasporogonic stage is located intracellularly within the epithelial cells of the collecting ducts. Infected cells become hypertrophied and show degenerative changes without xenoma formation. It is presumed that these stages are also able to become sporogonic if they enter the lumen of the ducts. Affected epithelium becomes hyperplastic, forming papillomatous folds into the lumen of the duct, which itself becomes extremely dilated. Gross pathology in juvenile fish resulting from these changes is reminiscent of PKD, with renal hypertrophy and a greyish mottled appearance being typical features (Fig. 13). Mature spores of M. lieberkuehni are fusiform with longitudinal striations typical of the genus and measure approximately 20 µm in length and 6 µm in width.

Chloromyxum species also infect renal tissues, although most do not cause significant pathology. Chloromyxum majori Yasutake and Wood, 1957 in salmonids can result in the destruction of glomerular capillaries, and plasmodia of Chloromyxum inexpectatum Baska, 1990 in sterlet (Acipenser ruthenus) from the River Danube also infect glomeruli, displacing the glomerular tuft. Stages within the lumen of the tubules and urinary bladder have no discernible pathogenic effect (Baska, 1990).

Hoferellosis in goldfish (C. auratus) caused by H. carassii gives rise to the condition known as 'kidney enlargement disease' (KED) (Molnár et al., 1989). The disease has been reported from Europe, Asia and North America. Clinical signs include abdominal distension, protrusion of scales and loss of balance. The disease is seasonal in nature, with fish becoming infected during the late summer and autumn, developing clinical signs the following spring, when mortalities may result (Yokoyama et al., 1990a). Internally, marked renal hypertrophy is caused by a cystic papillomatous hyperplasia of the tubule epithelium (Fig. 13). Extrasporogonic stages of H. carassii invade tubule epithelial cells. The stages consist of primary cells up to 17 µm in diameter, with secondary, tertiary and even quaternary cells contained within them. Affected cells become elongate and protrude into the lumen of the tubule. Division of the epithelial cells, presumably in response to the parasite, offers further opportunities for intracellular development and attachment of coelozoic plasmodial stages. These are approximately 100 µm in diameter and contain spores measuring 13 µm in length and 8.4 µm in diameter. The oligochaete Nais cf. elinguis has been identified as the host for the actinospore stage (Aurantiactinomyxon) of H. carassii. However, transmission experiments have not been successful in inducing typical disease symptoms in susceptible fish (Trouillier et al., 1996).

Cultured gilthead sea bream (Sparus aurata) from the Mediterranean and Atlantic coast of Spain are susceptible to glomerular disease caused by Polysporoplasma sparis Sitjà-Bobadilla and Alvarez-Pellitero, 1995. Only semi-intensive culture systems are affected and the infection is not seasonal. Prevalence rates increase with increasing fish size, reaching over 50% in fish greater than 100 g (Palenzuela et al., 1999). Infected fish do not exhibit external clinical signs, but internally glomeruli of the mesonephros harbour numerous sporogonic stages of the parasite within the capillaries, replacing blood cells and reducing the filtration capacity of affected glomeruli. Hypertrophy of the corpuscle and thickening of the capsule are typically seen and, eventually, necrosis and fibrosis result. Parasites can also infect the renal tubule epithelium and occupy the lumen. In heavy infections, atrophy of renal interstitial tissue, with inflammation and haemorrhage, is present (Palenzuela et al., 1999). Spores are subspherical, 19.8 µm in length, 21.3 µm in thickness and 18.1 µm in width, with large capsulogenic cells. The characteristic feature of the spores, which differentiates the species from the genus Sphaerospora, is the presence of between 4 and 12 sporoplasm cells (Sitjà-Bobadilla and Alvarez-Pellitero, 1995). Gilthead sea bream and common dentex (Dentex dentex) are hosts to Leptotheca sparidarum Sitjà-Bobadilla and Alvarez-Pellitero, 2001. Large numbers of sporogonic stages within renal tubules cause blockage and pressure atrophy of the tubule epithelium (Sitjà-Bobadilla and Alvarez-Pellitero, 1995).

