enteric septicaemia of catfish
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Top of page| Picture | Title | Caption | Copyright |
|---|---|---|---|
| Edwardsiella ictaluri | Channel catfish infected with Edwardsiella ictaluri. Fish A has haemorrhage in the skin under the jaw and isthmus (arrow). Fish B has white depigmented lesions on the pigmented skin (arrowheads), red ulcerated lesions on the lower gill cover (arrow) and abdominal distension, caused by ascitic fluid in the coelomic cavity. | J. A. Plumb | |
| Edwardsiella ictaluri | Channel catfish infected with Edwardsiella ictaluri exhibiting open lesion (large arrow) in the cranial region and inflamed nares (arrowhead) and exophthalmia typical of chronic infection. | J. A. Plumb | |
| Seasonal occurrence of Edwardsiella ictaluri | Seasonal occurrence of Edwardsiella ictaluri, showing greatest incidence of disease in May, June, September and October, when average water temperatures are 20-27°C. | J. A. Plumb | |
| Survival of Edwardsiella ictaluri | Survival of Edwardsiella ictaluri in pond water and mud at different temperatures. (From Plumb and Quinlan, 1986; reprinted with permission of the American Fisheries Society.) | J. A. Plumb | |
| Edwardsiella septicaemia | Edwardsiella ictaluri, E. tarda, Aeromonas hydrophila and Pseudomonas fluorescens on Edwardsiella isolation media incubated at 25°C for 48 h. (Photo by D. Earlix.) | J. A. Plumb | |
| Edwardsiella ictaluri isolated | Edwardsiella ictaluri isolated from organs and tissues 44, 51 and 65 days after initial exposure to the pathogen. HK, head kidney; BL, blood; BR, brain; L, liver; TK, trunk kidney; SP, spleen; G, gonads; GB, gall bladder; M, muscle. (From Mgolomba and Plumb, 1992, reprinted with permission of Elsevier Science Publishers.) | J. A. Plumb | |
| Edwardsiella ictaluri | Electron micrographs of Edwardsiella ictaluri in olfactory organ of channel catfish. (A) Scanning electron micrograph of the olfactory mucosal surface following direct experimental E. ictaluri infection with a cluster of bacteria attached to the epithelial surface. Note the fine filamentous processes extending from E. ictaluri (arrows) (x 13,500). (B) Transmission electron micrograph of the olfactory mucosal surface 1 h following direct experimental E. ictaluri infection. The bacteria (large arrows) are close to the epithelial surface (arrowhead), which has no cellular cilia but has increased secretory vesicles of host mucosal cells (small arrows) (x 2565). (C) Transmission electron micrograph showing several catfish leucocytes in the interlamellar space of the olfactory rosette. A number of E. ictaluri can be seen within cellular phagosomes (arrows) (x 4590). (Photographs courtesy of E.E. Morrison and K.G. Wolfe.) | J. A. Plumb | |
| Edwardsiella ictaluri | Paraffin sections of tissue from a channel catfish infected with Edwardsiella ictaluri. (A) Necrotic skeletal muscle (N) with infiltrating macrophages (arrow) typical of granulomatous inflammation. A giant cell (GC) is present within the accumulation of macrophages. (H & E, x 450.) (B) Focal accumulation of macrophages (M) in the liver. Within this lesion, the macrophages have displaced most of the liver tissue, but scattered hepatocytes (arrow) are still present. (H & E, x 1000.) (Photograph by A. Goodwin.) | J. A. Plumb |
Identity
Top of pagePreferred Scientific Name
- enteric septicaemia of catfish
International Common Names
- English: Edwardsiella ictaluri infection; Edwardsiella septicaemia (Edwardsiella ictaluri infection); hole in the head; hole-in-the-head
English acronym
- ESC
Overview
Top of pageThe genus Edwardsiella includes two species of bacteria that cause major diseases in fish. Edwardsiella tarda (Ewing et al., 1965) infects fish and other animals and Edwardsiella ictaluri (Hawke, 1979) infects fish only. A third species, Edwardsiella hoshinae (Grimont et al., 1980), infects birds and reptiles Edwardsiella tarda produces the disease commonly known as fish gangrene, emphysematous putrefactive disease of catfish or red disease in eels, and will be referred to as Edwardsiella septicemia (ES). E. ictaluri causes enteric septicemia of catfish (ESC). Because E. tarda and E. ictaluri produce distinctly different diseases, they are discussed separately.
