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furunculosis in fish

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furunculosis in fish

Summary

  • Last modified
  • 20 November 2019
  • Datasheet Type(s)
  • Animal Disease
  • Preferred Scientific Name
  • furunculosis in fish
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Identity

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Preferred Scientific Name

  • furunculosis in fish

International Common Names

  • English: Aeromonas salmonicida infection; carp erythrodermatitis; furunculosis; head ulcer disease of cultured Japanese eels; ulcer disease of goldfish

Overview

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Aeromonas salmonicida has been recognized as a pathogen of fish for over 100 years. Emmerich and Weibel (1894) made the first authentic report of its isolation during a disease outbreak at a Bavarian brown trout hatchery, the manifestations of the disease including furuncle-like swelling and, at a later stage, ulcerative lesions on infected trout. Since that time a number of subspecies of A. salmonicida have been recognized, although the taxonomy of the species is far from settled. A. salmonicida is one of the most studied fish pathogens, because of its widespread distribution, diverse host range and economically devastating impact on cultivated fish, particularly the salmonids (Austin and Austin, 1993). The continued importance of salmonids to rod fishermen, commercial fisheries and fish farmers and the extent of the impact that A. salmonicida has on these various methods of exploitation have served to maintain the status of A. salmonicida as an important fish pathogen. A number of excellent reviews of A. salmonicida have been published (McCarthy and Roberts, 1980; Austin and Austin, 1993; Bernoth et al., 1997). This datasheet attempts to summarize the current information, both research and anecdotal, available on A. salmonicida and its associated pathologies in a manner accessible to all those for whom this organism is of concern.

‘ATYPICAL’ AEROMONAS SALMONICIDA INFECTION

A salmonicida subsp. salmonicida is an important salmonid pathogen that has probably been studied more than any other fish pathogen. However, the species A. salmonicida comprises a large number of other strains, routinely referred to as ‘atypical’ strains, that is, strains that do not conform to the general guidelines presented for typical A. salmonicida. These atypical strains are responsible for several disease conditions in both salmonids and non-salmonid species. Our knowledge of ‘atypical’ strains is growing rapidly and their importance is being increasingly recognized. In the last decade, atypical isolates of A. salmonicida have become a growing concern in several countries, because of their frequent association with mortalities of both salmonids and non-salmonid species. Recent studies from Canada and northern Europe indicate that cases of ‘furunculosis’ in salmonids caused by atypical isolates are increasing, probably because of better awareness of these subspecies. This confirms the belief that atypical isolates are more widely distributed than previously suspected (Wichardt et al., 1989; Rintamäki and Valtonen, 1991; Olivier, 1992; Pedersen et al., 1994; Wiklund et al., 1994; Wiklund and Dalsgaard 1998).

The literature on ‘atypical’ strains of A. salmonicida is expanding rapidly but remains confusing, largely because of the problems associated with their taxonomy and the variety of disease conditions they cause in different species of fish. Atypical strains are often reviewed with typical isolates (McCarthy and Roberts, 1980; Trust, 1986; Austin and Austin, 1993), although they are distinct subspecies.

Future Research

There is still much to be learned about A. salmonicida and its associated pathologies. However, as long as there is continued availability of oil-based vaccines, furunculosis will no longer be perceived as a major problem for commercial marine salmonid farmers. Therefore, future research must focus on the appropriate and sustainable management of freshwater fisheries, in particular restoration programmes and native fisheries. Many of the disease control strategies available to commercial farmers such as vaccination and chemotherapy may not be appropriate in these situations. For example, the advisability of vaccinating fish with oil-based preparations prior to their release into river systems, from which they may be removed for consumption by anglers must be questioned. Oil based adjuvants are highly immunogenic, not only for fish but for consumers of those fish. Likewise, the presence of chemotherapeutic residues in the flesh of fish which are available for consumption is undesirable. Therefore, the control strategies that can be adopted by fresh water fisheries managers will be affected by different concerns than the more controlled commercial sector.

A number of important questions can be identified for freshwater fisheries in relation to furunculosis. These may be summarized as follows:

do fresh water hatcheries act as amplifiers of disease for the water body on which they are situated and how can this effect be minimized or eliminated?

what impact will the release of covertly infected fish into a water body have on their survival and on resident fish populations, and how can this impact be minimized or eliminated?

does vaccination of covertly infected hatchery stocks prior to release constitute the creation of ‘immune carriers’, what impact are these ‘immune carriers’ likely to have on non-vaccinated fish and how can this impact be minimized or eliminated?

how can hatchery reared stocks be protected from the impact of anadromous fish in the water body on which the hatchery is situated?

Many of these questions have yet to be answered. It is hoped, therefore, that future research into furunculosis will address the fundamental concerns outlined here. Only in this way will we limit both the commercial and biological impact of furunculosis among valuable and endangered fish populations. Non-culture-based methods may have a valuable contribution to make to the study of furunculosis epizootiology but much work on the validation of these techniques needs to be done before they can be applied to field studies. The presence of atypical strains in wild marine fish with ulcerations is also of concern and their impact on wild fish populations is still unknown. The importance of these isolates originating from wild marine fish could represent a threat to the future culture of these species in aquaculture conditions.

A second important problem associated with atypical A. salmonicida isolates is that they are highly variable in their biochemical profile and cannot be easily placed in known or accepted subspecies. Further work on their taxonomy will be necessary to unravel this basic problem. Results so far indicate that atypical strains are for the most part highly restricted to precise geographic areas. However, some strains including goldfish and carp isolates have a wide distribution probably due to the international trade of these species. The limited work carried out to date would indicate that atypical strains are restricted to their host of origin; this is especially true for atypicals isolated from non-salmonid fish. Most of the strains isolated from these various hosts have so far been biochemically different. More work on this issue will, however, be necessary to gain a fuller epizootiological picture of atypical A. salmonicida and its associated pathologies.

[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

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Animal nameContextLife stageSystem
Abramis brama (carp bream)
Alburnus alburnus (bleak)
Ammodytes tobianus
Anarhichas lupus (wolf-fish)
Anarhichas minor (spotted wolffish)Domesticated hostAquatic: All StagesEnclosed systems/Cages
Anguilla anguilla (European eel)Aquatic: All StagesEnclosed systems/Freshwater recirculating systems|Enclosed systems/Ponds|Enclosed systems/Raceways / running water ponds|Enclosed systems/Tanks
Anguilla japonica (Japanese eel)Wild host
Anguilla rostrata (american eel)Experimental settings
Anoplopoma fimbria (sablefish)Wild host
Aristichthys nobilis (bighead carp)
Bidyanus bidyanus (silver perch)
Blicca bjoerkna
Carassius auratus auratus (goldfish)Wild hostAquatic: All Stages
Catostomus commersonii (white sucker)Experimental settings
Centrolabrus exoletusExperimental settings
Clupea pallasii (pacific herring)
Coregonus autumnalis pollan
Cottus gobioExperimental settings
Ctenolabrus rupestris (goldsinny-wrasse)Experimental settings
Culaea inconstans
Cyprinus carpio (common carp)Experimental settings
Eopsetta
Esox lucius (pike)Experimental settings
Gadus morhua (Atlantic cod)
Gasterosteus aculeatus aculeatus (three-spined stickleback)
Hexagrammos
Hippoglossoides platessoides
Hippoglossus hippoglossus (Atlantic halibut)Domesticated hostAquatic: All StagesEnclosed systems/Pens
Hybognathus hankinsoni
Hyperoplus lanceolatus
Hypophthalmichthys molitrix (silver carp)
Ichthyomyzon castaneus
LabridaeExperimental settings
Labrus mixtus
Lates calcarifer (barramundi)Domesticated host
Latris lineata (striped trumpeter)
Limanda limanda
Luxilus cornutusExperimental settings
Melanogrammus aeglefinus (haddock)
Microgadus tomcod
Micropterus dolomieu (smallmouth bass)Experimental settings
Morone mississippiensis (yellow bass)Experimental settings
Notemigonus crysoleucas (golden shiner)Experimental settings
Oncorhynchus gorbuscha (pink salmon)Wild hostAquatic: All Stages
Oncorhynchus keta (chum salmon)Domesticated host, Experimental settings, Wild hostAquatic: All Stages
Oncorhynchus kisutch (coho salmon)Experimental settingsAquatic: All Stages
Oncorhynchus masou masou (cherry salmon)
Oncorhynchus masou rhodurus
Oncorhynchus mykiss (rainbow trout)Experimental settings, Wild hostAquatic: All Stages
Oncorhynchus nerka (sockeye salmon)Experimental settingsAquatic: All Stages
Oncorhynchus tshawytscha (chinook salmon)Domesticated host, Wild hostAquatic: All Stages
Ophiodon elongatusWild host
Paralichthys olivaceus (bastard halibut)
Perca flavescens (yellow perch)
Perca fluviatilis (perch)
Phoxinus eosExperimental settings
Phoxinus phoxinus (European minnow)
Pimephales promelas (fathead minnow)Experimental settings
Platichthys flesus (flounder)
Pleuronectes platessa (European plaice)
Pollachius virens
Polyodon spathula (mississippi paddlefish)Experimental settings
Psetta maxima (turbot)Domesticated host, Experimental settingsAquatic: All StagesEnclosed systems/Marine recirculating systems|Enclosed systems/Tanks
Rhombosolea tapirina (greenback flounder)
Rutilus rutilus (roach)
Salmo salar (Atlantic salmon)Domesticated host, Experimental settings, Wild hostAquatic: All Stages
Salmo trutta (sea trout)Domesticated host, Experimental settings, Wild hostAquatic: All Stages
SalmonidaeDomesticated host, Wild hostAquatic: All Stages
Salvelinus alpinus (Arctic charr)Aquatic: AdultOpen water systems/Enhancements and culture-based fisheries (inc. ranching and stock enhacement)
Salvelinus fontinalis (brook trout)Domesticated host, Experimental settings, Wild hostAquatic: All Stages
Salvelinus leucomaenis leucomaenis (whitespotted char)Experimental settingsAquatic: All Stages
Salvelinus malma
Salvelinus namaycush (lake trout)Domesticated host, Experimental settings, Wild hostAquatic: All Stages
Scardinius erythrophthalmus (rudd)
Sebastes schlegelii
Semotilus atromaculatusExperimental settings
Silurus glanis (wels catfish)Experimental settings
Sparus aurata (gilthead seabream)Experimental settings
Thaleichthys pacificus
Thymallus thymallus (grayling)Aquatic: Adult|Aquatic/Fry
Tinca tinca (tench)Wild host

Hosts/Species Affected

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Aeromonas salmonicida was traditionally thought of as a pathogen of salmonids, although this may be due in part to the volume of research carried out on this group. Salmonids are usually the most valuable species in developed countries, where good diagnostic bacteriological facilities are available (McCarthy and Roberts, 1980). It is now recognized that the host range of A. salmonicida is wide and that furunculosis is only one of several clinical diseases associated with A. salmonicida. Furthermore, infections of fish with A. salmonicida are not necessarily associated with clinical manifestations and may remain covert (Hiney et al., 1997b). Typical A. salmonicida have been associated with clinical or covert disease in a variety of salmonid and non-salmonid species in fresh water, brackish water and sea water.

Species susceptibility to typical Aeromonas salmonicida (furunculosis)

Table 1 Salmonid fish species from which typical Aeromonas salmonicida has been isolated (after Bernoth, 1997a).


Fish species

*Apparently healthy fish not showing signs of infection (see Hiney et al., 1997b).

†Not specified whether typical or atypical A. salmonicida but assumed to be typical.

 


Table 2 Freshwater non-salmonid species from which typical Aeromonas salmonicida have been isolated (after Bernoth, 1997a).


Fish species

 
Common nameScientific nameReference
American eelAnguilla rostrataClinicalNoga and Berkhoff (1990)
Brassy minnowHybognathus hankinsoniUnclear*McFadden (1970)
Brook sticklebackCulaea inconstansUnclearMcFadden (1970)
CarpCyprinus carpioClinicalMackie et al. (1930)
  UnclearBernoth (1997b)
CatfishSilurus glanisClinicalMackie et al. (1930)
Chestnut lampreyIchthyomyzon castaneumUnclearBernoth (1997a)
Common shinerNotropis cornutusClinicalOstland et al. (1987)
Creek chubSemotilus atromaculatusClinicalOstland et al. (1987)
  UnclearMcFadden (1970)
European eelAnguilla anguillaIncidentalSlack (1937)
Fathead minnowPimephales promelasClinicalMcFadden (1970)
GobyCottus gobioClinicalBernoth (1997b)
Golden shinerNotemigonus crysoleucasClinicalOstland et al. (1987)
GroperRoccus mississippiensisUnclearHerman (1968)
LampreyNot specifiedNot specified†McCarthy (1975)
MinnowPhoxinus phoxinusUnclearBernoth (1997a)
Mottled sculpinCottus bairdiIncidentalRabb and McDermott (1962)
Non-salmonids IncidentalBragg (1991)
Northern pikeEsox luciusClinicalBernoth (1997a)
Ornamental cyprinids UnclearBernoth (1997a)
PaddlefishPolyodon spathulaClinicalFord et al. (1994)
Redbelly daceChrosomus eos (Phoxinus eos)ClinicalMcFadden (1970)
Smallmouth bassMicropterus dolomieuiClinicalLe Tendre et al. (1972)
SticklebackGasterosteus aculeatusUnclearBarker and Kehoe (1995)
TenchTinca tincaClinicalMackie et al. (1930)
  Covert†Bernoth and Körting (1992)
White suckerCatostomus commersoniClinicalOstland et al. (1987)
Yellow bassMorone mississippiensisClinicalBuckley (1969)
Yellow perchPerca flavescensUnclearMcFadden (1970)

*Unclear from history whether isolation was from a clinical case or was an incidental finding.

†Not specified whether typical or atypical A. salmonicida but assumed to be typical.

 


Table 3 Marine non-salmonid species from which typical Aeromonas salmonicida have been isolated (after Bernoth, 1997a).


Fish species

 
Common nameScientific nameReference
Atlantic codGadus morhuaIncidentalWillumsen (1990)
CoalfishPollachius virensIncidentalWillumsen (1990)
Cuckoo wrasseLabrus bimaculatusClinicalTreasurer and Cox (1991)
Goldsinny wrasseCtenolabrus rupestrisClinicalTreasurer and Laidler (1994)
Rock cookCentrolabrus exoletusClinicalTreasurer and Laidler (1994)
Sea breamSparus aurataClinicalReal et al. (1994)
Striped trumpeterLatris lineataIncidentalBernoth (1997a)
Surf smeltThallichthys pacificusUnclear*Schiewe et al. (1988)
TurbotPsetta maximaClinicalNougayrede et al. (1990)
 Scophthalmus maximusClinicalToranzo and Barja (1992)
WrasseLabridaeClinicalTreasurer and Cox (1991)

*Unclear from history whether isolation was from a clinical case or was an incidental finding.