Several members of the genus Sphaerospora are pathogenic within the kidney of fish, although most frequently the changes produced are mild and induce only localized host reactions, especially where only the renal tubule lumens are infected (Fig. 14). Extrasporogonic stages appear to be typical for the genus and have been reported for many Sphaerospora species (Lom et al., 1985; Hedrick et al., 1988a, 1990; Feist et al., 1991; Supamattaya et al., 1993; McGeorge et al., 1994; Jones et al., 2004).

Sphaerosporosis in carp caused by S. renicola occurs amongst cultured common carp throughout Europe and in Israel. Prevalence may reach 100%. Sporogonic stages of the parasite are found in the lumen of renal tubules, comprising disporic pseudoplasmodia and subspherical spores measuring approximately 7.3 µm in diameter. These stages cause tubule dilatation and attenuation of the epithelium. In severe infections, the epithelium becomes atrophied and necrotic, with an associated inflammatory reaction in the interstitial tissues. Spores are infective to the oligochaete B. sowerbyi, with Neoactinomyxum actinospores being released after 98 days of development within the oligochaete host (Molnár et al., 1999a). However, experimental proof that these stages are infective to naïve carp is lacking. S. renicola has two proliferative extrasporogonic cycles. Blood stages, comprising primary cells containing a number of secondary and tertiary cells, invade the swim bladder, where they continue to proliferate, increase in size and induce swim bladder inflammation (see above) (Dyková et al., 1990). These swim bladder stages can be transmitted experimentally but have not been shown to give rise to the stages occurring within the renal tubule epithelium (Molnár and Kovács-Gayer, 1986). The marked cellular hypertrophy and stenosis of affected tubules are thought to be due to extrasporogonic stages of H. cyprini rather that S. renicola. Molnár (1988) reported that similar intracellular tubule epithelium stages occur in cyprinid fishes infected with Myxobilatus legeri (Cépede, 1905).

Mass mortality among cultured cobia (Rachycentron canadum) from Taiwan has been caused by infections with an unidentified Sphaerospora-like species (Chen et al., 2001). Cumulative mortality may reach 90% within 30 days of introduction to marine cages. Affected fish exhibit pale livers, ascites, gill pallor and marked renal hypertrophy, with the surface having a knobbly appearance and pale or haemorrhagic patches. Parasitic stages infect glomeruli, renal tubules and the interstitial tissues. Tubules become occluded with parasites and cellular debris, with the epithelium showing hypertrophy and hyperplastic changes. Ruptured tubules invoke a vigorous host response to the parasites, with granuloma formation being typical. Mature spores have not been described.

Wild and cultured groupers (Epinephelus malabaricus) from Thailand infected with Sphaerospora epinepheli Supamattaya, Fischer-Scherl, Hoffmann and Boonyaratpalin, 1991 exhibit highly vacuolated tubule epithelial cells with pycnotic nuclei. Other changes include dilation of glomerular capillaries, with degeneration of endothelial cells, thickening of the basement membrane and adhesion of the capillary membrane to the parietal layer of Bowman's capsule. Severely affected corpuscles become necrotic and surrounded by a fibrous layer (Supamattaya et al., 1993). Interstitial haematopoietic tissues are unaffected.

Serious outbreaks of histozoic sphaerosporosis involving significant losses among cultured juvenile tench (Tinca tinca) from Germany have been reported. Prevalence is generally low but can reach 100%. The causative agent, Sphaerospora tincae Plehn, 1925, produces large numbers of disporic pseudoplasmodia, replacing tissues of the pronephros and resulting in hypertrophy of the organ. Inflammation is usually absent. Concurrent infections with the coelozoic Sphaerospora galinae Evlanov, 1981 exert no discernible pathological effect (Lom et al., 1985).

Sphaerospora truttae Fischer-Scherl, El-Matbouli and Hoffmann, 1986 infects brown trout (S. trutta), grayling (Thymallus thymallus) and Atlantic salmon (S. salar). Hatcheryreared fish become infected during June, when extrasporogonic stages may be found in the blood and renal interstitial tissues. Primary cells contain up to 100 secondary cells, which in turn may have tertiary cells within them. These stages are transmissible to naïve fish by intraperitoneal injection. Coelozoic disporic pseudoplasmodia develop within renal tubules from August onwards, with mature spores present from September. These may be retained within the kidney for up to 18 months when held in fresh water but are lost within 4 months following transfer to sea water (McGeorge et al., 1996a, b).

