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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
International Common Names
- English: chlamydial infections of fish; epitheliocystis disease
Pathogen/sTop of page Chlamydia-like organisms
OverviewTop of page
The causative agent(s) of the disease referred to as EP are morphologically diverse and may represent a group of related organisms that produce similar pathology in varied hosts. The taxonomic placement of these diverse intracellular microorganisms is undefined but is considered to fit most appropriately in the order Chlamydiales (Moulder, 1984). Most exhibit the pleomorphic developmental cycle typical of this group of intracellular bacteria (Paperna et al., 1981).
Epitheliocystis occurs as a non-proliferative or proliferative disease, characterized by cysts in the branchial epithelia of the host. Clinical signs may include lethargy, flared opercula and rapid respiration. Cysts may appear as transparent white to yellow capsules on the gill filaments. Generally the host response is limited, and there is little or no mortality associated with infection.
[Based upon material originally published in Woo PTK, Bruno DW, eds., 1999. Fish diseases and disorders, Vol. 3 Viral, bacterial and fungal infections. Wallingford, UK: CABI Publishing.]
Host AnimalsTop of page
|Animal name||Context||Life stage||System|
|Acanthopagrus schlegelii (black porgy)|
|Acipenser gueldenstaedtii (russian sturgeon)|
|Acipenser transmontanus (white sturgeon)|
|Ameiurus nebulosus (brown bullhead)|
|Anoplopoma fimbria (sablefish)|
|Bidyanus bidyanus (silver perch)|
|Cyprinus carpio (common carp)|
|Dentex dentex (common dentex)|
|Dicentrarchus labrax (European seabass)|
|Gadus morhua (Atlantic cod)|
|Ictalurus punctatus (channel catfish)|
|Lates calcarifer (barramundi)||Domesticated host||Aquatic: Fry||Enclosed systems/Cages|
|Lepomis macrochirus (bluegill)|
|Limanda ferruginea (yellowtail flounder)||Aquatic: Adult|
|Liza aurata (golden grey mullet)|
|Lutjanus kasmira (blueline snapper)|
|Morone americana (white perch)|
|Morone saxatilis (striped sea-bass)|
|Mugil cephalus (flathead mullet)|
|Oncorhynchus gorbuscha (pink salmon)|
|Oncorhynchus keta (chum salmon)|
|Oncorhynchus kisutch (coho salmon)|
|Oncorhynchus mykiss (rainbow trout)|
|Oncorhynchus tshawytscha (chinook salmon)|
|Oreochromis aureus (blue tilapia)|
|Oreochromis mossambicus (Mozambique tilapia)|
|Oreochromis niloticus (Nile tilapia)|
|Pagrus major (red seabream)|
|Paralichthys dentatus (summer flounder)|
|Piaractus mesopotamicus (small-scales pacu)|
|Pseudopleuronectes americanus (winter flounder)|
|Salmo salar (Atlantic salmon)||Domesticated host, Wild host||Aquatic: Adult||Enclosed systems/Cages|
|Salvelinus namaycush (lake trout)|
|Seriola dumerili (greater amberjack)|
|Seriola lalandi (yellowtail amberjack)|
|Sparus aurata (gilthead seabream)|
|Takifugu rubripes (Japanese puffer fish)||Wild host|
|Zoarces americanus (ocean pout)|
Hosts/Species AffectedTop of page
Epitheliocystis was first described in the bluegill sunfish (Lepomis macrochirus), by Hoffman et al. (1969), who identified the aetiological agent as a bedsonia (now termed chlamydia). Molnar and Boros (1981) verified the chlamydia-like nature of the causative agent and determined that EP was identical to the mucophilosis disease of common carp described by Plehn (1920) and believed to be caused by a unicellular alga or fungus. These descriptions were followed by reports of the disease in freshwater, marine and anadromous fish from both warm and cold water environments. The documented host range includes species from more than 20 families of fishes. Probably additional hosts have gone unreported because of the self-limiting nature of the disease.
