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7 Infectious Diseases of Warmwater Fishin Fresh Water

Gilda D. Lio-Po1 and L.H. Susan Lim21Aquaculture Department, Southeast Asian Fisheries Development Center, Tigbauan,

5021 Iloilo, Philippines; 2Institute of Biological Sciences, University of Malaya,50603 Kuala Lumpur, Malaysia

Introduction

Cage culture of freshwater fish, whichbegan in Cambodia in the late 1800s, is nowcommonly practised in Southeast Asia andgaining popularity in India (Chapter 1). Indeveloping tropical countries, this type offish culture is still either at the subsistenceor semi-intensive level or is at the experi-mental stage, as for Chrysichthys spp. inAfrica (Aqua Farm News, 1993).

Fish cultured in cages in Southeast Asiainclude tilapia, carp, catfish, snakeheadsand eleotridids (Table 7.1). The tilapias, oneof the common species in freshwater cages,are also cultured in cages in warm marinewaters (Chapter 6) (Aqua Farm News, 1993).The catfish cultured include the Ictaluridae(Ictalurus spp.), Claridae (Clarias spp.),Pangasiidae (Pangasius spp.), Siluridae(Silurus glanis) and Bagridae (Hemibagrusspp.) (Aqua Farm News, 1993). Most catfishare of Southeast Asian origin, the exceptionbeing channel catfish cultured in theUSA, which have been introduced intocages in Indonesia (Rabegnatar et al., 1990).The most common catfish species culturedin cages in Southeast Asia is Pangasiushypophthalmus. Exotic Chinese carp,common carp (Cyprinus carpio), grasscarp (Ctenopharyngodon idellus), bigheadcarp (Aristichthys nobilis), silver carp

(Hypophthalmichthys molitrix) togetherwith Puntius gonionotus and Leptobarbushoevenii dominate the cyprinids (Table 7.1).Due to the variety of common names avail-able for a particular fish species in SoutheastAsia, the scientific names will be used asmuch as possible.

Publications and reports are availableon diseases of feral and cultured fish inwarm fresh water (Lio-Po, 1984; Kabata,1985; ADB/NACA, 1991; Lim 1991d, 1992;Paperna, 1991, 1996; Arthur, 1992; Thuneet al. 1993; Arthur and Lumalan-Mayo,1997; Fijan, 1999). However, there is apaucity of information on diseases of fishin freshwater cage culture, even thoughcage culture began in Southeast Asia(Chapter 1) (Christensen, 1989; Aqua FarmNews, 1993). Diseases are normally eithermentioned in passing or are not included,particularly in those publications dealingwith cage culture (Christensen, 1989;ADB/NACA, 1991; Dharma et al., 1992;Nasution et al., 1992; Alawi and Rusliadi,1993; Aqua Farm News, 1993). In addition,publications on diseases in fish culturedo not distinguish between diseasesfound in cage culture and pond culture(Davy and Chouinard, 1982; Arthur,1987; ADB/NACA, 1991; Aqua Farm News,1993). This is further exacerbated by thelack of comprehensive investigation into

©CAB International 2002. Diseases and Disorders of Finfish in Cage Culture(eds P.T.K. Woo, D.W. Bruno and L.H.S. Lim) 231

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diseases encountered in cage culture sys-tems in warm waters.

Disease outbreaks in cage culture have agreater impact because of high stocking

232 G.D. Lio-Po and L.H.S. Lim

Country Fish species References

Bangladesh

Cambodia

India

Indonesia

Malaysia

Philippines

Sri LankaThailand

Vietnam

Catla catlaCirrhina mrigalaCyprinus carpioHypophthalmichthys molitrixOreochromis niloticusChanna micropeltesCirrhinus microlepisLabeo sp.Clarias sp.Leptobarbus hoeveniiOxyeleotris sp.Pangasius sp.C. catlaC. mrigalaLabeo bataLabeo rohitaChanna striataOreochromis mossambicusC. striataOxyeleotris marmoratusTilapiaC. carpioAristichthys nobilisC. striataCtenopharyngodon idellusC. carpioHemibagrus nemurus (also knownas Mystus nemurus)H. molitrixL. hoeveniiO. marmoratusPuntius gonionotusTilapiaA. nobilisChanos chanosC. carpioH. molitrixO. niloticusO. niloticusClarias spp.C. carpioGoby sp.O. niloticusBarbus spp.Leptobarbus sp.C. striataO. marmoratusPangasius bocourtiPangasius conchophilus