Gall bladder and liver

A large number of coelozoic myxosporean species inhabit the gall bladder, especially in marine fish, and most infections are rather innocuous, without a significant host response. Infection proceeds via the hepatic vascular system into the biliary ductules and finally the gall bladder (Dyková and Lom, 1988b). Many myxosporeans cause changes to the colour and consistency of the bile, depending on the intensity of infection, and cellular necrosis of bladder epithelium is occasionally reported (Morrison et al., 1996). Plasmodia of pathogenic species frequently become lodged in the biliary collecting ducts in the liver, where they can be associated with proliferation of bile ducts, inflammation of the surrounding tissues (pericholangitis) and occasionally hepatocellular necrosis (Walliker, 1968; Bucher et al., 1992; Fig. 15). Attenuation of bile ductule epithelium in the liver is a common finding amongst gall-bladder-infecting species. It is likely that many myxosporeans inhabiting the biliary system are capable of inducing pathological changes similar to those described above. However, the majority of species have been described without histological examination of the bladder, bile duct or liver, which would reveal these.

Several Zschokkella species are known to be pathogenic. Zschokkella russeli Tripathi, 1948 in the five-bearded rockling (Ciliata mustela) are found at prevalences up to 89% at intertidal localities in Wales (Davies, 1985) and also occur in many other fish species. Plasmodia are found folded within the bile ducts in the liver, floating freely in the bile and loosely attached to the gall bladder epithelium. Pathological changes, including cholangiofibrosis, pericholangitis and flattened epithelium with reduced microvilli, have been reported in the hepatic bile ducts but the parasite induces no pathology in the gall bladder itself.

Zschokkella icterica Diamant and Paperna, 1992 infecting rabbitfish (Signanus luridus) from the Red Sea produces large plasmodia up to 400 µm in diameter in the hepatic bile ducts, whilst smaller plasmodia reside within the gall bladder, where they are associated with desquamation of the epithelial lining (Diamant and Paperna, 1992). Congestion of the ducts leads to cholestasis, breakdown of the ducts and, in severe infections, hepatocellular necrosis, and cholangitis results, with clinical signs of ascites and jaundice in affected fish. Mullets of the genera Mugil and Liza from the Mediterranean are hosts to Zschokkella mugilis Sitjà-Bobadilla and Alvarez-Pellitero, 1993. Sitjà-Bobadilla and Alvarez-Pellitero (1993) recorded prevalences up to 70%. Plasmodia measuring up to 50 µm in diameter induce degenerative changes in the bladder epithelium.

Fig. 15. Infections of the gall bladder, liver and neural tissues. A. Low-power view of papillomatous ingrowths of the gall-bladder epithelium of saithe (Pollachius virens) infected with Myxidium gadi. B. Semi-thin resin section from the previous specimen showing attachment of sporogonic M. gadi plasmodia. C. Hepatobiliary fibrosis associated with invasion of the bile ductules with elongate plasmodia of Myxidium truttae infecting brown trout (Salmo trutta). D. High-power view showing attenuation of gall-bladder epithelium and the presence of a small plasmodium of M. truttae within the hepatic parenchyma. E. Spores of Myxobolus cotti within the brain of bullhead (Cottus gobio).

Chloromyxum truttae Léger, 1906 is a common parasite of brown trout (S. trutta) and other salmonids (Alvarez-Pellitero et al., 1982; Lom and Dyková, 1992). Infections in wild fish do not usually exhibit clinical signs of disease, but heavy infections in cultured stocks may result in emaciation and yellowish discoloration of the skin and fins. A discoloured liver, swollen gall bladder and enteritis are the main presenting features, and mortalities due to severe infections have been reported (Schulman, 1966). Up to 100% prevalence of infections in cultured sea trout (S. trutta) (anadromous) and 48% in Atlantic salmon (S. salar) from Finland have been recorded without the presence of histopathological changes or mortalities (Feist and Rintamäki, 1994). Sporogonic plasmodia of Chloromyxum cristatum Léger, 1906 infecting common carp (C. carpio) and grass carp (Ctenopharyngodon idella) induce hepatocellular necrosis (Lom and Dyková, 1981). Similar pathology occurs in carp infected with Chloromyxum cyprini, where plasmodia invade the liver parenchyma. Regeneration of necrotic liver tissue in such cases is limited and the prognosis for infected fish poor (Dyková and Lom, 1988b). Necrosis of the gall bladder epithelium resulting from infections of Chloromyxum trijugum Kudo, 1919 has also been reported (Mitchell et al., 1980).