Many fish species would appear to be susceptible to infections by A. salmonicida subsp. salmonicida, but the level of susceptibility is variable. For example, among salmonids, susceptibility to infection is reported to be low in rainbow trout (Cipriano and Heartwell, 1986; Pérez et al., 1996), while brook trout, brown trout and many other salmon species appear to have a high susceptibility (McCraw, 1952; Evelyn, 1971; Klontz and Wood, 1972; Miyazaki and Kubota, 1975; McCarthy, 1977a; Cipriano and Heartwell, 1986; Austin and McIntosh, 1988). In addition, susceptibility may vary within the same fish species raised from different genetic lines (Dahle et al., 1996; Marsden et al., 1996) or with different histories of exposure to A. salmonicida (St Jean, 1992). Because of the potentially inheritable nature of some disease resistance, directed breeding programmes aimed at raising stocks inherently resistant to furunculosis have been investigated as a possible disease control strategy in salmonids (Gjedrem et al., 1991; Lund et al., 1995; Gjedrem, 1997) and non-salmonids (Sövényi et al., 1988; Hjeltnes et al., 1995). However, the multifactorial nature of inheritable characteristics complicates selective breeding programmes and much work remains to be done in this area (Gjedrem, 1997). Species susceptibility to infection by atypical A. salmonicida is discussed in a later section.

Among salmonids, susceptibility to furunculosis may also be age-related. Many early workers in furunculosis research believed that, in wild salmonid populations, furunculosis was mainly a disease of older fish (Plehn, 1911; Mettam, 1915; McCraw, 1952). Although this perception may, in part, have been due to the easier observation of large carcasses in rivers, experimental evidence did suggest that young fish (under 1 year old) are relatively resistant to A. salmonicida infections (Blake and Clarke, 1931; Mackie and Menzies, 1938; Scallan, 1983). The mechanisms of resistance in young fish are essentially unknown but are probably non-specific (Krantz and Heist, 1970). Furthermore, not all workers agree that age plays a significant part in susceptibility to furunculosis (McCarthy, 1977a; Inglis et al., 1993). McCarthy and Roberts (1980), referring to the disease in fingerlings, observed that fish of this size contract an acute form of the disease, which results in rapid death with little more than slight exophthalmos. Mortalities in infected fish in the 0+ age group can be high and have been reported to reach 93% to 40% during the egg to smolt stages (St Jean, 1992). In the experience of these authors, furunculosis can occur in Atlantic salmon alevin whose yolk sacs are still attached.

Amongst non-salmonid species, disease caused by typical A. salmonicida have been reported in turbot (Pedersen and Larsen 1996), wolffish and cod (Lillehaug et al., 2003) and wrasse (Treasurer and Laidler 1994).

Seasonal variation in the incidence of infection

There are predisposing factors, other than age and inherent susceptibility, that are associated with the precipitation of clinical furunculosis in hatchery stocks throughout the year. These include physical damage, poor water quality, presence of ectoparasites and other diseases, diet and physical and psychological stresses such as grading, tagging, injection and netting (Olivier, 1997; Pickering, 1997). However, a marked seasonality in the incidence of both clinical and covert furunculosis infections has been observed in hatchery stocks and wild populations. In hatchery stocks, both smolting and high water temperatures have been implicated in this apparent seasonality. The period of smolting is associated with major physiological changes, including chronic cortisol elevation, which can bring about severe depression of the fish’s defence system and increased susceptibility to bacterial infections (Maule et al., 1987; Pickering, 1997). In addition, high water temperatures (12-15°C) in late spring-early summer increase the likelihood of furunculosis outbreaks in both fresh water and sea water (Klontz and Wood, 1972; Johansson, 1977; Novotny, 1978; Lillehaug et al., 2003). In fact, Malnar et al. (1988) would contend that high temperature is the major factor influencing the development of furunculosis. High water temperature not only influences the stress response of fish but may act at the level of the pathogen (Groberg et al., 1978), and it has been demonstrated that the growth of A. salmonicida in the blood of cherry salmon (Oncorhynchus masou) correlated positively with water temperatures in the range 5-20°C (Sako and Hara, 1981). Not surprisingly, the seasonal nature of clinical infection by A. salmonicida is not confined to hatchery stocks. As early as 1926, Horne observed that the incidence of furunculosis in a riverine population of brown trout first appeared towards the end of May and declined in October (Horne, 1928). Blake and Clarke (1931) observed that spawning in salmon rendered them susceptible to furunculosis. There is no reason to suspect that the temperature effects observed in hatchery reared stocks will not also apply to wild stocks. The influence of spawning on increased susceptibility to furunculosis would appear to be similar in a wide range of salmonids (Nomura et al., 1993) and includes chronic cortisol elevation and associated lymphocytopaenia (Pickering, 1986), decline in antibody production (Yamaguchi et al., 1980) and immunosuppression associated with gonadal steroids (Slater and Schreck, 1993). The presence of wild spawning fish in the vicinity of freshwater hatcheries may also have an impact on the seasonality of furunculosis in stocks contained within those hatcheries. The occurrence of covert A. salmonicida infections in hatchery populations has also been observed to be seasonal (Jensen and Larsen, 1980; Scallan and Smith, 1984, 1993; Hiney, 1994). However, neither smolting, spawning nor high water temperatures can explain other peaks in the incidence of clinical and covert infections observed by these authors at a number of freshwater hatcheries and supported by anecdotal evidence from the industry. Scallan (1983) suggested that these peaks may result from the stress induced in fish by both high water temperatures and rapidly changing water temperature. As a general rule, both clinical and covert furunculosis are more likely to occur in smolting and spawning fish with the onset of higher water temperatures in spring or during periods of rapid temperature change. However, it is important to keep in mind that furunculosis outbreaks can also occur in very young fish (alevin and fry) and at temperatures as low as 2-4°C (Drinan, 1985).

Species susceptibility to atypical Aeromonas salmonicida


Table 10 Salmonid fish species from which atypical Aeromonas salmonicida have been isolated (after Bernoth, 1997a).


Fish species

   
Common nameScientific nameHabitatHistory of isolationReference
Atlantic salmonSalmo salarFresh waterClinicalGroman et al. (1992)
   IncidentalBenediktsdóttir and Helgason (1990)
  Brackish waterClinical Harmon et al. (1991)
  Sea waterClinicalOlivier (1992)
   IncidentalBenediktsdóttir and Helgason (1990)
Arctic charSalvelinus alpinusFresh waterClinicalWichardt et al. (1989)
  Sea waterClinicalOlivier (1992)
Brook troutSalvelinus fontinalisFresh waterClinicalLjungberg and Johansson (1977)
Brown troutSalmo trutta m. lacustrisFresh waterClinicalRintamäki and Valtonen (1991)
Chum salmonOncorhynchus ketaSea waterClinicalEvelyn (1971)
Coho salmonOncorhynchus kisutchFresh waterClinicalChapman et al. (1991)
GraylingThymallus thymallusFresh waterClinicalRintamäki and Valtonen (1991)
Lake troutSalvelinus namaycushFresh waterCliinicalLjungberg and Johansson (1977)
Masu salmonOncorhynchus masouFresh waterClinicalKimura (1969)
Pink salmonOncorhynchus gorbuschaFresh waterClinicalKimura (1969)
Rainbow troutOncorhynchus mykissFresh waterClinicalWichardt et al. (1989)
Sea troutSalmo trutta m. truttaSea waterClinicalKrovacek et al. (1987)
   IncidentalRintamäki and Valtonen (1991)
Sockeye salmonOncorhynchus nerkaSea waterClinicalEvelyn (1971)


 

Table 11 Freshwater non-salmonid species from which atypical Aeromonas salmonicida have been isolated (after Bernoth, 1997a).


Fish species

  
Common nameScientific nameHistory of isolationReference
American eelAnguilla rostrataClinicalOlivier (1992)
BreamAbramis bramaClinicalMcCarthy and Roberts (1980)
BigheadAristichthys nobilisClinicalCsaba and Szakolczai (1991)
CarpCyprinus carpioClinicalCsaba et al. (1984)
  UnclearBernoth (1997b)
ChubLeuciscus cephalusClinicalWilson and Holliman (1994)
European carpNot givenNot specifiedChart et al. (1984)
GoldfishCarassius auratusClinicalElliot and Shotts (1980)
Japanese eelAnguilla japonicaClinicalKitao et al. (1984)
MinnowPhoxinus phoxinusClinicalHåstein et al. (1978)
Northern pikeEsox luciusClinicalWiklund (1990)
  UnclearWichardt et al. (1989)
Ornamental cyprinids UnclearBernoth (1997a)
PerchPerca fluviatilisClinicalBernoth (1997a)
  UnclearWichardt et al. (1989)
River bleakAlburnus alburnusClinicalBernoth (1997a)
RoachRutilus rutilusClinicalAustin (1993)
  Unclear*Wichardt et al. (1989)
RuddScardinius erythrophthalmusUnclearBarker and Kehoe (1995)
Silver carpHypophthalmichthys molitrixClinicalCsaba and Szakolczai (1991)
Silver breamBlicca bjoerknaClinicalMcCarthy (1975)
Silver perchBidyanus bidyanusClinicalWhittington et al. (1995)

*Unclear from history whether isolation was from a clinical case or was an incidental finding.

 


Table 12 Marine non-salmonid species from which atypical Aeromonas salmonicida have been isolated (after Bernoth, 1997a)


Fish species

  
Common nameScientific nameHistory of isolationReference
Atlantic codGadus morhuaClinicalCornick et al. (1984)
  IncidentalOliver (1992)
American plaiceHippoglossoides platessoidesClinicalOlivier (1992)
Black rockfishSebastes schlegeliClinicalIzumikawa and Ueki (1997)
Common wolffishAnarhichas lupusClinicalHellberg et al. (1996)
DabLimanda limandaClinicalWiklund and Dalsgaard (1995)
FlounderPlatichthys flesusClinicalWiklund et al. (1994)
Goldsinny wrasseCtenolabrus rupestrisIncidentalFrerichs et al. (1992)
Greenback flounderRhombosolea tapirinaClinicalWhittington et al. (1995)
  IncidentalBernoth (1997a)
GreenlingHexogrammos otakiiClinicalIida et al. (1997)
HaddockMelanogrammus aeglefinusClinicalOlivier (1992)
Japanese flounderParalichthys olivaceusClinicalIida et al. (1997)
Pacific herringClupea harengus pallasiClinicalTraxler and Bell (1988)
PlaicePleuronectes platessaClinicalWiklund and Dalsgaard (1995)
SablefishAnoplopoma fimbriaClinicalEvelyn (1971)
Sand eelsAmmodytes lanceaClinicalDalsgaard and Paulsen (1986)
 Hyperoplus lanceolatusClinicalDalsgaard and Paulsen (1986)
Shotted halibutEopsetta grigorjewiClinicalNakatsugawa (1994)
Tom codGadus microgadusClinicalOlivier (1992)
TurbotScophthalmus maximusClinicalPedersen et al. (1994)
WrasseCtenolabrus rupestrisCovert*Frerichs et al. (1992)

*Apparently healthy fish not showing signs of infection (see Hiney et al., 1997b)

The number of hosts from which atypical A. salmonicida isolates have been cultured is rapidly increasing and includes several species of salmonids and non-salmonids, wild or cultured, in fresh, brackish or salt water. Strains of atypical A. salmonicida have been reported in wild and farmed populations of salmonids in northern Europe (Wichardt et al., 1989; Håstein and Lindstad, 1991; Rintamäki and Valtonen, 1991; Hirvelä-Koski et al., 1994; Pedersen et al., 1994; Gudmundsdóttir et al., 1995). The most common clinical sign in these infections is skin ulceration (Wichardt et al., 1989), but a number of other clinical signs have been recognized, some of which are similar to those seen in cases of typical furunculosis (Rintamäki and Valtonen, 1991).

Mortality of up to 60% has been reported from cultured sea trout in Sweden (Wichardt et al., 1989) and 15-20% in Finland and Iceland (Gudmundsdóttir et al., 1995: Rintamäki and Valtonen, 1991). In North America, an ulcer disease of trout, originally described by Sniezko et al. (1950), was associated with a fastidious organism (Haemophilus piscium), which was subsequently reclassified as an atypical strain of A. salmonicida (Paterson et al., 1980; Trust et al., 1980b; Adams and Thompson, 1990). On the Atlantic coast of Canada, fastidious atypical strains have been isolated from Atlantic salmon (Paterson et al., 1980; Groman et al., 1992; Olivier, 1992) with mortality of up to 50% being recorded in some instances (Groman et al., 1992). Thus, disease of atypical A. salmonicida aetiology can have a significant economic impact, depending on the fish species infected.

Some of the best-described conditions caused by strains of atypical A. salmonicida in non-salmonid freshwater hosts are: carp erythrodermatitis (Bootsma et al., 1977), the ulcer disease of goldfish Carassius auratus L. (Shotts et al., 1980) and the ‘head ulcer disease’ of cultured Japanese eels, Anguilla japonica (Ohtsuka et al., 1984; Kitao et al., 1985). Atypical strains of A. salmonicida have also been isolated from strictly marine hosts. Evelyn (1971) isolated an atypical strain of A. salmonicida from a marine species, the sablefish, Anoplopoma fimbria. A similar strain was isolated from lingcod, Ophiodon elongatus, in 1986 and from sablefish in 1987 and 1990 (Bell et al., 1990; McCormick et al., 1990; T.P.T. Evelyn, Nanaimo, 1990 personal communication). Atypical strains have been isolated from other marine species, some of which are now in the process of being developed for aquaculture purposes e.g. wolffish (Grontvedt et al., 2004), halibut (Gudmundsdottir et al., 2003) and cod (Magnadottir et al., 2002). According to some authors, disease conditions of atypical A. salmonicida aetiology may become a restricting factor for the mass culture of these species (Pedersen et al., 1994). In several cases, the disease has been diagnosed in wild fish transferred and maintained in aquarium facilities, reinforcing the hypothesis that there could be strains of A. salmonicida of marine origin (Cornick et al., 1984: Dalsgaard and Paulsen, 1986; Harmon et al., 1991; Olivier, 1992; Whittington et al., 1995). Furthermore, these findings indicate that marine species may act as carriers or reservoirs of some atypical strains and the disease condition will be expressed when animals are stressed.

Distribution

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At present, the geographical distribution of A. salmonicida subsp. salmonicida is almost worldwide, including Japan (Miyazaki and Kubota, 1975) and the mainland of Asia (Inglis et al., 1993), from where it was previously considered to be absent (Fryer et al., 1988). The possible exceptions to this distribution are South America and New Zealand, from which reports of the isolation of A. salmonicida have yet to be made (Bernoth, 1997a). The introduction of furunculosis into Sweden in 1951 was reported by Wichardt et al. (1989) and it was recognized in Norway in 1964 (Lunder and Håstein, 1990; Johnsen and Jensen, 1994). This more recent identification of A. salmonicida in Scandinavian countries has been tentatively traced to importation of live fish stocks, initially from other European countries (Egidius, 1987; Wichardt et al., 1989) and then within Scandinavia (Rintamäki and Valtonen, 1991). To date, there have been no reports of ‘typical’ furunculosis in salmonids in Australia, despite many attempts to isolate the organism (Bernoth, 1997a). Atypical A. salmonicida was, however, identified from diseased goldfish (Trust et al., 1980b). In South Africa, the first incident of infection of rainbow trout by an atypical A. salmonicida was noted by Boomker et al. (1984).