Distribution TableTop of page
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/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Atlantic, Northeast||Present||Paperna et al., 1981; Paperna and Alves, 1984; Nylund et al., 1998|
|Atlantic, Northwest||Present||Zachary and Paperna, 1977; Morrison and Shum, 1983; Lewis et al., 1992|
|Atlantic, Southeast||Present||Paperna and Sabnai, 1980; Paperna et al., 1981|
|Indian Ocean, Eastern||Present||Langdon et al., 1991; Anderson and Prior, 1992|
|Mediterranean and Black Sea||Present||Paperna, 1977; Paperna and Laurencin, 1979; Paperna and Sabnai, 1980; Paperna et al., 1981; Crespo et al., 1990; Grau and Crespo, 1991; Montero et al., 2004|
|Pacific, Eastern Central||Present||Venizelos and Beneti, 1996|
|Pacific, Northeast||Present||Groff et al., 1996; Kent et al., 1998|
|Pacific, Northwest||Present||Miyazaki et al., 1986; Egusa, 1987; Egusa et al., 1987; Kim et al., 2000|
|Pacific, Southeast||Present||Carvajal et al., 1990; Nowak and Clark, 1999|
|Pacific, Western Central||Present||Work et al., 2003|
|China||Present||Weng and Chen, 2000|
|Israel||Present||Paperna et al., 1978; Paperna and Sabnai, 1980; Paperna et al., 1981; Paperna and Alves, 1984|
|Japan||Present||Miyazaki et al., 1986; Egusa, 1987; Egusa et al., 1987|
|South Africa||Present||Paperna and Sabnai, 1980; Paperna et al., 1981|
|Canada||Present||Desser et al., 1988|
|USA||Present||Paperna and Zwerner, 1976; Bradley et al., 1988|
|-California||Present||Groff et al., 1996|
|-Connecticut||Present||Wolke et al., 1970|
|-Idaho||Present||Rourke et al., 1984|
|-Oklahoma||Present||Zimmer et al., 1984|
|-Oregon||Present||Lannan et al., 1999|
|-West Virginia||Present||Hoffman et al., 1969|
|Brazil||Present||Szakolczai et al., 1999|
|-Rio de Janeiro||Present||Lima et al., 2001|
|Chile||Present||Carvajal et al., 1990|
|Ecuador||Present||Venizelos and Beneti, 1996|
|France||Present||Paperna and Laurencin, 1979; Paperna et al., 1981|
|Portugal||Present||Cruz et al., 2000|
|Russian Federation||Present||Voronin and Chernysheva, 1997|
|UK||Present||Longshaw et al., 2004|
|Australia||Present||Langdon et al., 1991; Anderson and Prior, 1992|
|-Tasmania||Present||Frances et al., 1997|
DiagnosisTop of page
Epitheliocystis occurs as a non-proliferative or proliferative disease, characterized by cysts in the branchial epithelia of the host. Clinical signs may include lethargy, flared opercula, increased mucus production and rapid respiration. Cysts may appear as transparent white to yellow capsules on the gill filaments. Generally the host response is limited, and there is little or no mortality associated with infection.
Characteristic cysts are hypertrophic host cells filled with the causative bacterium. The enlarged host cells range from 10 to 400 mm in diameter and are frequently surrounded by squamous or cuboidal epithelial cells. The nature and origin of the target host cell is unclear; cysts may originate from a single cell (Zachary and Paperna, 1977) or be a coalescence of several (Wolke et al., 1970). The target cell may vary, as infection has been variously described in mucus, epithelial lining and chloride cells of an unidentified species of carp (Paperna and Alves de Matos, 1984); in capillary-forming cells of red sea bream (Pagrus major) and tiger puffer (Takifugu rubripes; Miyazaki et al., 1986); and in cells described as possible transformed macrophages in brown bullhead (Ictalurus nebulosus; Desser et al., 1988).
Preliminary diagnosis of EP is made by observation of the white to yellow cysts on the gills or skin of affected fish (Wolf, 1988). The thick capsule and granular contents, which are characteristic of EP cysts, are easily seen in wet mounts. The gill is the most frequently affected organ, but it has been suggested that the pseudobranch also be examined for cysts (Crespo et al., 1990).
No serological techniques are available for identification of the organism or for diagnosis of infection. Electron microscopy is required for definitive diagnosis of this disease. With this technique, the intracellular forms, too small to be clearly differentiated under the light microscope, can be observed and differentiated from cysts having a viral aetiology (Wolf, 1988).
List of Symptoms/SignsTop of page
|Finfish / Air gulping - Behavioural Signs||Aquatic:Adult||Sign|
|Finfish / Cysts - Gills||Aquatic:Adult||Diagnosis|
|Finfish / Fish swimming near surface - Behavioural Signs||Aquatic:Adult||Sign|
|Finfish / Generalised lethargy - Behavioural Signs||Aquatic:Adult||Sign|
|Finfish / Mucus build up - Gills||Aquatic:Adult||Sign|
Disease CourseTop of page
Molnar and Boros (1981) described the development of EP cysts in the gills of the common carp (Cyprinus carpio). In the early stages of the disease, infected cells are 10-15 mm in diameter and contain a central inclusion body, foamy cytoplasm and excentric nucleus. As infection progresses, infected cells increase in size to 70-80 mm in diameter and the granular inclusion fills the entire host cell, compressing or replacing both the nucleus and the cytoplasm.