Myxidium gadi Georgèvitch, 1916 is a well-known parasite of gadoid fish such as saithe (Pollachius virens) and pollack (Pollachius pollachius), occurring at high prevalences in fish from the North Sea. Infected gall bladders are atrophied and have a pale coloration. Depending on the intensity of infection, the bile may be slightly discoloured and viscous, leading to complete occlusion of the gall bladder and bile duct with parasitic stages. Plasmodia attach themselves to the bile duct epithelium, resulting in extensive papillomatous ingrowths of the epithelium and necrosis (Feist and Bucke, 1992; Fig. 15). Intracellular epithelial and intrahepatic stages have not been reported.

Ceratomyxosis caused by Ceratomyxa sparusaurati Sitjà-Bobadilla, Palenzuela and Alvarez-Pellitero, 1995 is a significant pathogen of cultured gilthead sea bream (S. aurata) in the Mediterranean (Sitjà- Bobadilla et al., 1995). Prevalence of infection can reach 60% in infected stocks and low-level mortalities have been reported. Severe infections induce abdominal distension with inflammation and ascites. Cytological damage to the bladder epithelium includes hypertrophy and vacuolization. Sloughing of the epithelium and inflammation of the underlying tissues can occur. The parasite is regarded as a threat to sea bream culture (Palenzuela et al., 1997).

Infections of the intestine

Members of several genera of myxosporeans infect intestinal tissues, although relatively few are exclusive pathogens of the intestine. Most infections are not regarded as pathogenic, typically forming discrete sporogonic cysts with minimal localized host reaction to the parasites. Examples of pathogenic species are given below.

Ceratomyxosis of hatchery-reared and wild salmonids from the Pacific north-west of North America is caused by C. shasta (Ratliff, 1983; Bartholomew et al., 1989a; Hendrickson et al., 1989). The parasite is responsible for losses among hatchery stocks, juvenile wild salmonids and pre-spawning adults. Differences in susceptibility between salmonid species and strains have been noted (Ibarra et al., 1991, 1992), and fish surviving the infection are typically undersized and emaciated (Tipping, 1988). Susceptible fish acquire the infection from actinospores released from the freshwater polychaete M. speciosa (Bartholomew et al., 1997). External clinical signs vary according to the salmonid species and include anorexia, lethargy, abdominal distension, darkened coloration and exophthalmia. Internally, there is widespread enteritis and haemorrhaging, with the formation of mucoid deposits and caseousmaterial within the intestine and pyloric caecae, resulting from destruction of the intestinal mucosa. As the infection proceeds, proliferative stages spread via the bloodstream and infect the visceral tissues and skeletal muscle by causing widespread haemorrhaging and necrosis (Bartholomew et al., 1989b).

The histozoic nature of C. shasta is unique amongst the usually benign coelozoic members of the genus inhabiting the urinary bladder and gall bladder of marine fish. The actinospore stage released from M. speciosa invades the epithelium of the posterior intestine, with small plasmodia initiating local lymphocytic infiltration by 18 days post-exposure at 12°C. Between 30 and 50 days post-exposure and the infective stage, all intestinal layers become invaded with the presporogonic stages of the parasite, necrosis is evident and mortalities ensue (Fig. 16). At 21°C the pathogenesis of the disease is more rapid. Sporogenesis is complete by day 18 and mortalities occur by day 20 (Bartholomew et al., 1989b). Gross lesions may also affect the spleen and renal and muscle tissue. Myxospores are arcuate, with a diameter of 14-23 µm, containing two uninucleate sporoplasm cells with a subspherical polar capsule positioned each side of the sutural line.