The widespread distribution of furunculosis is reflected by the fact that the disease has never been listed by the Office International des Epizooties (OIE) as one that merits special attention, being considered endemic in most countries and capable of control (C. Michel, personal communication). Likewise, the European Community assigned furunculosis to its List 3 diseases, that is, diseases which are endemic in many member states (Council Directive 91/67/ EEC). Individual member states may enforce control strategies on importation of stocks only with the approval of the Standing Veterinary Committee, which must ensure that valid reasons exist for the proposed controls, such as a disease-free status, and that they are not a concealed trade barrier (McLoughlin, 1993).

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.

Last updated: 10 Jan 2020
Continent/Country/Region Distribution Last Reported Origin First Reported Invasive Reference Notes

Asia

JapanPresentCABI (Undated)Original citation: Miyazaki and Kubota (1975)

Europe

NorwayPresentCABI (Undated);
SwedenPresentCABI (Undated)Original citation: Wichardt and et al. (1989)

Pathology

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Despite the significant amount of work that has been undertaken on the pathogenicity of A. salmonicida and the advances that have been made in this area, the mechanisms whereby this organism produces disease are only partly understood. Virulence mechanisms fall broadly into two categories, these being cell-surface structures and extracellular products (ECPs) excreted by the cell.

Cell-surface structures

In common with many pathogenic bacteria, A. salmonicida can manifest an additional surface protein microcapsule (Kay and Trust, 1997). These crystalline surface protein arrays are generally referred to as S-layers, but, for historical reasons, in A. salmonicida the term A-layer is more commonly used (Udey and Fryer, 1978). SDS-polyacrylamide gel electrophoresis (PAGE) analysis and X-ray diffraction studies have shown the A-layer of A. salmonicida to be a protein of approximately 50 kDa, with a tetragonal structural arrangement (Trust et al., 1980a; Kay et al., 1981; Evenberg et al., 1982; Garduño and Kay, 1992a; Kay and Trust, 1997). A-layers isolated from a wide variety of A. salmonicida strains were shown to be immunologically conserved (Kay et al., 1984). The evidence implicating the A-layer of A. salmonicida as a primary virulence factor is very strong. Authors have demonstrated that typical strains possessing the A-layer are both virulent for susceptible fish species and autoaggregating, while A-layer-negative variants are non-virulent and non-aggregating (Udey and Fryer, 1978; Ishiguro et al., 1981; Kay et al., 1984; Cipriano and Bertolini, 1988; Cipriano and Blanch, 1989; Noonan and Trust, 1995). Kay and Trust (1997) have suggested that the ability of the A-layer to bind immunoglobulins and other extracellular proteins may result in the masking of bacterial immunogenic receptors, thus allowing A. salmonicida to evade the host’s immune response. The A-layer has been reported to promote bacterial penetration and adhesion and to inhibit complement-mediated lysis in host serum (Trust et al., 1983; Sakai and Kimura, 1985; Garduño and Kay, 1992b; Garduño et al., 1995). It was also reported that the net negative charge of A-layer-containing A. salmonicida cells plays a crucial role in their long-term survival in a freshwater microcosm (Sakai, 1986) and that the production of exopolysaccharides under low nutrient conditions, which may protect the cell from desiccation, was greater in an A-layer-containing strain than in an A-layer-deficient mutant (Bonet et al., 1993).

Udey (1982) demonstrated that the incorporation of the protein-specific dye CBB into common growth media could provide a differentiation between strains containing an intact A-layer (A+), which grew as blue colonies, and those lacking it (A-), which grew as white colonies. Using CBB-containing media, Cipriano and Bertolini (1988) demonstrated a correlation between the A+ phenotype on this medium and virulence for brook trout, although Bernoth (1990a) reported that colony colour on CBB-containing media did not always correlate with virulence of A. salmonicida. Anomalies have been reported which might challenge the importance of the A-layer as a virulence factor. Specifically, one A- A. salmonicida strain was reported to be virulent for rainbow trout (Bernoth, 1990a), while Olivier (1990) reported that the presence of an intact A-layer did not always correlate with virulence. It must be remembered, however, that A. salmonicida strains lacking an A-layer are laboratory artefacts. To our knowledge, no strain lacking an intact A-layer has ever been isolated from a natural furunculosis infection.

The other major component of the cell surface of A. salmonicida, in common with all Gram-negative cells, is LPS. In A. salmonicida, LPS is normally composed of two types, a low-molecular-weight lipo-oligosaccharide (LOS), situated beneath the A-layer, and a high-molecular-weight LPS, containing attached O-polysaccharide chains, some of which traverse the A-layer (Ishiguro et al., 1983; Chart et al., 1984; Evenberg et al., 1985). The role of LPS in the structure of the A-layer and the virulence of A. salmonicida has been elusive. Observations of O-polysaccharide-deficient mutants have indicated that they play a role in securing the A-layer to the cell surface (Belland and Trust, 1985; Griffiths and Lynch, 1990), and Cipriano and Blanch (1989) reported that only A. salmonicida strains containing both an intact A-layer and LPS were virulent for brook trout.

It must be remembered, however, that virtually all of the studies cited above were carried out on A. salmonicida strains grown in vitro on artificial laboratory media. It is unlikely that these conditions will reflect the behaviour of A. salmonicida occurring naturally either in a fish host or in the environment. Garduño et al. (1993) found that cells grown in vivo in diffusion chambers which had been implanted in rainbow trout peritoneal cavities displayed enhanced resistance to host-mediated serum and oxidative killing, and expressed a polysaccharide capsular layer, which they suggested might act as a mechanism of phagocytosis resistance or evasion of the host immune system. Thornton et al. (1993) have also reported that A. salmonicida cells grown in vivo expressed novel surface antigens. It is reasonable to postulate that these additional cell surface components play an important role in the virulence of A. salmonicida, although this has not, as yet, been adequately demonstrated. Further in vivo studies will be required to clarify the exact role and importance of cell surface components in the pathogenicity of A. salmonicida.

A cytotoxin has been reported to be associated with the cell surface and termed AexT (Aeromonas salmonicida exoenzyme toxin) and is expressed following contact with host cells (Braun et al., 2002). It is secreted by a Type III secretion pathway directly into the cytosol of host cells (Burr et al., 2003). The Type III secretion pathway is encoded by genes carried on a large plasmid which is lost when the cells are exposed to high temperature (25°C) and this is accompanied by loss of virulence for cultured fish cells (Stuber et al., 2003).

Genes encoding pili have been reported and mutants deficient in their expression have reduced virulence (Masada et al., 2002).

As mentioned above, in vivo grown bacteria were more resistant to superoxide and hydrogen peroxide challenge but this acquired property was not dependent on capsule formation (Garduno et al., 1993). Of relevance to this, is that when A. salmonicida are grown in vitro under iron-restricted conditions, the bacterium produces manganese-superoxide dismutase (MnSOD) which is located in the periplasm, just below the cell surface. The expression of this enzyme is associated with increased resistance to superoxide challenge (Barnes et al., 1996) and as in vivo conditions are an iron-restricted environment, it is likely that the expression of MnSOD is also induced in vivo. Furthermore, levels of SOD production appear to be higher in virulent than in avirulent strains (Dacanay et al., 2003).

Extracellular products

In common with other pathogenic organisms, A. salmonicida has been found to produce a number of ECPs many of which have enzyme activity. Injection of crude extracellular material of A. salmonicida has been clearly demonstrated to kill susceptible fish (Munro et al., 1980; Ellis et al., 1981; Ellis, 1991) and a considerable amount of work has been performed to analyse the constituents of ECPs and to understand their role in virulence and pathogenesis. Ellis (1997a) has provided an excellent review of the known ECPs of A. salmonicida, which together make for a long list. These are grouped by Ellis (1997a) into three types, namely proteases, membrane-damaging toxins and other toxins, including Hlysin, which have not yet been fully investigated.

Tajima et al. (1983) were the first to report a lethal 70 kDa protease in A. salmonicida ECP, which was found to have an LD50 of 2.4 µg g-1 when injected into young salmon and to produce haemorrhaging and muscle liquefaction (Lee and Ellis, 1989). These effects were not as severe as when total ECP was used, but equivalent lesions were produced when the protease was combined with a haemolytic factor in the ECP, suggesting an interaction between these components (Lee and Ellis, 1991b). It is believed that the protease toxin is produced by A. salmonicida to digest host proteins as a nutrient source, although it has been found to have only limited specificity, mainly towards proteins with a relatively open structure (Price et al., 1990). The 70 kDa protease has also been shown to reduce the clotting time of trout blood, which may account for the presence of microthrombi in fish tissue in cases of clinical furunculosis and following injection of crude toxins (Ellis et al., 1988). There is, however, conflicting evidence for the exact role of the 70 kDa protease in virulence. A lack of correlation between the amount of protease in ECP and the killing ability of that ECP has been reported (Drinan et al., 1989), and Sakai (1985) reported that a protease-deficient mutant of A. salmonicida was avirulent, while other workers have identified virulent isolates which do not produce any protease under standard culture conditions (Hackett et al., 1984; Ellis et al., 1988). However, this confusion may be an artefact of in vitro studies, because apparent protease-deficient strains have been found to produce protease in vivo (Ellis, 1991). A second protease, whose preferred substrates are gelatine and collagen, has been identified (Sheeran and Smith, 1981), but its physiochemical properties have not yet been determined in detail.

A number of membrane-damaging activities of ECP have been described. Present evidence suggests that, of these membrane-damaging toxins, glycerophospholipid-cholesterol acyltransferase (GCAT) complexed with LPS, so called GCAT/LPS, is the most important factor in the lethal toxicity and pathology of the ECP (Ellis, 1997a). The LD50 of purified GCAT/LPS for Atlantic salmon has been reported to be 45 ng g-1 body weight (Lee and Ellis, 1990). In vitro studies of GCAT/LPS have shown it to be leucocytolytic, cytolytic and highly haemolytic for salmonid erythrocytes, although this haemolysis was incomplete in the absence of the 70 kDa protease (Lee and Ellis, 1990), which has been suggested by Ellis (1997b) to be necessary for activation of GCAT activity. However, there is no evidence for in vivo haemolysis in clinical furunculosis (Lee and Ellis, 1991a) and Ellis (1997a) has suggested that in vivo GCAT/LPS may function in destabilization of host red cell membranes rather than active haemolysis. The histopathological effects of GCAT/LPS are not extensive and cannot account for the death, within 20 h, of fish injected with purified product, and it has been suggested that in vivo toxicity may be due to metabolic effects, although this has yet to be confirmed (Ellis, 1997a). Although the 70 kDa protease and GCAT/LPS are clearly important ECPs, it has been demonstrated that mutants deficient in the production of either the protease or the GCAT were still virulent and could produce the manifestations of classical furunculosis (Vipond et al., 1998). Therefore, it has become clear, with respect to the pathogenesis of furunculosis and the lethal toxicity of the exotoxins, that, instead of one toxin being critical for pathogenesis, a combination of ECPs act in concert to produce disease (Barnes and Ellis 2004).

Host response to infection

Efforts over the last 10 years to improve the effectiveness of vaccines against furunculosis have led to a much improved understanding of the salmonid immune system, and a number of comprehensive reviews have been published (Warr and Cohen, 1991; Secombes, 1994a; Secombes and Olivier, 1997). In general, most components of the fish immune system are analogous to those of higher vertebrates. These include physical barriers and chemical barriers to prevent infection, inducible but non-specific humoral factors, phagocytes and non-specific cytotoxic cells, which mediate an inflammatory response, and, finally, specific immunity, effected by lymphocytes. It is this last type that is responsible for ‘immunological memory’, ensuring that responses to a second exposure are faster and stronger than the initial response, thus conferring immunity (Secombes and Olivier, 1997). Specific immune responses include both humoral immunity, based on the production of antibodies by B cells, and cellular immunity, based on the production of activated macrophages with enhanced bacterial activity, following release of cytokines from T cells (Secombes and Olivier, 1997).

Non-specific humoral factors

Salmonids present physical barriers to infection by A. salmonicida in the structure of their skin and intestinal mucosa and there is some evidence for active immune cells on both of these surfaces (Ellis, 1985; Cipriano, 1986). Fish also have a variety of non-specific humoral factors with which to counter infection by A. salmonicida (Lund et al., 1991). These include enzyme systems to lyse bacteria, such as lysozyme and complement, antiproteases to neutralize bacterial proteases, proinflammatory molecules, which can attract leucocytes and neutrophils to an infection site and increase local capillary permeability, substances to sequester essential nutrients and thus limit bacterial growth, and molecules which bind to the bacterial cell and trigger the complement system and thus facilitate phagocyte uptake (Secombes and Olivier, 1997; Ellis, 1999; Ellis, 2001). In addition to direct bactericidal effects, some components of the non-specific humoral response serve to neutralize toxins excreted by A. salmonicida, for example, alpha 2- macroglobulin, which has antiprotease activity (Salte et al., 1992). There is also evidence that the complement system may have a role in neutralizing A. salmonicida ECP (Sakai, 1984).

Non-specific cellular factors

Should A. salmonicida successfully breach the non-specific immune defences of fish, it will encounter specialized cells of the host immune system, namely the phagocytic cells. These include monocytes, or macrophages, and granulocytes (neutrophils, eosinophils and basophils). The macrophages of fish are similar to those of higher animals in both morphology and function and they act as antigen-presenting cells, cytokine-secreting cells and effector cells, with the ability to actively phagocytose bacterial cells (Secombes and Fletcher, 1992; Secombes and Olivier, 1997). Less is known about the function of neutrophils in the immune reaction of salmonids. Lamas and Ellis (1994a) demonstrated that neutrophils isolated from Atlantic salmon migrated in fish serum in response to the presence of A. salmonicida, although, strangely, A-layer-deficient strains acted as stronger chemoattractants. Neutrophils were also found to be phagocytic, but phagocytosis was low compared with macrophages (Lamas and Ellis, 1994b).

Eosinophilic granular cells (EGCs) may play an important role in the host response against infection by A. salmonicida, as they are widely distributed in connective tissues, especially in the intestine and gills (Ellis, 1985). In the intestine of salmonids, EGCs form a layer called the stratum granulosum between the muscle layer and the stratum compactum. A number of studies have suggested that fish EGC are the equivalent of mammalian mast cells (Ellis, 1982; Powell et al., 1991; Reite 1998), and in response to i.p. injection of A. salmonicida ECP, there is a rapid and explosive degranulation of the intestinal EGC accompanied by intense vasodilatation (Ellis, 1985; Vallejo and Ellis, 1989).