In non-proliferative infections, cysts may be surrounded by a layer of squamous or cuboidal epithelial cells. Typically, no host response is apparent, even in the presence of large numbers of cysts. However, in certain cases, the organism induces an extensive host response, with unrestricted proliferation of gill epithelia and extensive mucus production, a condition referred to as hyperinfection (Paperna, 1977; Rourke et al., 1984; Bradley et al., 1988). The proliferative form of the disease has been described in Sparus aurata (Paperna, 1977). Hyperplastic epithelial cells form concentric layers around the cysts and proliferate throughout the gill filaments. Infiltrating macrophages and eosinophils combine with the hyperplastic tissue to surround and obstruct the capillary network of the gill filament. These circumstances impair both gas transfer and osmoregulatory processes, which may result in the death of infected fish.
The causes for variation in the severity of EP infections are not clear, but proliferative infections with accompanying mortality often occur in cultured fish (Paperna, 1977; Miyazaki et al., 1986; Bradley et al., 1988; Crespo et al., 1990; Groff et al., 1996; Crespo et al., 1999; Montero et al., 2004). Non-proliferative infections are more frequently found in free-ranging fish (Grau and Crespo, 1991). Progression from the chronic to the progressive form may occur when host defence mechanisms are compromised by genetics or environmental factors (Paperna and Sabnai, 1980). Because the causative organism has not been isolated, the possibility that these observations can be explained by differences in virulence of the pathogen cannot be excluded.
EpidemiologyTop of page
Natural transmission of the chlamydia-like organisms (CLO) is not understood, but horizontal transmission apparently occurs within some host species. Hoffman et al. (1969) described experiments in the bluegill and Wolf (1988) listed experiments conducted by D.S. Wyand in goldfish (Carassius auratus) that demonstrated direct transmission. In both studies EP developed in the experimental animals 3-4 weeks after the addition of infected gill tissue to aquaria where uninfected fish were held. Paperna (1977) suggested that transmission from infected fish may occur in culture facilities through contaminated nets and other equipment. There is little information on interspecies transmission and it is not known if differences in the morphology of the agent relate to different host specificities or the CLO itself.
No information is available concerning temperature requirements for replication of the CLO or on transfer of the CLO from fish to humans or other warm-blooded animals. However, these microorganisms have only been reported in fish.
Impact SummaryTop of page
|Fisheries / aquaculture||Negative|
Zoonoses and Food SafetyTop of page
This species is not a zoonoses.
ReferencesTop of page
Carvajal J; Ruiz G; Gonzales L, 1990. Gill histopathologies in coho salmon (Oncorhynchus kisutch) and Atlantic salmon (Salmo salar) raised under net cages in southern Chile. Medio-Ambiente, 11:53-58.
Crespo S; Zarza C; Padrós F; Marín de Mateo M, 1999. Epitheliocystis agents in sea bream Sparus aurata [Pagrus aurata]: morphological evidence for two distinct chlamydia-like developmental cycles. Diseases of Aquatic Organisms, 37(1):61-72.
Cruz e Silva MP; Orge ML; Afonso-Roque MM; Grazina-Freitas MS; Carvalho-Varela M, 2000. Diplectanum aequans (Wagener, 1857) Diesing, 1858 (Monogenea, Diplectanidae) in sea bass (Dicentrarchus labrax (L.), 1758) from freshwater culture. Acta Parasitológica Portuguesa, 7(1/2):53-56.
Desser S; Paterson W; Steinhagen D, 1988. Ultrastructural observations on the causative agent of epitheliocystis in the brown bullhead , Ictalurus nebulosus Lesueur, from Ontario and a comparison with the chlamydiae of higher vertebrates. Journal of Fish Diseases, 11(6):453-460.
Draghi A; Popov VL; Kahl MM; Stanton JB; Brown CC; Tsongalis GJ; West AB; Frasca; S, 2004. Characterization of Candidatus Piscichlamydia salmonis (Order Chlamydiales), a Chlamydia-Like Bacterium Associated With Epitheliocystis in Farmed Atlantic Salmon (Salmo salar). Journal of Clinical Microbiology, 42:5286-5297.
Egusa S, 1987. Epitheliocystis disease. Fish Pathology, 22:165-171.
Egusa S; Miyazaki T; Shiomitsu T; Fujita S, 1987. Epitheliocystis-like disease among hatchery-reared fry of Oplegnathus punctatus. Fish Pathology, 22(1):33-34.