A few members of the genus Myxobolus infect intestinal as well as other tissues. Most species are located in the tissues of the lamina propria underlying the mucosal epithelium, suggesting that plasmodial development is initiated within capillaries (Molnár, 2002d). Rupture of the cysts is likely to result in the death of the host. Plasmodia vary greatly in size, depending on the species, from microscopic to over 5 mm in diameter. Myxobolus nodulointestinalis Masoumian, Baska and Molnár, 1996 produces large cysts in the intestinal wall of Barbus sharpeyi from south-west Iran, constricting the gut lumen. Thelohanellus kitauei (Egusa and Nakajima, 1981) causes serious disease in Israel carp (C. carpio nudus) in Korea. The parasite forms large cysts on the intestinal mucosa, surrounded by a thin connective tissue capsule. Development of the cysts is rapid during July, with maturation leading to cyst rupture and release of spores occurring in August and September (Rhee et al., 1993). Infections of catfish (Clarias gariepinus) from the River Nile, Egypt, with Henneguya suprabranchiae Landsberg, 1987 give rise to large intestinal cysts measuring up to 4.5 mm in diameter (El-Mansy, 2002). Prevalence was 21.2%. Plasmodia invade the outer layer of the intestine and cause pressure atrophy to the smooth muscle layer, which becomes discontinuous around the intestine.

Fig. 16. Infections of the intestine and associated tissues. A. Destruction of the intestinal mucosa of gilthead seabream (Sparus aurata) caused by Enteromyxum leei. B. Intraepithelial stages of E. leei in the intestinal mucosa of S. aurata. C. Section of pyloric caecae with large numbers of plasmodia and spores of Ceratomyxa shasta (arrows) in the underlying connective tissue. Inset: developing spores of C. Shasta. D. Intraepithelial stages of C. shasta in the mucosal epithelium and submucosal tissues (arrow). E and F. Cysts of Myxobolus sp. in the thin connective-tissue layer underlying the intestinal epithelium of minnow (Phoxinus phoxinus) and roach (R. rutilus), respectively.

Although Kudoa species are well-known parasites of striated muscle (see above), some species invade other tissues, including the smooth muscle of the intestinal wall. Kudoa dianae Dyková, Fajer Avila and Fiala, 2002, a parasite of the bullseye puffer, Sphoeroides annulatus, was found in 20% of adults collected off the coast of Mexico. Spores measure 4.5-5.5 µm in width and 5.5-6.5 µm in thickness. Cysts were also located in mesenteric tissues. Although there is no significant inflammatory reaction to intact cysts, spores from ruptured plasmodia are phagocytosed by macrophages and transported to the mucosal epithelium, where they can totally replace the epithelial tissue (Dyková et al., 2002). Kudoa intestinalis Maeno, Nagasawa and Sorimachi, 1993 infecting striped mullet, M. cephalus, from Japan also infects the smooth muscle of the intestine but is apparently non-pathogenic.

E. scophthalmi is also a highly pathogenic histozoic parasite which causes the disease enteromyxosis, characterized by severe enteritis and mortalites in cultured turbot (Scophthalmus maximus) (Branson et al., 1999; Palenzuela et al., 2002). Transmission is direct, via cohabitation with infected fish, exposure to water contaminated with spores and oral administration of infected intestinal tissue. Early developmental stages are intracellular within epithelial cells and can be detected in the anterior intestine as early as day 8 following oral intubation. Proliferative stages become abundant intraepithelially and within the lumen of the intestine (Redondo et al., 2002, 2003). Pathogenesis of the disease is dependent on temperature and mode of infection. Typical pathology includes vacuolation and sloughing of the epithelial cells and oedema of the subepithelial tissues. Mortalities occur within 3 weeks of exposure to infected material and reach 100% in most experimental trials. Spores (22.2 µm in length and 11.7 µm in width) are scarce in infected animals, are not formed until late in the infections and do not appear to be responsible for transmission to susceptible hosts (Redondo et al., 2002).

E. leei was first noted as a pathogen of cultured gilthead sea bream (S. aurata) by Diamant (1992) and subsequently in several other cultured marine species in the Mediterranean area (Diamant et al., 1994; Le Breton andMarques, 1995; Diamant, 1998). Direct fish-to-fish transmission occurs, presumably by ingestion of exfoliated intestinal tissues infected with spores or presporogonic stages of the parasite, or by coprophagy. Experimental infections in S. aurata resulted in prevalence levels of up to 33% and the onset of mortalities within 2 weeks of exposure by cohabitation or exposure to effluent from a tank containing infected fish (Diamant, 1997). Pathological changes are similar to those in enteromyxosis caused by E. scophthalmi (Fig. 16). Both species are serious threats to the mariculture of several fish species in the Mediterranean region.