Specific cellular responses

The specific immune response of salmonids includes three strategies, that is, lymphocyte proliferation, antibody production and cytokine response. In higher vertebrates, lymphocyte responses are characterized by their ability to proliferate following contact with bacterial antigens, generating a clone of antigen-specific cells. Some of these cells give rise to the primary response, while others remain dormant as specific memory cells, able to proliferate in response to subsequent contact with the same antigen (Secombes and Olivier, 1997). There are relatively few studies on lymphocyte proliferation in salmonids in response to A. salmonicida infection. However, those that have been carried out suggest that lymphocyte proliferation does occur in response to both formalin-killed and live A. salmonicida cells (Reitan and Thuvander, 1991; Vaughan et al., 1993), although this response is dependent on the dose and type of antigen presented (Erdal and Reitan, 1992; Secombes and Olivier, 1997).

Antibody production to A. salmonicida, on the other hand, has been extensively studied in salmonids, particularly following immunization with killed whole cells or ECP. High serum antibody titres can be routinely elicited in salmonids by both i.p. and i.m. injection of A. salmonicida (Secombes and Olivier, 1997). At permissible temperatures (10-15°C) antibody titres rise within 3-4 weeks and peak within 8-12 weeks, although water temperature is critical to this response and at lower temperatures responses may be slower or absent (Ellis et al., 1992; Eggset et al., 1997). The sites of antibody production in salmonids would primarily seem to be spleen and head kidney (Reitan and Thuvander, 1991; Davidson et al., 1993). Antibody-secreting cells have also been found in mucosal sites, such as the gut, but these can take up to 7 weeks post-vaccination to appear (Davidson et al., 1993). Antibody responses have also been elicited following administration of A. salmonicida by subcutaneous injection (Anderson, 1969), by bath (Anderson et al., 1979) and orally (Davidson et al., 1993). The inclusion of an adjuvant, such as an oil or glucan, in vaccines administered by a parenteral route has been shown to enhance the antibody titre and may act as a depot of antigen, allowing vaccination at low temperatures (Cipriano and Pyle, 1985; Anderson et al., 1997; Ellis, 1997b; Midtlyng, 1997; Secombes and Olivier, 1997). It should be noted, however, that high antibody titres do not necessarily correlate with protection and it is the specificity of the antibodies that appears to be important (Hirst and Ellis, 1994; Ellis, 1997b). In older fish, proliferation studies suggest that specific B-cell memory can be established to A. salmonicida and that a second exposure to the organism results in a faster and stronger antibody response (Secombes and Olivier, 1997).

In fish the regulatory role of T cells in the immune response is thought to be mediated by released cytokines analogous to interleukin-2, chemokines and macrophage-activating factor (MAF), following exposure to specific antigens (Secombes et al., 2001). Although little is generally known about cytokine release in response to A. salmonicida, release of MAF has been demonstrated in vitro from cultured cells, removed from fish immunologically primed with killed whole A. salmonicida or ECP, 2-3 weeks post-exposure, peaking 4-5 weeks post-exposure (Marsden et al., 1994). In vitro studies have also shown that MAF-treated macrophages acquired the ability to kill A. salmonicida (Graham et al., 1988). In fish, MAF production has also been demonstrated to correlate with both lymphocyte proliferation and antibody production following vaccination with whole cells (Secombes and Olivier, 1997). Therefore, unlike antibody production, any epitope on A. salmonicida can potentially induce a cell-mediated response, such as MAF release. In common with antibody production, however, cytokine release is temperature dependent and has been shown to be absent in fish cells kept at 7°C or less (Hardie et al., 1994).

Pathogenesis of atypical Aeromonas salmonicida

There is limited information on the pathogenicity of atypical strains of A. salmonicida. They generally produce cutaneous ulceration, but the infection does not necessarily become systemic and the cause of death is not always substantiated (reviewed by Trust, 1986). Work by Bucke (1980) provided evidence that atypical isolates could be pathogenic not only for their original host but for other species as well. In this particular study, Bucke (1980) inoculated i.m. brown trout, perch, rudd, roach, carp and goldfish with three strains of A. salmonicida, including a typical isolate, a cyprinid atypical isolate and a goldfish isolate (unfortunately, the author did not provide the exact number of bacteria injected). His results clearly indicated that there were differences in the virulence of the isolates towards different hosts; for example, the typical isolate was virulent for trout, roach and rudd, while the cyprinid isolate was only virulent for brown trout and the goldfish isolate was virulent for all the species tested.

In some cases, atypical strains isolated from salmonids can be almost as virulent as typical isolates. The atypical strain isolated from Atlantic salmon in Newfoundland, Canada, has an LD50 of less than 50 bacteria for juvenile Atlantic salmon, following an i.p. challenge (Olivier et al., 1990; Olivier, 1992). Other salmonid isolates from Sweden, Norway and Iceland were also virulent for salmonids, with LD50 values of 1 × 102-3 for Atlantic salmon (Olivier et al., 1990), confirming the results of Gudmundsdóttir et al. (1997) with similar isolates from Iceland.

Several atypical strains of A. salmonicida are virulent in their host of origin. The carp isolate (V234/81) produces 100% mortality in carp inoculated between dorsal scales with 1 × 106 cfu (Evenberg et al., 1988). The LD50 of an eel isolate was approximately 1 × 103 cfu by i.m. injection into healthy eels (Ohtsuka et al., 1984); similarly, herring isolates were highly virulent for their host, killing 50% of fish injected with fewer than 100 cfu (Traxler and Bell, 1988). A goldfish isolate is highly virulent for goldfish, with LD50 less than 1 × 104 cfu (Trust et al., 1980b), while a wrasse isolate had a low LD50 (5 × 102) when determined in wrasse by i.p. injection. Other isolates were not found to be as virulent for their host: isolates from cod, flounder, turbot and minnow had LD50 values higher than 1 × 106 (Håstein et al., 1978; Cornick et al., 1984; Wiklund, 1995b).

As salmonid culture represents an important asset in several countries, several non-salmonid isolates have been tested for their possible virulence in salmonids. There is no doubt that one goldfish isolate can be highly virulent for salmonids after i.p. injection, with LD50 values ranging from 1 × 102 to 1 × 103 injected cells. This has been confirmed by other studies, in which goldfish isolates were found to be virulent for Atlantic salmon, brook trout and rainbow trout under different challenge conditions, including i.p. injection and bath challenge, with or without prior skin abrasion, and by cohabitation (Carson and Handlinger, 1988; Whittington and Cullis, 1988). In addition, a sand eel isolate killed 50% of rainbow trout injected i.p. with 8 × 106 cfu (Dalsgaard and Paulsen, 1986). Isolates from other species, including flounder, turbot, wrasse and cod, were not found to be virulent for salmonids (Pedersen et al., 1994). Even if some atypical strains isolated from non-salmonids have been shown to be virulent for salmonids under laboratory challenge conditions, there are few reports of atypical strains of non-salmonid origin causing losses in salmonids in the field. However, in one instance a sablefish isolate was found to infect cultured Pacific salmon, but other virulent non-salmonid isolates, such as the goldfish strain, have not been reported from salmonids to date. Furthermore, there is little evidence of crossover of atypical strains from one non-salmonid species to another, although one goldfish isolate has been found to infect silver perch (Whittington et al., 1995) and the same isolate was found in two hosts, sablefish and lingcod, on the west coast of Canada.

Virulence factors

The virulence factors of atypical A. salmonicida strains are not as well characterized as those of typical strains (Trust, 1986). Atypical strains are able to sequester iron under conditions of iron limitation, but their ability is less than that reported for typical strains and some atypical strains are able to utilize siderophores produced by typical isolates (Chart and Trust, 1983). According to Hirst et al., (1991, 1994), the iron-uptake mechanism of atypical strains is siderophore-independent, in contrast to typical strains. These results were confirmed when Hirst and Ellis (1996) demonstrated that typical and atypical strains differed in their mechanism of utilization of non-haem protein-bound sources of iron. Typical strains utilize transferrin via a siderophore-mediated mechanism and are also able to digest transferrin with the extracellular serine protease. Atypical strains utilize transferrin by a siderophore-independent mechanism, probably involving the proteolytic degradation of transferrin by the extracellular metalloprotease.

As stated previously, most atypical strains examined so far possess surface structures (A-layer and LPS) similar to those of typical isolates. Small differences have been noted in the electrophoretic mobility of both the A-layer and LPS of atypical strains (Kay et al., 1984; Evenberg et al., 1985; Griffiths and Lynch, 1990). In the only report where a correlation between surface structure and virulence was tested, Trust et al. (1980a) demonstrated that an i.m. injection of 1 × 104 A-layer-positive cells in goldfish killed all test animals (8/8), whereas a similar injection of 108 cfu of an A-layer-negative isogenic strain killed only one goldfish. Except for these results, the role of surface structure in the pathogenicity of atypical strains has not been fully assessed, but it would not be surprising if the important role of the A-layer and additional surface structures as virulence factors were confirmed in atypical isolates.

Extracellular virulence factors produced by atypical strains are poorly understood. Pol et al. (1981) have reported that the ECP of one atypical strain was lethal for carp and similar results were obtained by Evenberg et al. (1988). Filter-sterilized culture supernatants of a carp isolate were lethal for carp by i.p. injection; interestingly, toxicity of the supernatants was dependent on growth conditions. Hastings and Ellis (1985) noted differences in the production of haemolysin and protease between typical and atypical strains, and Gudmundsdóttir et al. (1990) have purified and characterized a toxic protease from an atypical strain of A. salmonicida. The enzyme was caseinolytic and gelatinolytic, possessed properties of a metalloprotease and had a molecular mass of 20 kDa. This protease was only identified in atypical strains (5/9 strains tested) and not in typical isolates and thus may be specific for atypical strains. A further study of 25 atypical strains isolated from a variety of hosts indicates that the proteases produced by atypical strains are variable, and the author was able to differentiate five groups, based on protease properties (Gudmundsdóttir, 1996) and other exotoxins (Gudmundsdóttir et al., 2003). Compared with typical strains which were homogeneous in producing protease and gelatinase, atypical strains were heterogeneous.

The possibility that atypical strains can cause immunosuppression in carp was investigated by Evenberg et al. (1986). Infected carp given a sublethal i.m. injection showed a progressive decrease in total serum protein and immunoglobulin (Ig) levels but differential blood counts did not differ. In carp immunized with sheep erythrocytes, a sublethal infection produced a reduction of plaque-forming cells and serum antibodies against sheep erythrocytes, but the cellular response, tested by skin allograft rejection, was enhanced as the disease progressed (Pourreau et al.,1986). Further studies by Pourreau et al. (1987) indicated that crude supernatants of a virulent atypical strain modulated the carp immune response when tested by mitogen stimulation of carp leucocytes. Stimulation of leucocytes was enhanced by the supernatant from a young culture (20 h) of the virulent strain but was severely inhibited by supernatants from older cultures (96 h). This inhibitory activity was lost after heat treatment, suggesting it was due to a proteinaceous structure, but the substance was not characterized further.

With respect to atypical strains causing ‘furunculosis’ in salmonids, Gudmundsdóttir et al. (1995) have provided evidence that bacterins prepared from an atypical strain isolated from salmonids in Iceland can provide protection against the disease. Further work by Gudmundsdóttir et al. (1997) showed that Atlantic salmon vaccinated with detoxified ECP provided better protection than formalin-killed cells or a mixture of both bacterins. Since passive immunization with rainbow trout or rabbit antiprotease (AsaP1) was demonstrated to be efficacious and since the same antisera, but containing anti-A-layer antibodies, were not protective, the authors conclude that the protection of Atlantic salmon against atypical A. salmonicida is probably due to humoral immunity, with antibodies directed against bacterial toxins contained in the ECP.

Diagnosis

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The traditional description of A. salmonicida subsp. salmonicida is of a nonmotile, non-sporulating, fermentative, Gram-negative, aerobic bacillus (Popoff, 1984), which reduces nitrate, liquefies gelatine, hydrolyses starch (Popoff and Lallier, 1984) and produces cytochrome oxidase, although isolation of an oxidase-negative ‘typical’ A. salmonicida has been reported from coho salmon (Teska et al., 1992). Staining has a tendency to be bipolar, and the organism measures approximately 1.0 µm × 2.0 µm, varying morphologically from an almost coccoid form in freshly isolated cultures to distinct rods in cultures maintained on artificial media, the latter often proving avirulent. Aeromonas salmonicida is normally isolated from the kidney of infected fish, although it can also be isolated from lesions, blood and other organs (Daly and Stevenson, 1985). The size of colonies is variable, ranging from 0.5 to 3.0 mm in diameter after 72 h incubation (Drinan, 1985). Temperatures of 18-22°C are optimal for growth. A. salmonicida grows poorly at 4°C and does not grow at 37°C (Snieszko, 1957), although isolation of an atypical strain capable of growth at 37°C has been reported (Austin, 1993).

On agar media containing tryptone, typical A. salmonicida normally, though not always (Wiklund et al., 1993), produces a brown, melanin-like, water-soluble pigment. This pigment and its production pathway have been described in detail by Donlon et al. (1983). Virulent colonies are small and friable and, on initial isolation from fish, autoagglutinate in 0.85% physiological saline (Evenberg and Lugtenberg, 1982; Evenberg et al., 1982; Drinan, 1985). Prolonged subculture on laboratory media, or incubation of strains above optimum temperatures, produces non-aggregating variants with altered cell morphology (Ishiguro et al., 1981). The hydrophobic nature of A. salmonicida is due to the possession of an additional surface-protein layer (A-layer), first described by Udey and Fryer (1978). The A-layer has been well characterized (Kay and Trust, 1997), because of its association with the pathogenesis of A. salmonicida and its role in resistance to host defence mechanisms (Sakai and Kimura, 1985; Secombes and Olivier, 1997). However, the detection of an intact A-layer in laboratory culture cannot be used, in isolation, as a predictor of the virulence of an isolate (Ellis et al., 1988; Olivier, 1990).

Diagnosis in clinically infected fish

The boil-like lesions observed by Emmerich and Weibel (1894), from which classical furunculosis derives its name, are the exception in clinical cases, rather than the rule (Bernoth, 1997b) and are normally only observed during chronic infections in older fish. Table 4 [in the Disease Course section] summarizes the main clinical signs, gross pathology and histopathological features that have been described for the different forms of typical furunculosis (peracute, acute, subacute/chronic). The gross pathological signs of peracute and acute furunculosis in young fish are often indistinguishable from other bacterial septicaemias on preliminary examination. The inexperienced diagnostician may also have difficulty in differentiating the morphology and Gram-staining behaviour of A. salmonicida in fish tissue, as seen under a microscope, from other Gram-negative bacteria occurring in fish (Bernoth, 1997b). Thus, a firm diagnosis of clinical furunculosis requires isolation of the dominant infecting organism on agar media and identification by morphology, combined with either biochemical or serological tests, as A. salmonicida.