Enteric septicaemia of catfish (ESC) is caused by the bacterium Edwardsiella ictaluri, which belongs to the Enterobacteriaceae family (Hawke et al., 1981). ESC is one of the most important infectious disease problems in the commercial catfish industry in the USA. Most reported cases of disease caused by E. ictaluri are in channel catfish (Ictalurus punctatus), but the bacterium has been isolated from related North American catfish including blue catfish (I. furcatus), white catfish (Ameiurus catus), brown bullhead (A. nebulosus) (Hawke et al., 1981) and wild tadpole madtom (Noturus gyrinus) (Klesius et al., 2003). Edwardsiella ictaluri has also been reported from walking catfish (Clarias batrachus) in Thailand (Kasornchandra et al., 1987), Pangasius hypothalamus in Vietnam (Crumlish et al., 2002) and from several ornamental species (Kent and Lyons, 1982; Waltman et al., 1985). The susceptibility of other fish species including salmonids has been shown experimentally (Baxa et al., 1990). Several studies have shown that E. ictaluri is a biochemically, genetically and perhaps serologically, homogeneous species (Bertolini et al., 1990; Newton et al., 1988; Starliper et al., 1988; Waltman et al., 1986).
Acute outbreaks of ESC occur within a limited temperature range, from 18 to 28°C. This critical temperature window makes spring and autumn the most common periods for outbreaks in regions where channel catfish are normally cultured. However, low-level mortality due to ESC can occur in carrier populations outside of this temperature range. Other environmental factors (poor water quality, high stocking density and other stressors) predispose the host to ESC. Edwardsiella ictaluri is generally considered to be an obligate pathogen.
Two clinical forms of ESC occur in channel catfish, a chronic encephalitis and an acute septicaemia (Miyazaki and Plumb, 1985; Newton et al., 1989; Shotts et al., 1986). In the chronic form the bacterium infects the olfactory sacs, and migrates along the olfactory nerves to the brain, generating granulomatous inflammation (Morrison and Plumb, 1994). This meningo-encephalitis causes abnormal behaviour, with alternating listlessness and chaotic swimming. In late stages of this disease, swelling develops on the dorsum of the head as the inflammatory process erodes the connective tissue in this region. This swelling ulcerates, exposing the brain, which has lead to the term ‘hole in the head disease’, used in the industry. In the acute form of ESC the bacterium is thought to infect through the intestinal mucosa (Baldwin and Newton, 1993), and then to establish a bacteraemia. The affected fish display petechial haemorrhages around the mouth, on the throat, the abdomen and at the base of the fins. Multifocal distinct 2 mm diameter raised haemorrhagic cutaneous lesions that progress to depigmented ulcers also occur. Anaemia, moderate gill inflammation and exophthalmia are common signs. Internally, haemorrhages and necrotic foci are scattered in the liver and other internal organs. Haemorrhagic enteritis, systemic oedema, accumulation of ascitic fluid in the body cavity and enlargement of the spleen are nonspecific signs. Histological examination reveals a systemic infection of all organs and skeletal muscles, with the most severe changes being diffuse interstitial necrosis of the anterior and posterior kidney. Focal necrosis in the liver and spleen are also generally seen.