Groff JM; LaPatra SE; Munn RJ; Anderson ML; Osburn BI, 1996. Epitheliocystis infection in cultured white sturgeon (Acipenser transmontanus): antigenic and ultrastructural similarities of the causative agent to the chlamydiae. Journal of Veterinary Diagnostic Investigation, 8(2):172-180.
Hoffman GL; Dunbar CE; Wolf K; Zwillenberg LO, 1969. Epitheliocystis, a new infectious disease of the bluegill (Lepomis macrochirus). Antonie Van Leeuwenhoek Journal of Microbiology and Serology, 35:146-158.
Kent ML; Traxler GS; Kieser D; Richards J; Dawe SC; Shaw RW; Prosperi-Porta G; Ketcheson J; Evelyn TPT, 1998. Survey of salmonid pathogens in ocean-caught fishes in British Columbia, Canada. Journal of Aquatic Animal Health, 10(2):211-219.
Langdon JS; Elliott K; Mackay B, 1991. Epitheliocystis in the leafy sea-dragon. Australian Veterinary Journal, 68(7):244.
Lannan CN; Bartholomew JL; Fryer JL, 1999. Rickettsial and chlamydial infections. Fish diseases and disorders. Volume 3: viral, bacterial and fungal infections., 245-267; 5 pp. of ref.
Lima FCde; Machado APG; Borges AP; Lima CHA; Andrade Cde M; Mesquita Ede FMde, 2001. Epitheliocystis disease in Tilapia nilotica (Linnaeus, 1758) from Rio de Janeiro state, Brazil. Ciência Rural, 31(3):519-520.
Longshaw M; Green MJ; Feist SW, 2004. Histopathology of parasitic infections in greater pipefish, Syngnathus acus L., from an estuary in the UK. Journal of Fish Diseases, 27:245-248.
Miyazaki T; Fujimaki Y; Hatai K, 1986. A light and electron microscopic study on epitheliocystis disease in cultured fishes. Bulletin of the Japanese Society of Scientific Fisheries, 52:199-202.
Montero FE; Crespo S; Padrós F; Gándara Fde la; García A; Raga JA, 2004. Effects of the gill parasite Zeuxapta seriolae (Monogenea: Heteraxinidae) on the amberjack Seriola dumerili Risso (Teleostei: Carangidae). Aquaculture, 232(1/4):153-163.
Moulder JW, 1984. Order II. Chlamydiales Stortz and Page 1971, 334AL. In: Kreig N, ed. Bergey’s Manual of Systematic Bacteriology, Vol. 1. Baltimore, London, UK: Williams and Wilkins, 729-739.
Paperna I, 1977. Epitheliocystis infection in wild and cultured sea bream (Sparus aurata, Sparidae) and grey mullets (Liza ramada, Mugilidae). Aquaculture, 10:169-176.
Paperna I; Sabnai I, 1980. Epitheliocystis disease in fishes. Fish diseases--third COPRAQ session, 228-234; [7 fig.].
Paperna I; Sabnai I; Castel M, 1978. Ultrastructural study of epitheliocystis organisms from gill epithelium of the fish Sparus aurata (L.) and Liza ramada (Risso) and their relation to the host cell. Journal of Fish Diseases, 1(2):181-189.
Plehn M, 1920. Neue Parasiten im Haut und Kiemen von Fischen Ichthyochytrium und Mucophilus. Zentralblatt für Bakteriologie, Parasitenkunde de Infektionskrancheiten und Hygiene. Abt I Originale, 85:275-281.
Turnbull JF, 1993. Epitheliocystis and salmonid rickettsial septicaemia. In: Inglis V, Roberts RJ, Bomage NR (eds), Bacterial diseases of fish. Oxford, UK: Blackwell Scientific Publications, 237-254.
Voronin VN; Chernysheva NB, 1997. Epitheliocystis infection in common carp Cyprinus carpio L.: histopathology and pathogenicity. Bulletin of the European Association of Fish Pathologists, 17(3/4):137-139.
Wolke RE; Wyand DS; Khairallah LH, 1970. A light and electron microscopic study of epitheliocystis disease in the gills of Connecticut striped bass (Morone saxatilis) and white perch (Morone americanus). Journal of Comparative Pathology, 80:559-563.
Work TM; Rameyer RA; Takata G; Kent ML, 2003. Protozoal and epitheliocystis-like infections in the introduced bluestripe snapper Lutjanus kasmira in Hawaii. Diseases of aquatic organisms, 57:59-66.
Zachary A; Paperna I, 1977. Epitheliocystis disease in the striped bass Morone saxatilis from the Chesapeake Bay. Canadian Journal of Microbiology, 23(10):1404-1414.
Distribution MapsTop of page
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