Detection of typical Aeromonas salmonicida during clinical infections

Under ordinary circumstances, typical A. salmonicida can be recovered from clinically diseased fish, especially from the kidney and surface lesions, where present (Austin and Austin, 1993). However, culturing samples from several organs, rather than kidney alone, has been demonstrated to increase the detection rate in fish populations where the incidence of infection is low (Bernoth, 1997b). The recommended diagnosis of A. salmonicida in clinically diseased fish is based on isolation of the organism from the kidney on either tryptone soya agar (TSA) or brain-heart infusion agar (BHIA) (Department of Fisheries and Oceans, 1984; Shotts, 1984). The brown water-soluble pigment produced by typical A. salmonicida on TSA after 2-4 days’ incubation at 20-25°C is used as a presumptive identification. However, caution must be exercised when pigmentation on TSA is used as a presumptive identification of A. salmonicida. Other bacteria have also been found to produce a brown diffusible pigment on TSA, such as mesophilic aeromonads and Pseudomonas fluorescens (McCarthy, 1975; Frerichs and Holliman, 1991). Neither TSA nor BHIA is selective for A. salmonicida, allowing the growth of competing organisms, which may inhibit pigmentation or the growth of A. salmonicida (Austin and Austin, 1993). Inhibition of growth may result from the ability of faster-growing organisms to sequester the available nutrients in the medium (particularly iron) or from the production of inhibitory substances by these competing organisms (Cornick et al., 1969; Michel and Dubois-Darnaudpeys, 1980; Smith and Davey, 1993). Supplementing TSA with 0.01% (w/v) Coomassie brilliant blue (CBB) (CBB agar, CBBA) has been found by these and other authors to aid in the preliminary differentiation of A. salmonicida from competing bacteria (Cipriano and Bertolini, 1988; Markwardt et al., 1989; Cipriano et al., 1992). On this medium, A-layer-positive A. salmonicida colonies stain deep blue to navy and can be easily distinguished, the intensity of staining being dependent on the source of the dye and the batch of TSA. However, CBBA cannot be totally relied upon, because bacteria other than A. salmonicida can produce dark blue colonies on CBBA (Teska and Cipriano, 1993). None the less, the use of CBBA as a primary plating medium reduces the numbers of bacteria that need to be screened to ensure definitive identification.

Occasional failure to isolate typical A. salmonicida from diseased fish with macroscopic and histological signs of furunculosis has been noted. There are a number of reasons why A. salmonicida may fail to yield colonies on solid media, even in the absence of competing bacteria. Firstly, the number of detectable cells present in the original sample may be below the lower detection limit of cultural isolation (Bernoth, 1997b). Attempts to overcome this limitation, by incorporating a pre-enrichment step, carried out in liquid media, prior to plating on solid media, were reported by Daly and Stevenson (1985). Pre-enrichment of kidney samples in tryptone soya broth (TSB) for 48 h more than doubled the A. salmonicida detection rate in a fish population undergoing a furunculosis epizootic. A second reason for lack of growth might be the unsuitability of laboratory media, such as TSA and BHIA, to support the growth of A. salmonicida. Little is known about the specific nutrient requirements of typical A. salmonicida, other than that it requires methionine and arginine (Nerland et al., 1993). Most artificial media have been formulated for the isolation of medically important bacteria and do not, therefore, present an ideal environment for terrestrial and aquatic organisms. Another potential problem with the use of TSA as the primary isolation medium is that some batches may occasionally fail to support the growth of typical A. salmonicida which will grow on BHIA (Power et al., 1987) or blood agar (Bernoth and Artz, 1989). To exclude this possibility, the Galway laboratory now routinely checks the ability of each TSA batch to support the growth of a positive control A. salmonicida.

Biochemical identification of typical Aeromonas salmonicida

Presumptive identification of typical A. salmonicida colonies by pigmentation on TSA or dark blue to navy staining on CBBA after 2-4 days’ incubation and growth at 20-25°C but not 37°C is not sufficient to make a firm diagnosis of furunculosis, and the identity of the isolate should be confirmed by other tests. Gram-staining behaviour and cell morphology of a pure culture (for example, friability of colonies on an agar surface) are essential preliminary criteria and, in combination with a limited number of biochemical tests, should be sufficient to confirm the identity of an isolate as typical A. salmonicida and to exclude other members of the Vibrionaceae, Enterobacteriaceae and Pseudomonadaceae. A diagnostic protocol to confirm the identity of typical A. salmonicida is presented in Table 10.7. More extensive tables of biochemical tests for A. salmonicida characterization are presented elsewhere (Austin and Austin, 1993; Munro and Hastings, 1993), but performance of these tests is usually unnecessary, unless it is suspected that the isolate is an atypical A. salmonicida. A number of commercial rapid test kits for biochemical identification, such as APIBioMerieux and Biolog, have become available. These test systems were developed for bacteria of medical importance and function best at incubation temperatures of 30-37°C. Their application to the biochemical characterization of A. salmonicida has been reported to generate inconsistent results following incubation at lower temperatures (Bernoth, 1997b). However, incubation of A. salmonicida at 30°C, although above its optimal growth temperature (Snieszko, 1957), will generate consistent results.

Table 7 Confirmatory identification of typical Aeromonas salmonicida.

TestCharacteristic/resultCaveats/comments
Morphological tests
Growth on TSA or BHIABrown diffusible pigment; small, friable colonies after  2-4 days’ incubation at 18-22°CPigment may be inhibited by other bacteria; colony size varies from 0.5 to 3 mm after 72 h incubation
Growth on CBBADark blue to navy staining; small, friable colonies after  2-4 days’ incubation at 18-22°COther bacteria may also stain dark blue; A-layer-negative isolates may not stain
Growth temperatureGrowth at 18-25°C; no growth at 37°C
Sedimentation testCells autoagglutinate in 0.85% PBSAutoagglutination also occurs in broth
Gram stainShort Gram-negative rods; staining tends to be bipolar
MicroscopyNon-motile; almost coccoid rods; tending to aggregate
Hanging drop testNon-motile
Biochemical tests
Cytochrome oxidase productionOxidase positiveOxidase negative isolates have been described (Chapman et al., 1991); test colonies from TSA only
Catalase productionCatalase-positiveTest colonies from TSA only
Oxidation/fermentation test (Hugh/Leifson)FermentativeIncubate at 25°C; check after 48 h and 5 days
0/129 sensitivityResistant on blood agarIncubate at 25°C; check after 48 h and 5 days
Serological tests
Latex agglutinationClumping (souring) of antibody coated latex in solution with A. salmonicida suspensionAlways include positive and negative control strains because of possible autoagglutination of A. salmonicida
A. salmonicida-targeted ELISA*Colour response from binding of conjugated antibody to A. salmonicida antigen(s)Always include positive and negative control strains

*ELISA, enzyme-linked immunosorbent assay.

Serological identification of typical Aeromonas salmonicida

Aeromonas salmonicida subsp. salmonicida requires at least 48 h incubation to produce colonies suitable for morphological and biochemical identification and, as a result, diagnosis of furunculosis may take up to 1 week. This often represents an unacceptable time-lag for the fish farmer or veterinarian, who needs to make rapid decisions on the treatment and fate of infected fish. Rapid identification methods that could be applied directly to colonies after 48 h have the potential to overcome the current delays in identifying presumptive A. salmonicida. Bernoth and Artz (1989) suggested that serological tests may be more sensitive than cultural isolation for the detection of A. salmonicida in fish tissue. A comprehensive list of studies on the development of serological identification tests for typical A. salmonicida has been presented by Bernoth (1997b). She also provides a discussion of the technical difficulties that may be encountered, such as heterogeneity of cell surface of the target species and cross-reactivity with bacteria of other species or genera that may also be isolated. While there remain considerable difficulties in applying these tests in situ in infected tissue, they can facilitate more rapid confirmation of presumptive A. salmonicida subsp. salmonicida colonies on agar plates (Bernoth, 1997b).

Diagnosis of covertly infected fish

Diagnosis of covert A. salmonicida subsp. salmonicida infections poses a number of important problems for the diagnostician, veterinarian, fish health worker or regulator. As discussed by Bernoth (1997b), disease diagnosis in fish must be understood as diagnosis in a population, rather than in an individual fish, and therefore the method of sampling becomes an important diagnostic consideration when covert furunculosis is suspected. The first sampling consideration must be the number of fish that should be sampled in order to reflect the health status of the population as a whole. Ossiander and Wedemeyer (1973) have published sampling tables which specify statistically relevant numbers of fish that should be sampled from populations of a given size. These tables have been incorporated into regulations laying down the sampling plans and diagnostic methods for the detection and confirmation of fish diseases by the European Commission (Commission Decision 92/532/EEC), the USA (Department of the Interior, 1993; Thoesen, 1994) and Canada (Department of Fisheries and Oceans, 1984). However, Bernoth (1997b) was sceptical about the reliability of such sampling tables in situations where the prevalence of covert infection is low, but acknowledged that, in the absence of any detailed epizootiological statistics, such recommendations will remain, for the moment, our ‘best guess’. Even if statistically representative numbers of fish from a fish population are sampled, detection of covert furunculosis remains problematic. A number of approaches for the detection of covert infections have been presented over the history of the disease. Hiney et al. (1997b) grouped these methods according to what they demonstrated about the infection, i.e. the ability of infected fish to shed A. salmonicida into their environment and transmit disease, the presence of the organism or signs of the organism in/on covertly infected fish or the precipitation of clinical furunculosis following the application of stress.

Demonstration of shedding and transmission of typical Aeromonas salmonicida

The shedding of A. salmonicida by covertly infected fish can be demonstrated indirectly, by cohabitation studies, or directly, by detection of the organism in the environment of such fish. Cohabitation studies have been used by many authors to demonstrate shedding and transmission of furunculosis. A number of different transmission scenarios have been reported, including transmission of clinical furunculosis from clinically infected fish to healthy fish, transmission of clinical furunculosis from covertly infected fish to healthy fish, transmission of covert furunculosis from clinically infected fish to healthy fish and transmission of covert furunculosis from covertly infected fish to healthy fish - so called ‘silent transmission’ (McCarthy, 1977a; Scallan, 1983; Hiney et al., 1997b). Transmission experiments do not, however, represent a useful diagnostic strategy for detection of covert infections.

Direct shedding of A. salmonicida into the environment by subclinically or clinically infected fish has been demonstrated by culture methods (Scallan, 1983; Ford, 1994; Cipriano et al., 1996a), using immunological assays (Enger and Thorsen, 1992; Gilroy and Smith, 1995) and polymerase chain reaction (PCR)/DNA probe assays (Gustafson et al., 1992; O’Brien et al., 1994). Culture-based detection of A. salmonicida in the environment has traditionally been seen as problematic (Cornick et al., 1969). However, the success of Ford (1994) and Cipriano et al. (1996a), using dilution filtration and CBBA, in isolating typical A. salmonicida from hatchery water suggests that these methods are promising in routine monitoring of hatchery water supplies (Cipriano, 1998). None the less, the use of culture-based techniques are complicated, as A. salmonicida may enter a non-culturable-but-viable state once it has been shed into the environment (Roszak and Colwell, 1987; Enger, 1997). Therefore, techniques that do not rely on culture, such as PCR and enzyme-linked immunosorbent assay (ELISA), have received considerable attention and may have the potential to overcome the problems encountered by culture-based techniques. However, these non-culture-based detection techniques present significant problems of interpretation when applied in the environment (Hiney, 1994; Hiney et al., 1997a; Hiney and Smith, 1998) and will be discussed later in this section.

Detection of Aeromonas salmonicida subsp. salmonicida or its components

The difficulty of isolating A. salmonicida or its components (antigens, DNA) from covertly infected fish, in terms of not only the lack of selective bacteriological methods (Cornick et al., 1969; Gustafson et al., 1992) but also the reliability of current diagnostic techniques (Inglis et al., 1993; Crane and Bernoth, 1996; Hiney and Smith, 1998) and the time-consuming nature of some of these techniques, has been discussed (Austin and Austin, 1993). The problem of deciding which organs of the fish should be examined is central to all methods that fall into this group. However, as discussed, the location of A. salmonicida in covertly infected fish remains uncertain.

Attempts to culture A. salmonicida from the internal organs of fish suspected of covert infection constitute the method used by many authors. However, there has been some disagreement about precisely which organ or organs are most likely to yield the organism where no clinical signs are apparent. Some workers have favoured the kidney as the organ of choice for detection of covert infections (Mackie et al., 1935; Mackie and Menzies, 1938; McCarthy, 1977a), but others have demonstrated that examination of a number of organs is more efficient (McDermott and Berst, 1968; Daly and Stevenson, 1985; Sutherland and Inglis, 1992). In older salmonids, Nomura et al. (1991a,b, 1992, 1993) reported the isolation of A. salmonicida from coelomic fluid, as well as the kidney.

The application of non-culture-based techniques to the detection of components of A. salmonicida in tissues of covertly infected fish has not resolved the issue of location in this infection type. In two studies which applied an A. salmonicida-targeted ELISA to examination of the kidneys of similar salmonid populations with SIF, A. salmonicida antigens were detected in one study (Rose et al., 1989) but not in the other (Hiney et al., 1994). Similarly, using PCR amplification and A. salmonicida-targeted DNA probes, positive responses have been detected in a variety of salmonid organs, including spleen and kidney (Gustafson et al., 1992; Høie et al., 1997) and blood (Mooney et al., 1995; Høie et al., 1997). However, in none of these studies was covert infection demonstrated, and few successfully isolated the organism in primary culture from the samples tested, making it difficult to interpret the meaning of these data.

The use of external surfaces, including the intestine, as sampling sites when attempting to detect typical A. salmonicida or its components has also been reported. Although the intestine has been demonstrated as a site of colonization of A. salmonicida in apparently healthy fish (Horne, 1928; Mackie et al., 1930) and in fish that have been artificially challenged (Hodgkinson et al., 1987; Markwardt and Klontz, 1989a,b), it has never been a popular sampling site. The main reason for this is rapid overgrowth by concomitant flora, which complicates isolation of A. salmonicida (Bernoth, 1997b). None the less, the intestine is currently recommended as a sampling site for covertly infected fish in the UK (Munro and Hastings, 1993) and USA (Shotts, 1984). This recommendation is considered misplaced. The difficulties encountered with culture from the intestine make it unlikely that a covert infection will be detected by this means. The use of non-culture-based detection techniques for A. salmonicida may have the potential to overcome the difficulties of bacteriological culture from the intestine. Intestinal contents have been reported to yield positive results using both an A. salmonicida-targeted ELISA assay (Rose et al., 1989; Hiney et al., 1994) and PCR-based assays (Gustafson et al., 1992; Padley et al., 1997). However, until these assays have been adequately validated for routine analysis they will remain research tools.