Fish from a population that has recovered from the disease are considered to be immune carriers that may have high levels of E. ictaluri-specific antibodies. Occasional losses due to recurrent ESC will occur in these populations, especially after stress is induced. Edwardsiella ictaluri has been detected in the kidney of such fish well over 4 months after exposure (Antonio-Baxta and Hedrick, 1994; Klesius, 1992), suggesting that carrier fish act as the natural reservoir for the organism. It is believed that shedding with faeces is the main mode of dissemination into the environment. The pathogen persistence and the common practice of continual partial harvest and stocking within a production pond have contributed to the success of this pathogen and the prevalence of ESC in the industry. Moreover, the agent can survive in pond sediments for an extended period of time (Plumb and Quinlan, 1986), and this may be another important factor in disease resurgence in given areas. Researchers have found the bacterium in the gut of fish-eating birds by performing fluorescent antibody tests on ingesta, but generally no E. ictaluri could be cultured indicating that the bacteria were not viable (Taylor, 1992; Waterstrat et al., 1999). This suggests, that birds are not an important means of disseminating this pathogen.
ESC may be controlled through chemotherapy and/or prophylactic measures. The most common antimicrobial treatments are oral application of the potentiated sulfonamide sulfadimethoxine ormethoprim or oxytetracycline, but plasmid-mediated resistance to these antibiotics does occur (Cooper et al., 1993). Many producers are now focusing on alternative methods to reduce losses. This relies on management to reduce stress in fish, the cessation of feeding when ESC-induced losses are detected (Wise and Johnson, 1998) and on vaccination (Klesius and Shoemaker, 1999; Shoemaker et al., 1999; Wise et al., 2000; OIE, 2003)
Enteric septicaemia of catfish, caused by E. ictaluri, was first reported by Hawke (1979). However, Mitchell and Goodwin (1999) showed by histochemistry on formalin preserved diseased catfish that the bacterium may have been present in Arkansas in the late 1960’s. Since its discovery, this disease has become the most important infectious disease of the catfish industry in the USA especially in the southeast (Wagner et al., 2002). Estimates of the cost of the disease to the catfish industry have been in the US$l0’s of millions annually, but a carefully calculated assessment of losses is not available. Because of its comparatively narrow host specificity, ESC is not a great economic problem in regions where channel catfish are not cultured, although the bacterium has been isolated from other catfishes and species of fish in the wild in other parts of the world (Plumb, 1999b).
[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 Animals
Top of pageHosts/Species Affected
Top of pageEdwardsiella ictaluri has a narrower host range than that of E. tarda; however, it is still diverse. Cultured channel catfish are most severely affected, but less susceptible ictalurids include white catfish (Ameiurus catus), blue catfish (Ictalurus furcatus), rarely brown bullhead (Ameiurus nebulosus), wild tadpole madtom (Noturus gyrinus) (Klesius et al., 2003) and wild channel catfish in California (Chen et al., 1994). Experimental infections in blue catfish have been difficult but an occasional natural infection has occurred in this species and variances in susceptibility in channel catfish strains have been demonstrated (Wolters and Johnson, 1994). Additionally, Wolters et al. (1996) showed that, following experimental infection, channel catfish had the lowest survival (62%) and blue catfish had the highest survival (90%), while survival of hybrids of the two species was intermediate (74%). Natural infections in non-ictalurids include walking catfish (Clarias batrachus) (Kasornchandra et al., 1987), freshwater catfish (Pangasius hypothalamus) (Crumlish et al., 2002) and several ornamental species(Waltman et al., 1985; Kent and Lyons, 1982; Humphrey et al., 1986). Experimental infections were established in chinook salmon (Onchorhynchus tshawytscha) and rainbow trout (Oncorhynchus mykiss) (Baxa et al., 1990), but Plumb and Sanchez (1983) could not experimentally infect golden shiners (Notemigonus chrysoleucas), tilapia (Tilapia aurea), largemouth bass (Micropterus salmoides) or bighead carp (Aristichthys nobilis). Plumb and Hilge (1987) demonstrated that the European catfish (sheatfish) (Silurus glanis) was only slightly susceptible.
There is no indication that E. ictaluri poses a health threat to aquatic animals other than a limited number of fish species. The temperature limitations under which E. ictaluri grows essentially preclude this bacterium from being a pathogen for humans or other warm-blooded animals (Janda et al., 1991).