Karim and Harun-al-Rashid Khan (1982)

Thana (2000)

Guerrero (1979)

Natarajan et al. (1983)

Sukumaran and Sanjeeviraj (1983)Jameson (1983)Indra (1982)

Jangkaru and Rustami (1979)Annual Fisheries Statistics (1998)

Palisoc (1988)

Siriwardena (1982)Tugsin (1982)

Pantulu (1979)

T.T. Dung (personal communication)

Table 7.1. Freshwater fish species cultured in cages in some tropical countries.



densities and close proximity of culturedfish with each other as well as with feral fish.For example, there were 64 reported diseaseoutbreaks in cage-cultured channel catfishin the USA during 1990 with mortalityin 91% of these cases (Masser et al.,1991). Also, diseases appear to occur morefrequently in cages than in ponds (Collins,1988). Cage culture exposes fish topathogens of feral fish and perhaps to agreater number of intermediate hosts inparasitic diseases. Fish reared in cages mayalso present a potential health threat toman, especially when they are reared inunsanitary waters in areas where fish-bornezoonotic diseases are prevalent (seeKo, 1995) or when located in pollutedareas. Diseases afflicting pond-reared andcage-cultured fish are in most cases similar,hence those that are important in pondaquaculture will be treated as potentialproblems for the cage culture. For example,Piscinoodinium pillulare, the causativeagent of velvet disease of cyprinids, was firstreported on pond-reared fish but is nowfound on cage-cultured fish (F. Shaharom,personal communication). The paucity ofinformation on diseases in cage-culturedfish is partly due to the lack of studies onidentification of pathogens/disease mecha-nisms and/or the absence of mandatoryreports on disease outbreaks in manycountries. Hence, we expect diseases tobecome more prevalent in the future as wemove into more intensive fish culture,find out more about infectious agents, andadopt a system where it is mandatory toreport disease outbreaks. In the currentreview, we have also included unpublishedinformation from colleagues as well asfrom personal observations, and whereverpossible we have provided the correct iden-tification of pathogens and supplementaryinformation on them.

Viral Infections

Viral infections can cause mass mortality,especially in fry or fingerlings, while olderfish may develop resistance or are hardly

affected. Most viral infections occur infish at low water temperatures. This mayexplain the paucity of viral infectionsrecorded in warm freshwater fish. Stressfrom handling, poor water quality, watertemperature, age of fish, high stockingdensity and poor nutrition are factorsthat facilitate the development of viraldiseases.

Among viral infections in fish, thechannel catfish virus disease has the mostimpact on cage culture while the grass carphaemorrhagic virus and the spinning tilapiasyndrome are also potential viral problems.In addition, other viral epizootics have beenreported in common carp and tilapia thatmay have implications in fresh warmwatercage culture systems (Sano et al., 1993;Oyamatsu et al., 1997; Fijan, 1999).The epizootic ulcerative syndrome (EUS),a disease associated with a rhabdovirus,bacteria and the pseudofungi, Aphanomycesinvadans, is discussed in the section onDiseases of Complex Infectious Aetiology.