Emaciation disease of tiger puffer (Takifugu rubripes) in the Kyushu district of Japan has been noted since 1996 (Tun et al., 2000). Clinical signs are principally general emaciation of the body, including enophthalmia and the appearance of bony ridges on the head. The disease can result in mortalities of up to 60%. Three myxozoans have been detected in the intestine of infected fish (namely M. fugu, Myxidium sp. TP and Leptotheca fugu Tun, Yokoyama, Ogawa and Wakabayashi, 2000) (Ogawa and Yokoyama, 2001). Of these, Myxidium sp. TP and M. fugu can be transmitted by feeding infected tissues, cohabitation with infected fish and exposure to effluent from tanks containing infected fish (Yasuda et al., 2002). Histopathological changes include infiltration with inflammatory cells into the lamina propria and epithelium, with eventual detachment of the mucosal layer as infection with Myxidium sp. TP and L. fugu progresses. Myxidium fugu does not appear to be pathogenic (Tun et al., 2002). The tissue tropism, morphological characteristics and pathogenicity of M. fugu appear similar to those of Enteromyxum infections, and further studies are required to clarify relationships between these parasites.

Infections of the swim bladder

There are several cyst-forming myxosporean parasites that infect the wall of the swim bladder. In most cases, there is little significant host reaction to the parasite since sporogonic stages are not exposed to the host immune system. However, as is the case for such infections in other locations within the host, rupture of cysts results in a vigorous inflammatory response. In the case of swim bladder infections, the likely result is organ dysfunction, impaired swimming performance and possibly death. However, such an outcome has rarely been reported. In chub (L. cephalus) infected with Myxobolus cycloides Gurley, 1893, infiltration of trophozoites with fibroblasts results in the isolation of small clusters of spores by fibrous tissue and macrophages. Eventually there is complete encapsulation of the trophozoite and necrosis of the trapped spores. M. cycloides is not regarded as a serious pathogen (Holzer and Schachner, 2002).

The swim bladder is one site for the extrasporogonic proliferative stage of S. renicola in common carp fingerlings, which causes swim bladder inflammation (SBI). The stages comprise primary cells containing up to approximately 50 secondary cells, each containing one or two tertiary cells. Rupture of the primary cell releases the secondary cells, which may then grow, with internal division of the enclosed cells to repeat the cycle. The presence of large numbers of parasites in the swim bladder wall elicits a strong inflammatory response. Hyperplasia of the connective and epithelial tissues results in extreme thickening of the swim bladder wall, and haemorrhaging is frequently present. Affected fish display reduced swimming capability and poor growth. In the acute phase, peritonitis and renal hypertrophy can occur. Prognosis for fish with SBI is poor and mortalities may occur (Körting, 1982; Csaba et al., 1984; Dyková and Lom, 1988a; Molnár, 1988).

Diagnosis

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Diagnosis of myxozoans is still mainly reliant on morphological characteristics of mature spore stages, using established criteria (Lom and Arthur, 1989; Lom et al., 1997). Detection of developmental stages from fresh material, smear preparations and histological sections is dependent on the experience of the investigator and the intensity of the infection. However, specific identification is generally not possible. Ultrastructural investigations using sectioned material are rarely able to identify species, but scanning electron microscopy provides valuable diagnostic data on the surface morphology of spore stages (Lom and Dyková, 1993). Fresh or fixed spores are best visualized using phase contrast or differential interference contrast microscopy, whilst smears or imprints can be stained with Giemsa, May-Grünwald-Giemsa or silver nitrate (Wolf and Markiw, 1979; Clifton-Hadley et al., 1983; Baska and Molnár, 1988). Biotinylated lectins have been used to characterize the carbohydrate types within myxozoans and have some diagnostic value (Castagnaro et al., 1991; Marín de Mateo et al., 1996; Muñoz et al., 1999a, 2000), and the fluorescent dye 5(6)-carboxyfluorescein diacetate succinimidyl-ester (CFSE) has been used to determine routes of entry by actinospores into the fish host (Yokoyama and Urawa, 1997).