Distribution
Top of pageEdwardsiella ictaluri has been confirmed only in the USA, Thailand (Kasornchandra et al., 1987) and Australia (Humphrey et al., 1986); however, there is a report of its presence in Taiwan (Chung and Kou, 1983), Vietnam (Crumlish et al., 2002), China (Tan et al., 2003) and there have been unconfirmed reports of clinical signs typical of ESC in other parts of the world where catfish are cultured. In the USA, E. ictaluri is found primarily across the southeastern region, where channel catfish are grown commercially (i.e., Alabama, Arkansas, Louisiana, Mississippi, Texas, Florida, Georgia, North Carolina and South Carolina). However, the bacterium has been reported to cause disease among cultured channel catfish in other states, such as Arizona, California, Idaho, Indiana, Kansas, Maryland, New Mexico and Virginia. The bacterium was recently isolated from wild tadpole madtom in New Jersey. With the continual worldwide dissemination of channel catfish for aquaculture purposes and an inadequate method of detecting non-clinical infections, it is likely that E. ictaluri will occur in other geographical regions (Plumb, 1999b).
Distribution Table
Top of pageThe distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
Last updated: 10 Jan 2020Pathology
Top of pageHistopathologically, the trunk kidney and spleen are the most severely affected organs in channel catfish, both of which are necrotic, while the liver is oedematous and necrotic (Areechon and Plumb, 1983). Proliferation occurs in interlamellar tissue in gills (Jarboe et al., 1984; Miyazaki and Plumb, 1985; Shotts et al., 1986). Also, a mild focal infiltration, necrosis and granulomatous inflammation take place in the underlying musculature in areas where the epidermis is missing. Intact E. ictaluri cells are also seen in macrophages, such that occurs with E. tarda.
The most complete pathological study of E. ictaluri was that of Newton et al. (1989). Following experimental exposure of channel catfish to 5 × 108 cfu ml-1, 93% of the affected fish developed acute ESC and 7% developed chronic infection. Acute disease was characterized grossly by haemorrhage and ulceration, and microscopically by enteritis and by olfactory sacculitis at 2 days postexposure, followed by hepatitis and dermatitis. Chronic ESC, seen at 3-4 weeks, post-exposure was characterized by dorsocranial swelling and ulceration, granulomatous inflammation and meningoencephalitis of the olfactory bulbs, olfactory tracts and olfactory lobes of the brain. The granulomatous inflammation in chronic E. ictaluri infection is a key histopathological characteristic of ESC. Skeletal muscle becomes necrotic, with infiltration of macrophages, while internal organs, especially the liver, have normal tissue displaced by macrophages.
Diagnosis
Top of pageEnteric septicaemia of catfish may be mild, chronic or acute. Diseased fish are listless at the surface, with ‘head-up, tail-down’ posture and sometimes spin in circles before death. Other characteristic clinical signs are petechial haemorrhage or inflammation in the skin under the jaw, on the operculum and belly; haemorrhaging often becomes so severe that the skin is bright red. Haemorrhage also occurs at the base of fins. Small white (1-3 mm) depigmented areas appear on the skin and they progress into similar-sized inflamed cutaneous ulcers. An open lesion develops between the frontal bones of the skull posterior to, or between, the eyes in chronically ill fish hence the common name ‘hole-in-the-head’. It should be noted that other bacteria (Aeromonas hydrophila, for example) can cause the same lesion. Infected fish also have pale gills, exophthalmia and sometimes abdominal distension with ascites fluid that is usually cloudy and/or bloody and rarely clear yellow. The kidney and spleen are hypertrophied, while the spleen is dark red. Inflammation occurs in adipose tissue, peritoneum and intestine, and the liver is either pale or mottled with congestion.
Clinical signs of E. ictaluri infection are more pathognomonic than most other infectious fish diseases and therefore are helpful in ESC diagnosis. However, isolation of the organism and/or serological tests or PCR are essential for confirmation (OIE 2003). Edwardsiella ictaluri is isolated from clinically infected fish on BHI or tryptic soy agar (TSA), but Shotts and Waltman (1990) developed EIM which enhances E. ictaluri isolation and aids identification. The organism forms small, translucent, greenish colonies on EIM, while inhibiting Gram-positive and most Gram-negative contaminating organisms. Colonies of E. tarda have black centres and A. hydrophila colonies are brownish and larger, while Pseudomonas fluorescens colonies are blackish and punctate on EIM. Using the biochemical and biophysical characteristics, E. ictaluri can be easily separated from E. tarda, because the former is indole-negative and does not produce H2S on TSI agar. Commercial identification systems, such as Minitek and API 20E, are not as accurate for E. ictaluri as they are for some other fish pathogens (Taylor et al., 1995).