An insufficient number of susceptiblefish cell lines hampers isolation and diagno-sis of viral pathogens. Cell lines currentlyused for isolation of warm freshwatervirus are from: bluegill fry (BF-2) (Wolfand Quimby, 1966), brown bullhead (BB),channel catfish ovary (CCO) (Bowser andPlumb, 1980), Epithelioma papulosumcyprini (EPC) (Fijan et al., 1983), grass carpkidney (GCK-84), grass carp gonad (GCG),grass carp fin (GCF) (Wolf, 1988), rainbowtrout gonad (RTG-2), snakehead fry (SSN-1)(Frerichs et al., 1993), catfish spleen (CFS)and snakehead spleen (SHS) (Lio-Po et al.,1999).

Electron microscopy for the diagnosisof viral infections is not commonly useddue to inaccessibility to this equipment inmost tropical countries. As an alternative,serological tests are applied such as neutral-ization index determination, Western blot,ELISA, fluorescent antibody technique(FAT) and indirect fluorescent antibodytest (IFAT). Recent molecular biologytechniques such as PCR, RT–PCR andgene probes are becoming popular for thediagnosis of fish viral infections.

Infectious Diseases of Warmwater Fish in Fresh Water 233



Channel catfish virus disease (CCVD)

Channel catfish (lctalurus punctatus) is theprincipal host of channel catfish virus(CCV). Outbreaks occur in most southernstates in the USA, while low-grade mortal-ity can be induced in blue catfish (Ictalurusfurcatus) and channel catfish × blue catfishhybrids by experimental injection (Plumbet al., 1975).

Pathology. CCV causes acute infection incultured channel catfish fry and fingerlingsless than 10 cm in length. It can also infectchannel catfish juveniles and adults follow-ing waterborne exposure to CCV (Plumb,1971; Hedrick et al., 1987). Clinical signsare abdominal distension, exophthalmia,pale or haemorrhagic gills and petechialhaemorrhage at the base of the fins andthroughout the skin (Fig. 7.1). Infected fishswim erratically at the surface in head-highor hanging position. Mortality approaching100% in channel catfish younger than4 months old occurs at water temperaturesabove 25°C within 7–10 days. The virus doesnot induce mortality below 15°C. Secondaryexternal lesions caused by bacteria, e.g.Flavobacterium columnare or Aeromonashydrophila, or by aquatic stramenopiles maydevelop.

CCVD develops into a haemorrhagicviraemia after replicating in the kidney

and in the spleen. Thereafter, the virus istransported via the blood to the intestine,liver, heart and brain (Plumb and Gaines,1975). Thus, hyperaemia of the visceralcavity, enlarged spleen, and empty stomachand intestine have been observed (Plumb,1994). Necrosis of the renal haematopoietictissue and tubules, oedema, necrosis andcongestion of the liver, intestinal oedemaand congestion and haemorrhage in thespleen are characteristic histopathologicalfindings. Skeletal muscle haemorrhage isseen in experimentally infected fish. Thevirus can be isolated from the kidney, intes-tine, liver, spleen, brain and muscle tissues(Plumb, 1971; Plumb and Gaines, 1975).The portal of entry for CCV from water isthrough the gills and the gut (Nusbaum andGrizzle, 1987). Channel catfish survivinga CCV infection grow slowly; e.g. experi-mentally induced CCVD survivors rangedfrom 11 to 15 g compared with 73–93 g inunexposed channel catfish 6 months after astandardized feeding regime (McGlameryand Gratzek, 1974).

The virus remains viable in deadfish kept on ice for 14 days and at −20°Cfor 100 days (Plumb et al., 1973). Itremains infective for 2 days in pond waterat 25°C and for 11 days in dechlorinatedtap water. However, it is rapidly inacti-vated in pond mud and by drying (Plumb,1994).


Fig. 7.1. Channel catfish (Ictalurus punctatus) infected with the channel catfish virus (CCV) (courtesy ofDr John Plumb).