Monoclonal and polyclonal antibodies have been developed for a number of myxozoans that are important fish pathogens and used to detect both extrasporogonic and spore stages and to study parasite antigens. Antibodies have been raised against Tetracapsuloides bryosalmonae (Adams et al., 1992; Marín de Mateo et al., 1996; Saulnier and de Kinkelin, 1996; Morris et al., 1997, 2000a), Ceratomyxa shasta (Bartholomew et al., 1989c), Kudoa thyrsites (Chase et al., 2001), Henneguya salminicola (Clouthier et al., 1997), Myxobolus cerebralis (Griffin and Davis, 1978; Markiw and Wolf, 1978; Hamilton and Canning, 1988), Sphaerospora dicentrarchi Sitja-Bobadilla and Alvarez-Pellitero, 1992 (Muñoz et al., 1998, 1999b) and M. rotundus (Lu et al., 2003).

The advent of molecular biology-based tools has led to the development of sensitive methods for the detection of myxozoan parasites of economic concern by targeting specific rDNA sequences, in particular, the use of PCR to link the alternate stages of myxozoan life cycles (Andree et al., 1997; Bartholomew et al., 1997; Lin et al., 1999; Pote et al., 2000; Feist et al., 2001; Hanson et al., 2001). Primers have been developed that target both the 18S and the ITS regions of the rDNA (Andree et al., 1999a) and include generic myxozoan primers (Kent et al., 2001), genus-specific (Andree et al., 1999b) and species-specific primers (Hervio et al., 1997; Saulnier and de Kinkelin, 1997; Andree et al., 1998; Kent et al., 1998; Morris et al., 2002a). DNA sequence data from the PCR product provide confirmation on species identity and in addition are routinely used for phylogenetic analysis. In addition, restriction fragment length polymorphism (RFLP) on PCR products can be used to discriminate species and has been used to demonstrate phylogenetic links between species (Xiao and Desser, 2000c; Eszterbauer et al., 2001; Eszterbauer, 2002).

One of the major constraints of the PCR technique is that, whilst it demonstrates the presence of parasite DNA, it does not provide information on the viability or pathological effect of the parasite. Therefore, a further refinement of the PCR method has been in the development of the in situ hybridization technique to demonstrate the presence of myxozoans in tissue sections (Antonio et al., 1998; Frasca et al., 1999; Morris et al., 1999, 2000b; Longshaw et al., 2002).

Epidemiology

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Myxozoans can show seasonal and annual variations in prevalence and this is related to a number of factors, including availability of hosts and ambient temperature influencing parasite development. Whilst the definitive hosts (oligochaetes) can release actinospore stages throughout the year, most are released in spring and summer in temperate regions (El-Mansy et al., 1998a, b; Xiao and Desser, 1998b; Özer et al., 2002; Oumouna et al., 2003). Similarly, Tetracapsuloides bryosalmonae is present within the bryozoan definitive host throughout the year, but is released mainly in spring and summer (Gay et al., 2001; Tops and Okamura, 2003). A corresponding seasonal pattern of infection in the fish host occurs, though in many cases the pattern is less clear. The confusing patterns observed may in part be due to the examination of different host age classes within a sample, as it is known that prevalence of infection alters with age (Roubal, 1994). However, in temperate climates, it appears that prevalences are generally highest in the autumn and winter (Andrews, 1979; Gonzalez-Lanza and Alvarez-Pellitero, 1985; Mitchell, 1989; Brummer-Korvenkontio et al., 1991). Furthermore, inter-annual variations in prevalences within fish hosts are known to occur (Awakura et al., 1995; Molnár, 1998; Molnár and Székely, 1999; Pampoulie et al., 2001; M. Longshaw and S.W. Feist, unpublished data).

Extrasporogonic stages of T. bryosalmonae infections in fish tend to be short-lived, less than a year. Fish surviving the infection become resistant to reinfection and generally do not display disease symptoms. Myxospore infections in fish hosts, however, can be extremely long-lasting, potentially persisting for the life-time of the host. Infections in oligochaetes can persist for at least a year under laboratory conditions and possibly for the life-time of the oligochaete under natural conditions (El-Matbouli et al., 1999a, b; Granath and Gilbert, 2002).