Careful observation of culture plates is essential to detect E. ictaluri, because of its slow growth and the possible presence of more rapidly growing bacteria, such as Aeromonas spp. Definitive identification is by using biochemical characteristics or serological identification with specific antiserum agglutination or other serological tests. These include polyclonal and monoclonal antibody in indirect FAT (Ainsworth et al., 1986) and ELISA (Rogers, 1981; Hanson and Rogers, 1989; Klesius, 1993; Earlix et al., 1996). Twenty E. ictaluri isolates were positive using FAT and ELISA, while no cross reactivity was detected with E. tarda, Salmonella sp. or A. hydrophila (Rogers, 1981). Ainsworth et al. (1986) also used monoclonal
antibodies against E. ictaluri in an indirect FAT application for diagnosing ESC. They compared the FAT techniques with bacterial isolation from brain, liver, spleen, anterior kidney and posterior kidney, and 90.3% of the culture-positive fish were also FAT-positive. The greatest discrepancy occurred in the brain samples, but tissues from the spleen were 90% positive by FAT and 85% by culture. All of these studies emphasized the time-saving advantage of immunoassay techniques of 2 h for results versus 48 h required for culture results. It is also possible to diagnose E. ictaluri in the carcasses of dead fish, using an ELISA system (Hanson and Rogers, 1989). Apparently, upon death, the bacterium escapes from lysed macrophages and uses the nutrients of these cells to proliferate, thus providing a large number of organisms.
Molecular techniques to detect E. ictaluri have been developed. Bilodeau et al. (2003) developed a specific real time PCR method that was able to detect E. ictaluri in blood and tissues. However, limitations of PCR should be realized (i.e. dead bacteria can be quantified) when interpreting data generated using these methods.
Detection of E. ictaluri carrier fish when there is no clinical disease may present a problem; however, Mgolomba and Plumb (1992) and Klesius (1992) found significant bacteria in the blood and all organs of fish 65 and 270 days, respectively, after initial exposure to E. ictaluri. Edwardsiella ictaluri is also readily phagocytized by macrophages in naïve fish, but immunization increases the phagocytic activity (Shoemaker et al., 1997). Indications are that the phagocytized bacteria are not destroyed in these cells, which could lead to a lengthy carrier state and could provide a site of identifying carrier fish (Miyazaki and Plumb, 1985; Klesius et al., 1991; Klesius, 1993;). Application of the Falcon screening test (FAST)-ELISA was utilized by Klesius (1993) to rapidly and accurately detect E. ictaluri antibody in adult fish and he proposed that the method could be used to identify possible E. ictaluri carrier fish. Earlix et al. (1996) utilized an ELISA and monoclonal antibody in conjunction with tissue homogenization, digestion of tissue with Triton X-100 and filtration on 0.45 mm nitrocellulose membrane to detect an 80% E. ictaluri carrier state in asymptomatic channel catfish. This is compared with 24% carrier-state detection by conventional bacteriological isolation methods. Implementation of these procedures could be useful in determining E. ictaluri carrier populations.
Serum agglutination, passive haemagglutination, complement-dependent passive haemolysis, indirect immunofluorescence, agar gel immunodiffusion and agglutination with fractionated immunized fish sera were used for detecting humoral antibody to LPS of E. ictaluri (Saeed and Plumb, 1987). All tests were sensitive to the LPS antibody, with the complement-dependent haemolysis titres being the highest (average titres of 1 : 2360). Waterstrat et al. (1989) found a close relationship of antibody titres measured by optical density and corresponding agglutination titres to E. ictaluri in channel catfish. Using the FAST-ELISA system employing a monoclonal antibody against an immunodominant epitope of E. ictaluri, Klesius et al. (1991) found no cross reactivity with sera from experimentally immunized channel catfish against E. tarda or A. hydrophila. The ELISA system was further refined so that either E. ictaluri antigen or antibody in fish can be detected in 30 min. Time is usually of the essence in diagnosing clinical E. ictaluri; hence, the rapid serological techniques should be used in conjunction with bacterial isolations.