Perhaps the most extensive work on detection from external locations has been carried out by Cipriano and co-workers. Using CBBA as a differential medium (Cipriano and Bertolini, 1988; Markwardt et al., 1989), they demonstrated that A. salmonicida could be isolated from the external mucus of hatchery-reared salmon (Cipriano et al., 1996a), hatchery trout (Cipriano et al., 1992, 1994) and feral salmon returning to spawn (Cipriano et al., 1996c) more frequently than from the kidneys of the same fish. The method of Cipriano et al. (1992) has the advantage of being non-lethal, which would be welcome in restoration programmes and when assessing valuable brood stock (Cipriano, 1997). Culture of A. salmonicida from mucus, gill and kidney from covertly infected fish was more frequent than obtaining positive PCR tests of the samples (Byers et al., 2002).

Detection of covert furunculosis by the application of stress

Bacteriological culture from one or more organs may detect preclinical infections or covert infections present at high levels in a fish population, but culturing is unlikely to detect low levels of covert infections (McCarthy, 1977a; Scallan, 1983; Hiney, 1995; Cipriano et al., 1997). Bullock and Stuckey (1975) found that injection of the corticosteroid triamcinolone acetone (at 20 mg kg-1 body weight), combined with heat stressing at 18°C for 14 days, activated latent infections in covertly infected fish and facilitated their detection. The stress test proposed by Bullock and Stuckey (1975) was demonstrated to be more efficient at detecting covert infection than either heat stressing alone, as used by Plehn (1911), or bacteriological culture alone (Blake and Clark, 1931; Mackie et al., 1935; Mackie and Menzies, 1938). A number of modifications to the method of Bullock and Stuckey (1975) were later published by McCarthy (1977a), Jensen (1977), Scallan (1983) and Scallan and Smith (1985). Scallan (1983) described a method for the bath administration of the corticosteroid prednisolone acetate to fish of 5 g or less, which might not survive injection administration. She also demonstrated that the amount of corticosteroid injected into fish was not a critical factor in precipitating overt disease, with concentrations over a four-fold range being effective. In a comparison of the stress test and culture-based assays, Cipriano et al. (1997) found that the probability of detecting A. salmonicida in apparently healthy salmon and trout, where the prevalence of covert infection was assumed to be low, was 17 times greater using stress testing than direct culture of either external mucus or kidney. When a 24 h pre-enrichment of kidney and mucus was included prior to plating, the probability of detecting A. salmonicida was 10 and 27 times greater for stress testing, as compared with kidney and mucus culture, respectively (Cipriano et al., 1997). It should be noted that infections detected by the stress test of Bullock and Stuckey (1975) or its later modifications should properly be termed SIF, and the relationship between SIF and covert infections detected by other methods should be considered unknown (Hiney et al., 1994).

Limitation on the use of the stress-inducible furunculosis test for detection of covert furunculosis infections

In Ireland, reliance on stress testing is widespread at the level of individual companies and has been successful, when used, in limiting the spread of furunculosis to marine farms (Smith, 1992; Scallan and Smith, 1993). Stress testing of salmon smolts prior to their transport to sea is a regulatory requirement in New Brunswick, Canada, and has also been found to be a successful measure there (Olivier, 1992). However, although stress testing is valuable in the field, it is not without its problems. Stress testing tends to result in the isolation of mixed cultures from the kidney, making identification of A. salmonicida from this tissue difficult (Cornick et al., 1969). Pigmentation of A. salmonicida strains, often used as a preliminary confirmation of their presence, is inhibited in the presence of other organisms, requiring careful examination and repeated subculture of kidney streaks to confirm diagnosis (McCarthy, 1977a). Even so, the presence of A. salmonicida may be missed (Scallan, 1983). No selective medium exists for the isolation of A. salmonicida from mixed cultures, although CBBA has been found to simplify identification of putative A. salmonicida colonies isolated from the kidneys of stress tested fish. Furthermore, the stress test is by definition lethal. It requires 14-16 days for a definitive diagnosis and uses large numbers (150-200) of fish to ensure detection of the infection where prevalence is low. However, Bullock and Cipriano (1997) found, following sampling of the mucus and gills of stress-tested fish, that, from day 5 of the test onwards, A. salmonicida could be detected in these samples, thus eliminating the need to continue the test to 14 days. This method may, therefore, represent a more rapid alternative to the current stress test.

Prior vaccination of SIF-tested fish may complicate interpretation of the results obtained from a stress test. Hiney (1995) demonstrated that, following stress testing by the method of McCarthy (1997a), A. salmonicida could be isolated from the kidney of Atlantic salmon and brown trout that had been vaccinated against furunculosis. Attempts to culture A. salmonicida from the kidneys of parallel, unstressed, groups of salmon and trout were unsuccessful, confirming that the infection was of a stress-inducible nature. The results of Hiney (1995) suggested that vaccinated fish could become or remain covertly infected following vaccination and that the extent of protection provided by vaccination was not sufficient to prevent SIF. Hiney et al. (1997b) and Smith (1997) have suggested that covertly infected fish with increased systemic immunity (i.e. vaccinated) might act as ‘immune carriers’. If such fish exist, then they will present a number of important questions for managers of fish farms and wild fisheries. For example, should potential ‘immune carriers’ be treated with antimicrobial agents immediately prior to transfer to a sea site or restocking into fresh water in order to eliminate any carried A. salmonicida? In theory, ‘immune carriers’, while remaining disease-free, could act as a source of infection for unvaccinated fish. However, at present not enough is known about the interactions of covert infection, vaccination and immunity to address the issue of ‘immune carriers’.

Rapid diagnostic methods

An important development in diagnostic microbiology over the last 25 years has been the move to detect microorganisms directly in clinical samples or in the environment of the host, without the need to culture. In line with this general trend, non-culture-based microbial detection techniques are being increasingly developed for the identification of typical A. salmonicida. Since 1990, in particular, a number of genetic-based assays for A. salmonicida have been described, and these are presented in Table 8 (below), along with immunological assays for the organism developed during the same period. For a more exhaustive list of immunological assays developed prior to 1990 readers are referred to Bernoth (1997b). Although many of the techniques presented in Table 8 are intended for clinical diagnosis of furunculosis, their potential for detecting A. salmonicida in covertly infected fish and their environment has also been investigated. In theory, non-culture-based techniques can be designed to specifically detect low numbers of a target organism in a mixed microbial flora. In addition, because culturing of the target organism is not a requisite for its detection, they circumvent the numerous problems of loop and plate microbiology (Bernoth, 1997b; Pace, 1997). Non-culture-based techniques developed for the detection of A. salmonicida fall into two broad categories, those based on immunological principles and those based on genetic principles.

Table 8 Non-culture-based techniques developed for, or applied to, the detection of typical Aeromonas salmonicida since 1990.

Target sample

Detection principleAssay typeClinicalEnvironmentalReference
ImmunologicalELISAKidneyAdams and Thompson (1990)
ELISAKidneyBernoth (1990b)
IFAT, ELISAKidney, liverLallier et al. (1990)
ImmunofluorescenceWater, sedimentEnger and Thorsen (1992)
ELISABloodYoshimizu et al. (1992)
ELISAKidney, intestineHiney et al. (1994)
Immunodot-blotVarious tissuesBanneke and Bernoth (1994)
ELISAKidneySedimentGilroy and Smith (1995)
ELISAKidney, intestineSedimentHiney et al. (1997a)
ELISAKidney, mucus, gills, spleenBullock et al. (1998)
Genetic16S rDNA-PCRPure cultureBarry et al. (1990)
DNA-PCRSpleen, kidneyFish faeces, tank effluentGustafson et al. (1992)
DNA-PCRPure cultureHiney et al. (1992)
DNA-PCRFreshwater microcosmMorgan et al. (1993)
DNA-PCRMarine sediment, waterHiney (1994)
DNA-PCRHatchery effluentO’Brien et al. (1994)
DNA-PCRBloodMooney et al. (1995)
DNA-PCRFreshwater microcosmPickup et al. (1996)
16S rDNA-PCRKidney, spleenHøie et al. (1996)
DNA-PCRKidneyMiyata et al. (1996)
DNA-PCRKidney, intestineFreshwater sedimentHiney et al. (1997a)
DNA-PCRKidney, gill swabsHøie et al. (1997)
DNA-PCRKidneyJençiç and Piano (1997)
DNA-PCRKidney, intestineFreshwater sedimentPadley et al. (1997)
DNA-PCRKidneySørum et al. (1998)

IFAT, indirect fluorescent antibody test; rDNA, ribosomal DNA.

Immunological detection techniques

Immunological detection techniques offer many advantages over culture-based techniques, particularly when attempting to detect organisms such as A. salmonicida, whose growth on laboratory media can be inhibited by the presence of other bacteria. These techniques have been revolutionized in recent years by the introduction of monoclonal antibodies and ELISAs, which allow for more specific assays that can be semiautomated. As a result, a number of ELISAs have been developed for screening of clinical samples for signs of A. salmonicida (Bernoth, 1997b; see table 8 above). In a comparison of ELISA and an indirect fluorescent antibody test (IFAT), similar to the fluorescent antibody microscopy (FAM) technique, Lallier et al. (1990) found that ELISA was more sensitive than IFAT when tested on pure cultures of A. salmonicida and both methods were found to be more sensitive than bacteriological culture. However, it has been argued that the use of IFAT coupled with experience overcomes the problems of lesser specificity of this technique, making it comparable to ELISA (E.-M. Bernoth, CSIRO, 1995, personal communication). Perhaps the most useful application of immunological assays would be in the detection of covert A. salmonicida infections, and a number of ELISA tests have been applied for this purpose. Good correlation was found between detection of covert A. salmonicida infection by stress testing and ELISA of kidney material from non-stress-tested fish (Scallan, 1983; Rose et al., 1989). These findings were supported by Hiney et al. (1994), who found that ELISA examination of the kidney, mucus and intestine of covertly infected fish detected A. salmonicida antigens in 45% of fish as compared with culture of the organism from the kidney of 24% of a parallel group of stress tested fish. On the other hand, Bullock et al. 1997 found that culture of mucus, gills, kidney and spleen was more sensitive than ELISA for identification of A. salmonicida in fish that had been stress-tested. Enger and Thorsen (1992) have reported on an IFAT which was applied to detection of A. salmonicida antigens in the environment of a fish farm. None of these tests has, however, been applied successfully under true field conditions, nor are they recommended with any conviction in diagnostic manuals (Crane and Bernoth, 1996).

There are a number of problems to resolve when attempting to detect bacterial antigens in situ in tissue or environmental samples. Many of the immunological assays developed do not appear to offer any greater sensitivity or reliability than conventional bacteriological methods (Inglis et al., 1993). The lower limit of detection has been found to be 103 cells ml-1 or greater in both clinical and environmental samples (Sakai et al., 1986; Adams and Thompson, 1990; Bernoth, 1997b). A major problem with cross-reactivity of antisera or antibodies to epitopes expressed by ubiquitous motile Aeromonas species or other aquatic microorganisms has been reported, which requires that confirmatory bacteriological isolation of the pathogen be performed (Bernoth, 1990b). Importantly, immunological assays do not differentiate between live and dead cells (Rose et al., 1989) and A. salmonicida-targeted ELISA has been shown to generate positive results from the spleen and intestine of fish which were immunized with a killed whole-cell furunculosis vaccine (Gilroy and Smith, 1997). Another problem with immunological assays is that the antiserum raised against bacteria grown in vitro must detect the organism as it occurs in situ in a clinical or environmental sample. For A. salmonicida, at least, cells grown in vivo have been found to express novel antigens, including an antigenically new form of lipopolysaccharide, which were not induced under in vitro growth conditions (Garduño et al., 1993; Thornton et al., 1993). The question of how relevant the current methods of antiserum generation from in vitro-grown A. salmonicida components are for diagnosis in situ has to be addressed.

Genetic detection techniques

A second family of non-culture-based detection techniques that have been investigated for the detection of components of A. salmonicida is based on genetic principles. Typical A. salmonicida is homogeneous at the genetic level (Vaughan, 1997) and presents, in theory, an ideal candidate for a genetic-based detection technique. Since 1990, a number of assays targeted against 16S ribosomal ribonucleic acid (rRNA) or DNA sequences of A. salmonicida have been developed for this organism (see table 8 above). Most of these assays also utilize the ability of PCR to amplify the number of copies of the target sequence in a sample between several thousand- and 1 million-fold, making detection of initially minute quantities of that sequence possible. However, it is difficult to estimate the true lower detection limits of PCR-based assays for A. salmonicida DNA sequences, because the kinetics of PCR amplification are essentially non-linear and amplification is prone to variable inhibition by facets of both clinical and environmental samples (Wilson, 1997).

Similar to immunological assays, the ultimate goal of genetic techniques is to specifically detect DNA sequences of A. salmonicida in the tissues or environment of both clinically and covertly infected fish. Clearly, then, the key parameter of this type of technique is its specificity for the target organism. Because of the relative newness of genetic detection techniques in comparison with immunological techniques, there have been few reports to date about cross-reactivity of these techniques. None the less, Høie (1995) found that his PCR-based assay for an A. salmonicida DNA sequence cross-reacted with an uncharacterized aquatic organism, and Hiney and Smith (1998) have expressed concern about the relevance of laboratory studies of specificity for field applications. In addition, PCR-based techniques are prone to false-positive results through sample contamination, especially if a double cycle of PCR amplification is employed, so-called ‘nesting’ (Byers et al., 1997). As with immunological assays, many PCR-based assays do not differentiate between live and dead cells, and they have been demonstrated to detect DNA from killed whole cells in the spleen and head kidney of fish vaccinated against furunculosis (Høie et al., 1996), suggesting that this type of assay is unsuitable for application to vaccinated stocks. Molecular-based diagnostic methods for A. salmonicida have been reviewed by Colquhoun and Cunningham (2002).

Validation of non-culture-based detection techniques

The biggest drawback to the use of non-culture-based detection techniques for A. salmonicida is the difficulty of interpreting the results generated by these techniques in field samples. Genetic techniques ‘see’ a small segment of the microbial genome, while immunological techniques ‘see’ one or more epitopes on the microbial surface. Therefore, the key characteristic of these techniques is that they are proxy measurements of microbial presence, that is, indirect indices that are presumed to signal the presence of the target organism in a sample (Wildavsky, 1995). Where the target organism is a pathogen, the target sequence or epitope(s) may also be used as a proxy measurement of disease potential. In fish health studies, it is probable that this will, in fact, be the meaning attributed to any data generated by a non-culture-based detection technique. The use of proxy measurements does not preclude the generation of meaningful data but does present difficulties about the interpretation of results (Wildavsky, 1995). Hiney (1997) and Hiney and Smith (1998) have argued strongly that these difficulties can only be overcome by validation, that is, demonstration that a technique does what it is supposed to do (Thrusfield, 1986). Although important for all techniques, validation is essential for techniques that involve proxy measurements.

Validation is not the property of a technique but, rather, is a property of its application. It establishes that a technique can be correctly and properly used for a particular purpose. A formal structure for the validation of non-culture-based detection techniques in laboratory studies was presented by Hiney and Smith (1998). However, no amount of laboratory studies can validate the application of such techniques under field conditions. Comparative and predictive validation represent the only two available strategies for the validation of such applications. In comparative methods, the technique under test can be compared either with a technique that has been previously validated or with one which is, itself, also unvalidated. This second approach, mutual covalidation, is the more frequently encountered but is, however, limited in power.