Ecological factors affecting transmission

Actinospore release follows a circadian pattern in oligochaetes, with most spores being released during late evening and early morning (Yokoyama et al., 1993a; Özer and Wootten, 2001). The maximum number of spores released daily by an individual worm can be as many as 80,000 for Echinactinomyxon types (Özer and Wootten, 2001). Actinospores are released from the oligochaete host throughout the year; however, most are released during the summer (El-Mansy et al., 1998a, b; Özer et al., 2002). There have been limited studies on the longevity of actinospores following release from the oligochaete, with estimates ranging from 11 to 25 days (Yokoyama et al., 1993a; Xiao and Desser, 2000a). Actinospores appear to release their sporoplasms in response to mucus from the specific fish host (Yokoyama et al., 1993a, 1995a).

The general paucity of data on actinospore biology for most species contrasts markedly with those gained on the actinospore of Myxobolus cerebralis. The susceptibility of different genetic strains of Tubifex tubifex to M. cerebralis myxospores and the factors that influence the distribution of actinospore infections in oligochaetes have been determined (Allen and Bergersen, 2002; Beauchamp et al., 2002). The M. cerebralis actinospore develops optimally at 15°C, and developmental stages degenerate at temperatures above 25°C (El-Matbouli et al., 1999a). The effect of different chemicals and salinity on the viability of the Triactinomyxon stages has also been investigated (Wagner et al., 2003). Whilst it is recognized that sediment type affects the distribution of oligochaete species (Xiao and Desser, 1998b), Blazer et al. (2003) showed that the substrate type affected the number and duration of release of M. cerebralis actinospores. Actinospore release was greatest in oligochaetes maintained in mud and sand, and least in a leaf litter substrate (Blazer et al., 2003). Stevens et al. (2001) found that the initial myxospore dose affected the number of actinospores produced and that the parasite reduced the biomass, abundance and individual weights of oligochaetes. There is a need to conduct more wide-ranging studies on actinospore biology and ecology in order to ascertain whether the patterns that are apparent in M. cerebralis actinospores apply to other actinosporean infections.


Vector specificity and factors affecting vector



Most actinospores have been recorded in oligochaetes, predominantly in the families Tubificidae and Nadidae, although there are a few records in polychaetes (Ikeda, 1912; Bartholomew et al., 1997; Hallett et al., 1998; Køie, 2000, 2002; Køie et al., 2004). The life cycle for Ceratomyxa shasta is the only one that has been completed, in which a freshwater polychaete has been implicated. Whilst Bartholomew et al. (1997) also reported the presence of an Aurantiactinomyxon type in the same host, thus far all other records of actinospores in marine polychaetes are of the Tetractinomyxon type (Ikeda, 1912; Køie, 2000, 2002). Despite a number of publications on the actinospore fauna of marine oligochaetes and polychaetes, no links have been made between the stages in this host and in a vertebrate counterpart (Caullery and Mesnil, 1905; Hallett et al., 1995, 1997, 1998, 1999, 2001; Hallett and Lester, 1999).

Parasite specificity in the invertebrate host is unclear. Actinospores of M. cerebralis appear to be specific to genetic lineages of Tubifex tubifex, whilst other actinospores may have a wide variety of oligochaete types as hosts (Xiao and Desser 1998a, b; Koprivnikar and Desser, 2002; Székely et al., 2003). Whilst the malacosporean Tetracapsuloides bryosalmonae appears to be relatively specific to the salmonid host (with the exception of pike, Esox lucius), it has been reported from a wide range of bryozoan hosts in both Europe and the USA (Longshaw et al., 1999; Okamura et al., 2001; Okamura and Wood, 2002).

As with any free-living organism, polychaetes, oligochaetes and bryozoans are affected by the environment in which they are found. Tubifex tubifex distributions are determined by the composition and organic content in the substratum and the presence of other oligochaetes (Granath and Gilbert, 2002) and, whilst they are often associated with sites of poor water quality, they can survive in many different habitat types. In general, poor water quality, low oxygen content and low flow rates are detrimental to the definitive hosts of myxozoans.

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