List of Symptoms/Signs
Top of page| Sign | Life Stages | Type |
|---|---|---|
| Finfish / Blood spots in muscle of body cavity wall - Body Cavity and Muscle | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Build up of bloody fluids - Body Cavity and Muscle | Aquatic:Adult,Aquatic:Fry | Diagnosis |
| Finfish / Cessation of feeding - Behavioural Signs | Aquatic:Adult,Aquatic:Fry | Diagnosis |
| Finfish / Change in feed-conversion ratio - Miscellaneous | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Change in feed-conversion ratio - Miscellaneous | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Change in feed-conversion ratio - Miscellaneous | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Change in feed-conversion ratio - Miscellaneous | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / 'Dropsy' - distended abdomen, 'pot belly' appearance - Body | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Fish swimming near surface - Behavioural Signs | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Fish swimming near surface - Behavioural Signs | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Generalised lethargy - Behavioural Signs | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Generalised lethargy - Behavioural Signs | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Haemorrhagic lesions - Skin and Fins | Aquatic:Adult,Aquatic:Fry | Diagnosis |
| Finfish / Haemorrhaging - Body Cavity and Muscle | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Haemorrhaging - Body Cavity and Muscle | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Hole-in-head (longitudinal opening in cranial foramen) - Miscellaneous | Aquatic:Adult | Diagnosis |
| Finfish / Liver - white / grey patches (haemorrhage / necrosis / tissue damage) - Organs | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Liver - white / grey patches (haemorrhage / necrosis / tissue damage) - Organs | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Liver - white / grey patches (haemorrhage / necrosis / tissue damage) - Organs | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Loss of balance - Behavioural Signs | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Loss of balance - Behavioural Signs | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Mortalities -Miscellaneous | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Mortalities -Miscellaneous | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Mortalities -Miscellaneous | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Mortalities -Miscellaneous | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Mucus-filled intestines - Organs | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Paleness - Gills | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Paleness - Gills | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Pop-eye - Eyes | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Pop-eye - Eyes | Aquatic:Adult,Aquatic:Fry | Sign |
| Finfish / Red spots: pin-point size (petechiae) - Skin and Fins | Aquatic:Adult,Aquatic:Fry | Diagnosis |
Disease Course
Top of pageThe pathogenic mechanism of E. ictaluri infection in channel catfish is not fully understood. Janda et al. (1991) demonstrated that the bacteria did not invade HEp-2 cell monolayers at 35°C, nor did they produce cell-associated haemolysin or siderophores, as do E. tarda. In examining ECP being associated with the pathogenesis of E. ictaluri, Stanley et al. (1994) found a fibrillar network connecting virulent cells that could aid in attachment. Virulent isolates had greater amounts of capsular material and surface proteins, and they demonstrated a greater ability to degrade chondroitin than did avirulent cells. These authors reported no clear correlation between haemolytic activity and virulence.
Edwardsiella ictaluri infects fish by several routes. Water-borne bacteria can invade the olfactory organ via the nasal opening and migrate into the olfactory nerve, then into the brain meninges and finally to the skull and skin (Miyazaki and Plumb, 1985; Shotts et al., 1986; Morrison and Plumb, 1994). Injury to the nasal passage includes loss of sensory cilia and microvilli from the olfactory mucosal surface (Morrison and Plumb, 1994) within 1 h of exposure to E. ictaluri. By 24 h, the olfactory receptors and supporting cells were degenerating; electron microscopy confirmed the presence of E. ictaluri on the mucosal surface and within the epithelium. Host leucocytes migrated through the olfactory epithelium into the lamellar lumen and phagocytized the bacterium. With regard to the attachment mechanism of E. ictaluri, Wolfe et al. (1998) showed that bacterial lectins were instrumental in this attachment by utilizing specific sugar residues, specifically D-mannose, N-acetylneuraminic acid and L-fucose, in the nasal mucosa.