Transmission of CCV occurs bothhorizontally and vertically. The virus isreadily transmitted from fish to fish. Theexact mode of transmission is unknown butis most likely through the branchial andintestinal epithelium. Upon intraperitonealinjection, the virus is detected in the kidneyafter 24–48 h, in the intestine and liver after72–96 h and in the brain after 96–120 hpost-injection (Plumb, 1971). In experi-mental infections, fry die within 3 daysof exposure (Wolf, 1988). The virus alsooccasionally persists in apparently healthyadult channel catfish broodfish but in mostcases CCV cannot be isolated from adultfish, and has been isolated from fingerlingsin only two of seven farms with positivebroodfish (Bowser et al., 1985).

Diagnosis. CCV, designated as Herpesvirusictaluri, is a herpesvirus of the familyHerpesviridae (Wolf and Darlington, 1971).It is enveloped, with icosahedral symmetryand measures 90–100 nm in diameter.The virus can be isolated from the kidneyof fish with active infections using CCOor BB cells. Inoculated cells developcytopathic effects (CPE) 24–48 h post-exposure, with optimal viral replication at25–30°C. Identification is confirmed usingelectron microscopy, serum neutralizationtests, IFAT, ELISA using monoclonal anti-bodies, CCV DNA probes and PCR (Wiseet al., 1985; Office International desEpizootie (OIE), 1995; Baek and Boyle,1996).

Prevention and control. Detection of CCVin catfish broodstock will help preventits spread to young catfish. The use ofvirus-free stock is the best preventivemethod. Alternatively, the use of resistantfish stocks or hybrids of channel catfishis recommended. Quarantine and killing ofCCV-infected stock including surveillancefor feral fish carriers should be practised.This should be of utmost considerationbefore introduction of channel catfish intotropical countries. Vaccination is still atthe experimental stage and there is nochemotherapy.

Grass carp haemorrhagic disease (GCHD)

The disease was first observed in Chinain the 1980s. It commonly affects grasscarp but can also infect black carp(Mylopharyngodon piceus), topmouthgudgeon (Pseudorasbora parva) and rareminnow (Gobiocypris rarus). It can alsoreplicate in silver carp and in Chineseminnow (Hemiculter bleekeri) without anyclinical signs. Outbreaks occur in SouthernChina during the summer when watertemperatures range from 24 to 30°C (Nieand Pan, 1985; Wolf, 1988; Jiang, 1995;Fijan, 1999).

Pathology. Acute infections cause sig-nificant mortality of more that 80% infingerlings and up to 70% in yearlings.Clinical signs include exophthalmia andsevere haemorrhage of the gills and finbases. Internally, haemorrhage occurs in themusculature, oral cavity, intestinal tract,liver, spleen and kidney. Naturally andexperimentally infected fish have reducederythrocytes, plasma protein, calcium andurea nitrogen but serum potassium iselevated. Experimental infection by bathand by injection induced typical signsof the infection. Disease and mortality areobserved within 1–2 weeks exposure of fishin water at temperatures of 25°C or higher(Wolf, 1988; Fijan, 1999).

Diagnosis. The grass carp haemorrhagicvirus (GCHV) is a non-enveloped, doublyencapsidated icosahedron with 5:3:2 sym-metry, 92 capsomeres, with an overalldiameter of 60–80 nm and a 40 nm innercapsid (Wolf, 1988). It is resistant to etherand chloroform. It is presently classifiedunder the genus Aquareovirus (FamilyReoviridae) (Li et al., 1997). The virus canbe propagated in GCK-84, GCG and GCFcells yielding titres of 108–109 TCID50 ml−1.In vitro viral replication is optimum at28–30°C inducing CPE in 3–4 dayspost-inoculation (Wolf, 1988). The virus infish with clinical signs and in carrier fishcan be confirmed using RT–PCR (Li et al.,1997).




Prevention and control. Experimental vac-cination using inactivated virus induced80% protection by day 4 at temperaturesabove 20°C, by day 20 at 15°C, and by day30 at 10°C, and this protection lasts for upto 14 months (Wolf, 1988). Zhu et al. (1993)as cited by Fijan (1999) reported that the‘Kelieao–Yufukang’, a combination of twodrugs, has in vitro and in vivo anti-GCHVactivity.