Predictive validation requires that the application intended for the technique must be clearly defined in terms which are empirically meaningful. For example, the intended application might be the prediction of the disease incidence for hatchery smolts covertly infected with A. salmonicida following transfer to a sea site. An empirically meaningful measure of ‘disease’ could, in this case, be the frequency of the isolation of A. salmonicida from the kidney of moribund fish, following transfer. The process of predictive validation would then involve measuring the degree of correlation between the results generated by the non-culture-based technique and the incidence of positive isolation of A. salmonicida. Such a correlation, if it is to be useful, must be determined over a number of years and in a variety of environmental contexts. When, and only when, it has been established that a satisfactory correlation exists can the technique be used alone and the data it generates be interpreted as a proxy measurement of disease. Simply stated, techniques can only be used to predict an event if they have been shown to predict it.

Despite the importance of validation, few papers that have reported on the development of non-culture-based assays for A. salmonicida have paid adequate attention to demonstrating that these techniques have field validity. Hiney et al. (1997a) illustrated the danger of assuming that a positive response, generated by a non-culture-based assay for A. salmonicida in a field sample, was indicative of the presence of the organism in a form capable of causing disease. In their study, positive responses generated from hatchery inflow sediment by A. salmonicida-targeted ELISA and a DNA/PCR assay showed no correlation with the health status of fish at that hatchery over a 2-year period, as assessed by routine bacteriological analysis and stress testing. Furthermore, Stanley et al. (2002) found that in a humic acid-rich microcosm A. salmonicida could be detected for over 200 days by PCR and ELISA methods, but the ability to culture the bacteria ceased after one day and these samples were incapable of producing disease in fish. These results suggest that interpretation of the results generated by either of these techniques as indicators of the presence of a disease risk would be absolutely invalid. More seriously, the use of this type of data by regulators would be both unwarranted and dangerous. There is little doubt that the current developments in non-culture-based techniques for the detection of A. salmonicida have the potential to improve our understanding of, and limit the impact of, diseases caused by this pathogen. However, as discussed, adequate validation of the development and utilization of these techniques is critical to their correct interpretation, especially if that interpretation will be used to impose regulatory limitations on aquaculture activities. More recently, these authors have argued that laboratory-scale studies are not adequate to validate the application of proxy methods to field samples (Smith et al., 2003).

Diagnostic methods for atypical Aeromonas salmonicida

Diagnosis is usually based on culturing of the pathogen. The identification of non-fastidious atypical isolates of the subspecies achromogenes is relatively straightforward, because all these strains are reported to grow on conventional media, such as TSA and BHIA. The strains can be chromogenic or not; however, growth is often slower than that of typical isolates (Evelyn, 1971; Wichardt et al., 1989; Rintamäki and Valtonen, 1991; Olivier, 1992; Wiklund and Dalsgaard, 1995). Samples of ulcers or lesions, in addition to kidney tissues, should be cultured to confirm the presence of atypical isolates, because some organisms may not be found in internal organs. In Iceland, the recovery of A. salmonicida subsp. achromogenes was superior when gills of subclinically or covertly infected Atlantic salmon were cultured, rather than kidney tissues (Benediktsdottir and Helgason, 1990).

Several problems are associated with the culture of fastidious strains belonging to the subspecies nova. A culture medium containing blood is required for the primary isolation of atypical strains of this subspecies, including isolates from carp, goldfish and eel, and some fastidious isolates recovered from salmonids (Bootsma et al., 1977; Paterson et al., 1980; Böhm et al., 1986; Ishiguro et al., 1986; Olivier, 1992). Ishiguro et al. (1986) investigated this phenomenon and found that fastidious atypical strains isolated from goldfish and salmon (including Haemophilus piscium) could grow well if conventional medium (TSA) was supplemented with haemin (10 µg ml-1): this was confirmed by Nakai et al. (1989) with atypical strains isolated from Japanese eel (Anguilla japonica).

Isolation plates are often contaminated with what is believed to be opportunistic pathogens, such as motile aeromonads and Pseudomonas spp., thus complicating the diagnosis. This phenomenon has been reported by several investigators in both natural infections and during experimental challenges with strains causing carp erythrodermatitis (Csaba et al., 1984; Evenberg et al., 1986), goldfish ulcer disease (Whittington et al., 1987) and the causative agent of the ulcerative disease of eels (Nakai et al., 1989). Often, the organism is only isolated from skin lesions and the infection is not systemic (Elliot and Shotts, 1980; Csaba et al., 1984; Böhm et al., 1986; Noga and Berkhoff, 1990). In rare instances, the diagnosis can be complicated by the presence of both typical and atypical strains as reported by Noga and Berkhoff (1990) in American eels (Anguilla rostrata).

Biochemical identification

Similar to typical isolates, the presumptive identification of atypical isolates is based on a few characteristics, i.e. the isolates are Gram-negative coccobacilli, oxidase-positive and fermentative and do not grow at 37°C. It is, however, important to note that some atypical strains are oxidase-negative, including isolates recovered from flounder (Wiklund and Bylund, 1991), herring (one out of four isolates) (Traxler and Bell, 1988), tomcod (Olivier, 1992) and turbot (Scophthalmus maximus) (Pedersen et al., 1994). These results strongly suggest that care is needed to properly identify some of these atypical isolates, and that even discrepancies in basic characteristics, such as the oxidase reaction, should not necessarily cause a tentative identification to be rejected. Additional tests are needed to ensure that atypical strains are not implicated in the condition under investigation. A schematic representation of the various steps that should be taken for the culture and identification of atypical strains of A. salmonicida is presented in Table 13.

Table 13 Diagnostic procedure to culture and identify atypical Aeromonas salmonicida.

TestCharacteristic/resultCaveats/comments
Sampling procedure
External samplesSample from lesion materialSampling preferable during early or acute phase; test several fish
Internal samplesSamples from kidney, spleen, heartTest several fish
Morphological tests
Growth on TSA or BHIAOnly non-fastidious strains will grow on these agars; delayed pigment production after extended incubation at 18-22°CMany strains do not produce pigment; incubate for at least 7 days
Growth on blood agarFastidious stains require this medium; small, creamy colonies strongly adhering to the mediumIncubate for at least 7 days
Growth temperatureGrowth at 18-22°C; no growth at 37°CStrains which grow at 37°C have been reported (Austin, 1993)
Sedimentation testCells autoagglutinate in 0.85% PBS
Gram stainShort Gram-negative rods
Hanging drop testNon-motile
Serological tests
Slide agglutination (presumptive identification only)Clumping (souring) of antibody-coated latex in solution with cell suspensionAlways include positive and negative control strains because of possible autoagglutination of A. salmonicida
Biochemical tests
Cytochrome oxidase productionOxidase-positiveOxidase-negative isolates have been described (Traxler & Bell, 1988; Olivier, 1992; Pederson et al., 1994; Wiklund et al., 1994)
Catalase productionCatalase-positiveCatalase-negative isolates have been described (Csaba et al., 1984; Böhm et al., 1986)
Indole productionIndole-positiveIndole-negative isolates have been described (Austin et al., 1989; Wiklund, 1990; Olivier, 1992)
Oxidation/fermentation test (Hugh/Leifson)FermentativeIncubate at 25°C; check after 5 and 7 days
Substrate utilizationSucrose-positive; do not produce gas from glucoseNot all strains will be positive for these tests
Resistance to antimicrobial agentsResistant to ampicillin (25 mg) and cephaloridin (15 mg)Incubate at 25°C; check after 5 and 7 days; not all strains will be positive for these tests

Differentiation between typical and atypical isolates is achieved based on the following properties; atypical isolates are generally achromogenic, lack the capacity to produce gas from glucose, utilize sucrose, produce indole and are gelatinase-negative (Popoff, 1984). All of these characteristics are useful for the differentiation of atypical isolates, although it must be borne in mind that exceptions to the above have been reported. A more comprehensive list of biochemical characteristics is presented in Table 14, although this level of investigation may not be necessary for routine diagnosis.

Table 14 Biochemical characteristics distinguishing between typical and atypical strains of Aeromonas salmonicida.

Differential characteristics of Aeromonas salmonicida subspecies based on McCarthy (1978), Popoff (1984), Austin et al. (1989), Wichart et al. (1989) and Olivier (1992, unpublished results).
+, –, V indicate that > 80%, < 20% and 21-79% of the strains gave positive reactions, respectively.
0 indicates not tested.

Pigment

salmonicida +: masoucida -: acromogenes V: nova -: smithia -

Haemin requirement

salmonicida -: masoucida -: acromogenes -: nova +: smithia 0

Gas from glucose

salmonicida +: masoucida +: acromogenes -: nova -: smithia 0

Arbutine

salmonicida +: masoucida +: acromogenes V: nova -: smithia 0

Sucrose

salmonicida -: masoucida +: acromogenes +: nova V: smithia V

Salicin

salmonicida +: masoucida -: acromogenes V: nova -: smithia 0

N-Acetylglucosamine

salmonicida +: masoucida -: acromogenes V: nova -: smithia 0

Dextrin

salmonicida +: masoucida +: acromogenes +: nova -: smithia 0

Fructose

salmonicida +: masoucida +: acromogenes +: nova V: smithia 0

Galactose

salmonicida +: masoucida +: acromogenes V: nova V: smithia -

Glycerol

salmonicida +: masoucida +: acromogenes +: nova -: smithia -

Mannose

salmonicida +: masoucida +: acromogenes +: nova V: smithia 0

Maltose

salmonicida +: masoucida +: acromogenes +: nova V: smithia -

Trehalose

salmonicida +: masoucida +: acromogenes +: nova V: smithia -

Indole

salmonicida -: masoucida +: acromogenes V: nova +: smithia -

Aesculin

salmonicida +: masoucida +: acromogenes -: nova -: smithia -

Gelatinase

salmonicida +: masoucida +: acromogenes V: nova -: smithia +

Lecithinase

salmonicida +: masoucida -: acromogenes -: nova -: smithia -

Elastase + – – – –

salmonicida +: masoucida -: acromogenes -: nova -: smithia -

Protease

salmonicida +: masoucida -: acromogenes +: nova -: smithia V

Resistance to

Ampicillin (25 µg)

salmonicida -: masoucida +: acromogenes +: nova +: smithia +

Cephaloridine (15 µg)

salmonicida -: masoucida +: acromogenes +: nova V: smithia 0

Polymyxin B (300 IU)

salmonicida -: masoucida -: acromogenes -: nova V: smithia 0

Salmonid source

salmonicida +: masoucida +: acromogenes V: nova V: smithia -

Serological identification of atypical Aeromonas salmonicida

Because it may be difficult to grow atypical A. salmonicida, other diagnostic methods have been explored, including immunofluorescence (fluorescent antibody test (FAT)), as described by Böhm et al. (1986). Using this method, serological confirmation of atypical furunculosis in carp and goldfish was at least 50% higher than by culture. Sövényi (1986) described a coagglutination test for the diagnosis of carp erythrodermatitis and found that all experimentally induced ulcerative lesions were positive by this test, with no cross reactions with other fish pathogens being observed. The ELISA assay described by Adams and Thompson (1990) was also able to detect atypical strains. Methods used to detect typical isolates should be effective in detecting atypical isolates, because of their serological similarities, and simple methods, such as FAT, could be used as an initial screening for these pathogens. Modern techniques, such as PCR, will undoubtedly become available to detect these isolates, although, as has been discussed above, the routine diagnostic use of these methods cannot be considered without extensive validation of the techniques.

List of Symptoms/Signs

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SignLife StagesType
Finfish / Cessation of feeding - Behavioural Signs Aquatic:Adult Sign
Finfish / Cessation of feeding - Behavioural Signs Aquatic:Adult Sign
Finfish / Darkened coloration - Skin and Fins Aquatic:Adult Sign
Finfish / Darkened coloration - Skin and Fins Aquatic:Adult Sign
Finfish / Darkened coloration - Skin and Fins Aquatic:Fry Sign
Finfish / Generalised lethargy - Behavioural Signs Aquatic:Adult Sign
Finfish / Generalised lethargy - Behavioural Signs Aquatic:Adult Sign
Finfish / Haemorrhagic lesions - Skin and Fins Aquatic:Adult Sign
Finfish / Haemorrhagic lesions - Skin and Fins Aquatic:Adult Sign
Finfish / Haemorrhaging - Body Cavity and Muscle Aquatic:Fry Sign
Finfish / Increased respiratory rate (increased opercular movements) - Behavioural signs Aquatic:Adult Sign
Finfish / Increased respiratory rate (increased opercular movements) - Behavioural signs Aquatic:Fry Sign
Finfish / Liver - white / grey patches (haemorrhage / necrosis / tissue damage) - Organs Aquatic:Adult Sign
Finfish / Paleness - Gills Aquatic:Adult Sign
Finfish / Pop-eye - Eyes Aquatic:Adult Sign
Finfish / Pop-eye - Eyes Aquatic:Fry Sign
Finfish / Splenomegaly - spleen swelling / oedema - Organs Aquatic:Adult Sign
Finfish / Splenomegaly - spleen swelling / oedema - Organs Aquatic:Adult Sign

Disease Course

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Clinical infection by typical Aeromonas salmonicida

Classical furunculosis derives its name from the boil-like lesions observed by Emmerich and Weibel (1894) on the skin and in the musculature of infected fish. However, development of ‘furuncles’ on the dorsal body are the exception rather than the rule (Bernoth, 1997b) and, in the experience of this author, only occur in older fish suffering from the chronic form of the disease. The clinical manifestations of furunculosis are often divided into peracute, acute and subacute or chronic forms and will be discussed here under these headings (see table 4 below). It should be noted that clinical manifestations of more than one form of the disease may be present in individual fish within a population (Bernoth, 1997b). Reviews on the macro- and microscopic features of furunculosis are given in Ferguson (1977), McCarthy and Roberts (1980), Frerichs and Roberts (1989), Armstrong (1992), Austin and Austin (1993) and Bernoth (1997b).

Table 4 Diagnosis of clinical infections of typical A. salmonicida aetiology (documented signs).