Edwardsiella ictaluri apparently colonizes capillaries in the dermis and causes necrosis and depigmentation of the skin of infected fish. In the intestine, E. ictaluri enters the blood through the intestinal wall and is engulfed by macrophages, resulting in septicaemia (Shotts et al., 1986; Newton et al., 1989). Channel catfish exposed to E. ictaluri via oral infection developed enteritis, hepatitis, interstitial nephritis and myositis within 2 weeks of infection. Francis-Floyd et al. (1987) described gastrointestinal lesions, including petechia or ecchymoses in the mucosa of the gastrointestinal tract and intestinal distension associated with gas production. The gill is also a primary site of E. ictaluri invasion. Using radiolabelled E. ictaluri, Nusbaum and Morrison (1996) demonstrated that, during immersion, the organism colonizes the gill epithelium in large numbers in 2-72 h. Bacterial numbers then increased rapidly in the liver and less rapidly in the trunk kidney, gut and brain. It is worth noting that heat-killed radiolabelled bacteria did not appear to cross the gill epithelium membrane.
Epidemiology
Top of pageEnteric septicaemia of catfish is considered a seasonal disease, occurring primarily in late spring to early summer and again in autumn. This pattern generally coincides with, but is not confined to, water temperatures of 18-28°C. Francis-Floyd et al. (1987) demonstrated that the highest mortality in experimentally infected channel catfish fingerlings was at 25°C, lower at 23 and 28°C and no deaths at 17, 21 or 32°C. Several experiments have further substantiated the temperature preference of ESC. Baxa-Antonio et al. (1992) used immersion of channel catfish in a bath containing E. ictaluri to show peak mortality at 25°C (98%) and lower mortality at 20°C (47%), 30°C (25%), 35°C (4%) and 15°C (0%). The effect of 25°C on clinical ESC was further demonstrated by Plumb and Shoemaker (1995) using a naturally infected population of channel catfish, in which 10% were culture-positive for E. ictaluri while being held at 15°C. When these fish were elevated to 25°C, 77% mortality occurred due to E. ictaluri, which was significantly higher than that at 18°C or 30°C (10% and 23%, respectively).
In spite of the compelling experimental data that implicates a mid-20°C optimum temperature, an increasing incidence of ESC in diagnostic case work during winter and summer has been noted, indicating possible adaptation of E. ictaluri to a broader temperature range. During the early years following the discovery of ESC, relatively few outbreaks of the disease were reported, but the number of E. ictaluri isolates soon began to occur at an alarming rate. In 1981, there were 47 outbreaks diagnosed in southeastern USA and, in 1985, there were 1420 diagnosed outbreaks, thus accounting for 28% of all reported fish-disease cases in the region (A.J. Mitchell, Fish Farming Experiment Station, Stuttgart, Arkansas, 1996, personal communication). In 1988, there were 1605 documented reports of ESC (30.4% of reported fish diseases); however, the prevalence levelled off during 1990 and 1991 and has remained constant since then. In a survey by Wagner et al. (2002), it was found that 78.1% of all channel catfish operations and 42.1% of production ponds experienced columnaris and/or enteric septicemia, or a combination of the two with no distinction being made between the two. Average losses were from 90 to 900kg of fish per disease episode.
Mortality in naturally infected channel catfish populations varies from less than 10% to over 50%. It occurs in juvenile as well as food-sized fish, and under all types of cultural conditions (including ponds, raceways, recirculating systems and cages). Few fish diseases occur without some environmental stressor preceding the infection, but E. ictaluri can probably cause disease independent of stressors. This is not to suggest that adverse environmental conditions do not influence the severity of infection, because Wise et al. (1993a) showed that, when channel catfish were stressed by confinement in tanks prior to E. ictaluri exposure, there was 97% mortality in stressed fish and 77% mortality in non-stressed fish. To further illustrate the effects of stress, Ciembor et al. (1995) netted and handled channel catfish and then exposed them to water-borne E. ictaluri, leading to a mortality of 53% in stressed fish and 16% in unstressed fish. It was also shown by Plumb et al. (1993b) that stocking density in ponds may affect susceptibility to E. ictaluri.
In aquaculture, infected channel catfish are the primary source of E. ictaluri, with natural transmission occurring primarily through the water. Horizontal transmission of E. ictaluri was demonstrated by Klesius (1994), in which naïve fish showed clinical infections 12 days post exposure to fish that had died of ESC. Experimental transmission of E. ictaluri is easily achieved by water-borne exposure, intramuscular or intraperitoneal injection, intestinal intubation or introducing the bacterium into the nares only (Plumb and Sanchez, 1983; Shotts et al., 1986; Newton et al., 1989; Morrison and Plumb, 1994). Acute clinical disease usually appears at 5-7 days post-bath exposure at 25°C. Nusbaum and Morrison (1996) showed that the pathogen invaded the gills and then migrated to other organs and tissues. The nares are a primary site of E. ictaluri invasion, and exposure of this organ to E. ictaluri can initiate chronic ESC (Morrison and Plumb, 1994). It was shown by Mgolomba and Plumb (1992) and Klesius (1992) that survivors of an epizootic still carry E. ictaluri long after clinical disease has disappeared; therefore, they can serve as reservoirs of the pathogen. Research indicated that, in a pond where fish were dying of E. ictaluri, the water in the vicinity of dead fish had significantly higher E. ictaluri counts than waters where there were no carcasses (Earlix, 1995). Removal of dead fish should reduce the number of bacteria to which non-infected fish are exposed. Transmission from adults to offspring during spawning is likely but remains as yet unproved.
MacMillan and Santucci (1990) were unable to isolate E. ictaluri from the intestine of channel catfish, but Earlix (1995) isolated the organism on Edwardsiella isolation media (EIM) from intestines of approximately 50% of clinically infected channel catfish.
Once E. ictaluri is introduced into a particular body of water, the bacterium probably remains there as a source of infection, either in carrier fish or in the environment. The way in which E. ictaluri is transferred from farm to farm is speculative, but there is little doubt that the transfer of infected fish is the primary instrument of transmission, as well as other means; for example, seines, nets, etc. that are not disinfected or thoroughly air-dried between use could be sources of infections.
Impact Summary
Top of page| Category | Impact |
|---|---|
| Biodiversity (generally) | Negative |
| Fisheries / aquaculture | Negative |
| Native fauna | Negative |
Impact: Economic
Top of pageThe economic impact of E. ictaluri on the channel catfish industry is significant. Enteric septicemia along with the bacterial disease "columnaris" are the two most serious infectious diseases affecting the industry (Wagner et al., 2002). While not specifically measuring the economic impact of ESC they emphasized "that the majority of producers reported them (enteric septicemia and columnaris) as the diseases causing the greatest economic impact." Earlier Plumb and Vininantharat (1993) reported that Edwardsiella ictaluri was responsible for US$20 to 30 million losses annually and Shoemaker et al. (2003) estimated that ESC costs the catfish industry US$60 million annually.
Zoonoses and Food Safety
Top of pageEdwardsiella ictaluri poses no infectious threat to non-fish species or to any warm blooded animals, nor does it present a food safety problem. However, Skirpstunas and Baldwin (2002) showed that E. ictaluari can attach to IEC-6 (rat small intestinal cells) and Henie 407 (human embryonic intestinal epithelium cells) in culture at a low rate, thus indicating that under some circumstances this bacterium may invade intestinal cells of warm blooded animals but is unlikely to flourish because of its lower temperature intolerance.
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Distribution References
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Links to Websites
Top of page| Website | URL | Comment |
|---|---|---|
| Enteric Septicaemia of Catfish | http://www.oie.int/eng/normes/fmanual/a_00029.htm | Included in OIE Manual of Diagnostic Tests for Aquatic Animals. |
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