Type of clinical infection

PeracuteAcuteSubacute/Chronic
Age of fishVery young fishGrowing fishUsually older fish
Clinical signsDarkening of skin; tachybranchia (rapid breathing); slight exophthalmus (pop-eyed)Darkening of skin; inappetence; lethargy (sluggishness); tachybranchia; small haemorrhages at base of finsSlight darkening of skin; inappetence; lethargy; congested blood vessels at base of fins; slight exophthalmus; expression of serosanguineous fluid from nares and vent; may have pale or congested gills; may have furuncles on flank or dorsal surface
Gross pathologyDilated blood-vessels; punctate haemorrhages in parietal and visceral peritoneum and over myocardium; may have focal haemorrhages in gillsHyperaemia of serosal surfaces; punctate haemorrhages scattered over abdominal walls, viscera and heart; soft and friable or liquefied kidney; enlarged, cherry-red spleen with round edges; pale liver with subcapsular haemorrhages or mottled appearance; stomach and intestine void, may contain sloughed epithelium, mucus and blood; swim-bladder wall clouded and hyperaemic; haemorrhagic patches along body side or raised furuncles in dermisGeneral visceral congestion and peritonitis; multiple haemorrhages in muscle and liver; splenomegaly; necrotic kidney; adhesions between viscera and between viscus and abdominal cavity; intestinal inflammation; septic, necrotic and haemorrhagic muscle lesions (furuncles)
HistopathologySmall bacterial colonies in branchial mesenchyme, myocardium, anterior kidney and spleen; limited localized necrosis; cardiac damage as possible cause of deathPrincipal bacterial focus may be in any single organ or be multiply located; toxic haematopoietic necrosis; myocardial and renal tubular degeneration; focal hepatic necrosis; initial lesions in gills may cause lamellar thrombosisHeart and spleen most infected organs; microcolonies in vascular endothelia; may be massive destruction of spleen ellipsoids, resulting in vascular collapse; damage to spleen ellipsoids may be accompanied by reticular cell proliferation and lymphocyte accumulation; degeneration of cardiac ventral epicardium fibrinoid and collagen; marked inflammation of epicardium; reticulin and collagen damage in spleen and heart; toxic cardiac necrosis, especially of atrial lining
Outcome of infectionMay die rapidlyMay die within 2-3 daysLow mortality, healed furuncles may leave scar tissue

PERACUTE FURUNCULOSIS

Because the peracute form of furunculosis is usually restricted to young fish, whose defences against a severe bacterial septicaemia will be poor, this form of the disease normally results in rapid death with little more that slight exophthalmus (McCarthy and Roberts, 1980; Frerichs and Roberts, 1989). The gross pathology of peracute furunculosis is typical of a peracute septicaemia. Microcolonies can be observed histologically in a number of organs, with no inflammatory infiltration and only limited necrosis. McCarthy and Roberts (1980) considered that cardiac damage was the most likely cause of death in young fish.

ACUTE FURUNCULOSIS

In growing fish, furunculosis tends to occur in an acute form, which is manifested clinically by a generalized bacterial septicaemia displaying the ‘standard’ features (table 4 above). As its name implies, acute furunculosis is often fatal in 2-3 days and, because of the short duration of the disease, furuncle development is unusual. Fish with an acute infection show signs of a haemorrhagic septicaemia, including bloody anal vents. Skin lesions may be haemorrhagic patches or blotches along the side or on the dorsal body surface, or, more typically, raised furuncles, which usually develop in the dermis rather than the hypodermis (Bernoth, 1997b).

SUBACUTE/CHRONIC FURUNCULOSIS

The chronic form of furunculosis is common in older fish and is probably the form first observed by Emmerich and Weibel (1894). In chronic cases, fish may show a lesser degree of skin darkening and inappetence than in the acute form. Other signs are summarized in table 4 above. Furuncles are likely to be observed during the progress of a chronic infection and, where they do occur, more than one lesion may be present. These furuncles may be large and, when ruptured, the viscous fluid may contain more necrotic material than furuncles found in acute cases (Bernoth, 1997b). For a description of the histopathology of furuncles, see the reviews of McCarthy and Roberts (1980) and Frerichs and Roberts (1989).

INTESTINAL FURUNCULOSIS

A fourth form of furunculosis associated with low mortality, intestinal furunculosis, has been described by Amlacher (1961) (cited in Austin and Austin, 1993). The only clinical sign of this form of the disease was prolapse of the anus, although examination revealed haemorrhage and intestinal inflammation.

Epidemiology

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The identification of the reservoir of the pathogen, the vector system it employs to move from this reservoir to a new host and the mode of entry into the new host are key questions in developing an understanding of the epizootiology of any disease. Unfortunately for A. salmonicida and its associated disease furunculosis, none of these questions can be answered with certainty.

Standard textbooks of bacterial taxonomy refer to A. salmonicida as an obligate fish pathogen (Bergey, 1984), suggesting that the bacterium is incapable of sustained growth outside its piscine hosts. A number of laboratory microcosm studies have suggested short survival times for the bacterium in aquatic environments. It should, however, be noted that such experimental systems are notoriously difficult to work with and the data they generate should only be extrapolated to the real world with great caution (Enger, 1997). It is also true that there are few reports of the isolation of the bacterium from the non-piscine environment. However, colony formation by A. salmonicida can be inhibited by a number of bacteria common in the aquatic environment, and Austin and Austin (1993) have argued that the failure to culture A. salmonicida from water may be a function of the inadequacy of our methods, rather than the absence of the organism. A second factor that may have contributed to our failure is the extreme hydrophobicity of the bacterium. This surface property would suggest that, in nature, A. salmonicida is more likely to be isolated from particles (Sakai, 1986), making traditional culture techniques unsuitable for the organism from the environment. Michel and Dubois-Darnaudpeys (1980) demonstrated that A. salmonicida can survive in a virulent form for many months in a sterile water microcosm and there is some indirect evidence that, at least in marine sediment, the organism can remain virulent for up to 6 months (Smith et al., 1982). In natural fresh water, survival is highly matrix dependent and is very short in humic acid-rich sediments (Hiney et al., 2002). In a survey of a freshwater hatchery, in which they used both ELISA and a PCR/DNA probe assay, Hiney et al. (1997a) found no correlation between positive detection of the signs of A. salmonicida in hatchery sediment and the disease status of hatchery stocks over a 2-year period, as assessed by kidney culture and stress testing. These non-culture-based assays are, however, capable of detecting non-culturable-but-viable cells and dead cells, and the results generated by them cannot be used, therefore, as an indication of virulence.

Covert infection by typical Aeromonas salmonicida

The existence of covert furunculosis, that is, clinically inapparent infections, has been recognized almost as long as the disease itself (Plehn, 1911). The epizootiological importance of fish with covert furunculosis in the maintenance and spread of the disease within and between susceptible fish populations was understood by early workers. In 1935 the Scottish Furunculosis Committee concluded that clinically inapparent infections by A. salmonicida could persist in fish populations, that these infections could be latent and that clinical furunculosis could be precipitated in covertly infected fish populations by stress. In addition, fish with covert infections were capable of acting as carriers and could shed sufficient bacteria to transmit the infection to other fish (Mackie et al., 1930, 1933, 1935). However, despite the large body of work that exists on covert furunculosis, a number of important questions on the nature, persistence, location, lack of clinical signs, host defence mechanisms and transmission of covert infections remain unanswered or have not been answered satisfactorily.

NATURE OF COVERT INFECTIONS

There is a paucity of information available on the nature of covert furunculosis, despite more than 100 years of research on A. salmonicida. The studies that have been performed have employed a variety of methods, of differing efficiency, to detect covert infections. In addition, the variety of names that have been applied to these infections makes it extremely difficult to present comparisons of these studies (Hiney et al., 1997b). In an effort to clear up some of the confusion that surrounds both the nomenclature and the exact nature of what is being studied, Hiney et al. (1997b) proposed a number of alternative definitions of clinically inapparent infections, strictly related to the diagnostic methods that have been used. Using this approach, three categories could be identified and all studies on covert infections were reassigned to one of these three categories:

1. Covert infection: demonstration of A. salmonicida, its antigens or its deoxyribonucleic acid (DNA) in, or on, a fish that manifests no clinical signs of furunculosis.

2. (Covert) carrier infection: demonstration of the shedding of A. salmonicida or its antigens or DNA into the environment by a fish that manifests no clinical signs of furunculosis; demonstration of the ability of fish that manifest no clinical signs of disease to transmit furunculosis to fish free of this disease, in cohabitation experiments.

3. (Covert) stress-inducible infection (stress-inducible furunculosis (SIF)): demonstration of clinical disease following the stressing of a fish that manifests no clinical signs of furunculosis.

It is apparent from these definitions that information on the location of A. salmonicida in covertly infected fish would do much to improve our understanding about the mechanisms of these types of infections. When considering covert infections, these clinically inapparent infections of fish may, in fact, represent a number of infection types, which are mediated by fundamentally different processes (Hiney et al., 1997b). However, the underlying processes of covert infection will remain unknown until we develop methods that can distinguish between different infection types in individual fish.

LOCATION OF AEROMONAS SALMONICIDA IN COVERT INFECTIONS

Despite almost 80 years of speculation, we have no certainty as to the location of A. salmonicida subsp. salmonicida in covertly infected fish. Convincing evidence exists for an external location of A. salmonicida on the mucous layer, on the gills and in the intestine during a covert infection (Klontz, 1968; Markwardt and Klontz, 1989b; Cipriano et al., 1992; Hiney et al., 1994). Cipriano et al. (1996b) successfully eliminated SIF by the topical administration of chloramine T and have isolated typical A. salmonicida from the mucus of apparently healthy fish (Cipriano et al., 1992, 1994, 1996a,c). The pathogenicity of mucus-isolated A. salmonicida has been demonstrated by Hiney et al. (1994), who could induce clinical furunculosis in disease-free brown trout by injection with a mixture of mucus and gill scrapings collected from Atlantic salmon with SIF.

An internal primary location of A. salmonicida in covertly infected fish has been proposed by a number of authors (cited by Hiney et al., 1997b). However, the experimental protocols employed in many of these studies make meaningful interpretation of the results difficult. In particular, experiments that involved the transport of fish would have been stressful and, when examined in the laboratory, fish that originally had a commensal infection might be experiencing the early phase of a clinical infection. Preliminary work carried out in Galway, Ireland, on the salmon intestinal mucosa suggests that bacteria resident in the intestine may breach this barrier following a brief transport stress. Thus, arguments for an internal location of A. salmonicida in fish must be viewed with caution in the absence of detailed descriptions of the protocols employed for handling and transport. Nomura and his coworkers in Japan have, however, presented evidence suggesting an internal location for A. salmonicida in covertly infected adult fish (Nomura et al., 1991a,b, 1992).

LACK OF CLINICAL SIGNS IN COVERT INFECTIONS

The lack of clinical signs of disease during a covert furunculosis infection is relatively simple to understand if the location of A. salmonicida is external to the fish. The clinical signs of furunculosis are those of a systemic bacteraemia and are unlikely to be present where the bacterium is confined to the gills, mucus or intestine. If, however, the location is internal, then the absence of disease becomes much more difficult to explain. A number of studies have demonstrated that the median lethal dose (LD50) of virulent A. salmonicida subsp. salmonicida strains for salmonids is between 102 and more than 10 colony-forming units (cfu), when injected either intramuscularly (i.m.) or intraperitoneally (i.p.) (McCarthy, 1977a; Cipriano et al., 1981; Cipriano and Starliper, 1982; Drinan and Smith, 1985; Olivier et al., 1985; Shieh, 1985; Bernoth and Körting, 1992). However, Scallan (1983) could isolate over 106 cfu of A. salmonicida from the kidneys of dead salmonids within 4 days of their being stressed. Knowledge of the possible mechanisms whereby A. salmonicida might avoid the fish’s internal defence systems during a covert infection would facilitate an understanding of the processes underlying these infections. In practically the only attempt to offer any explanation for the temporary reduction of virulence that would appear to be necessary if an internal location were accepted, McIntosh and Austin (1991) hypothesized that A. salmonicida might reside in fish as L-forms (spheroplasts) (McIntosh et al., 1991). They argued that the renal medulla of fish may provide a favourable niche for L-forms, because of the high electrolyte concentration in this tissue. This environment inhibits complement activity, and McGee (1986) has suggested that it may protect L-forms from the lethal effects of the antibody-complement system. However, infectivity studies with these naturally occurring A. salmonicida L-forms failed to produce disease, even after stressing experimentally infected fish (McIntosh and Austin, 1990).

PERSISTENCE OF COVERT INFECTIONS

Although persistence of covert infections has frequently been suggested, there is no definitive evidence that covert infections of individual fish can persist for extended periods. Early furunculosis research demonstrated the existence of persistent covert infections (Plehn, 1911), with no doubt that the duration of these infections had major epizootiological significance (Mackie et al., 1935). A study by Scallan (1983), involving the regular sampling of a population, would suggest that high frequencies of stress-inducible infections may persist in such populations for at least 1 year. However, studies of populations cannot provide information on the persistence of covert infections in individual fish over time. Where the parental population is maintained in a natural water body, the possibility of continual reinfections by A. salmonicida derived from the water cannot be eliminated. Persistence in a population may also result from repeated, overlapping, short-term infections in individual members of the population (Scallan, 1983).

In summary, a number of observations can be made about covert furunculosis infections. It is likely that covert infections are heterogeneous in nature and that there is, in theory, more than one form of covert infection, whose underlying processes differ fundamentally. Covert infections can be seasonal (Scallan and Smith, 1984, 1993) and transitory in occurrence (Andrews, 1981; Scallan et al., 1993). It is possible that they may persist for long periods in a population without obvious harm to the fish. Scallan (1983) also demonstrated, using a quantitative stress test, that the level of covert infection in a population varied throughout the year. Cipriano (1997) also reported that the numbers of A. salmonicida that could be isolated from the mucus of salmonids were variable over the growing season. Covertly infected fish may act as carriers of disease or be latently infected. Both internal and external locations have been proposed for covert infections, but, to date, the locations of A. salmonicida in fish suffering from commensal covert infections have not been resolved.

Disease transmission in ‘atypical’ strains

The transmission of atypical A. salmonicida has not been thoroughly investigated, due to a lack of understanding of the ecology of the organisms. Only a few reports describe some ecological aspects of atypical A. salmonicida. Evelyn (1971) reported that the atypical strain isolated from sablefish survived better in salt water compared with fresh water. In another study, Wiklund (1995a) investigated the survival of atypical strains isolated from flounders in microcosms. Best survival was observed in the presence of sediments and at high water temperature and the highest survival was observed in brackish water compared with fresh and salt water. Additional studies on the ecology of these strains is required if the epizootiology of these isolates is to be better understood.

In several cases, transmission of the disease seems to have been linked to the transfer of infected fish (Wichardt et al., 1989; Håstein and Lindstad, 1991). The best evidence has been provided by the example of the goldfish ulcer disease agent, which was introduced into Australia through the importation of live infected or subclinically infected goldfish in 1974. Following this introduction, the agent has been recovered from wild goldfish and the disease is now thought to be endemic in several areas of Australia (Whittington et al., 1987; Humphrey and Ashburner, 1993).

Impact Summary

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CategoryImpact
Fisheries / aquaculture Negative

References

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Distribution References

CABI, Undated. Compendium record. Wallingford, UK: CABI

CABI, Undated a. CABI Compendium: Status as determined by CABI editor. Wallingford, UK: CABI

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Tony Ellis
FRS Marine Laboratory, University of Aberdeen, Victoria Road, Aberdeen, Scotland, AB11 9DB, UK

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