Spinning tilapia (ST) syndrome

This virus was recently detected in Moz-ambique tilapia, blue tilapia (Oreochromisaureus), Nile tilapia (Oreochromis nilo-ticus), and mango tilapia (Sarotherodongalilaeus) in Australia. The disease iscaused by an iridovirus (Ariel and Owens,1997).

Pathology. Affected tilapia fry swim in aspiral pattern, sink to the bottom then riseand hang at a 45° angle just under the watersurface, gasping for air. They do not feed, aredarker in colour and exhibit ‘fin clamping’.Tilapia fry manifesting the spinning syn-drome die within 24 h and a 100% mortalityoften occurs within 60 days. Naive tilapiafry experimentally exposed to diseased fryvia cannibalism developed similar signsafter 12 days. Histopathologically, the renaltubules are shrunken, haemorrhaging andinfiltrated with eosinophilic granular cells.In addition, focal myolysis occurs inmuscles. These histopathological lesionsand the size range (110–140 nm) of the virusare similar to those caused by the Bohleiridovirus (BIV) in tilapia fingerlings (Arieland Owens, 1997). The Bohle iridovirus alsoinfects amphibians (Cullen et al., 1995).

Diagnosis. So far, the virus has not beenisolated in cell culture from diseased tilapiabut the disease is usually diagnosed basedon clinical signs and is confirmed byelectron microscopy.

Prevention and control. No treatment isavailable but prevention through quarantine

and restriction of transfer of stocksfrom endemic to non-endemic areas isrecommended.

Bacterial Diseases

High stocking density of fish leads toincreased feed rations and waste. Thisalso results in bacterial problems with con-comitant increases in ammonia and nitritetoxicity (Mitchell, 1997). Stress and traumafrom handling are also predisposing factors.Most bacterial pathogens produce enzymesthat facilitate their entry/invasion into thefish host tissues. Although they may causeprimary infection, they may also act assecondary disease agents to a primary virusor parasite. The major bacterial infectionsamong warm freshwater fish are motileAeromonas septicaemia, Pseudomonas sep-ticaemia, edwardsiellosis, enteric septicae-mia, columnaris disease and streptococcalsepticaemia/meningoencephalitis.

Motile Aeromonas septicaemia (MAS)

This disease was formerly known as haem-orrhagic septicaemia, infectious dropsy,infectious abdominal dropsy, red pest, reddisease, red sore or rubella. The syndromeis caused by the motile A. hydrophila(previously named Aeromonas punctataor Aeromonas liquefaciens). Aeromonassobria and Aeromonas caviae are rarelyassociated with fish epizooties.

MAS affects freshwater and occasion-ally brackishwater and marine warmwaterfish worldwide. It is the most frequentlydiagnosed bacterial fish disease and was themost severe disease problem among cage-cultured channel catfish in the USA between1972 and 1980 (Plumb, 1994). Subsequently,it became the third most common bacterialinfection (1987–1991) among cage-culturedchannel catfish in the USA, accounting for13–22% of disease outbreaks (Duarte et al.,1993). The infection occurs mostly from Feb-ruary to July, with some outbreaks occurringin September and November. In the tropics,




MAS infections are often reported in pond-and pen-cultured milkfish (Chanos chanos),common carp, grass carp, Nile tilapia andgiant gourami (Osphronemus goramy) (Kou,1972; Ruangpan et al., 1985; Karunasagaret al., 1986; Lio-Po et al., 1986; Saitanu et al.,1986; Supriyadi, 1986; Areerat, 1987; Angkaet al., 1988; Okaeme et al., 1989; Yambot,1997). Moreover, A. hydrophila has beenassociated with epizootic ulcerative syn-drome (EUS)-affected striped snakeheads(Channa striata, also known asOphicephalus striatus) and walking catfish(Clarias batrachus) in the wild, as well asin ponds and cages (Llobrera and Gacutan,1987; Boonyaratpalin, 1989; Subasingheet al., 1990; Torres, 1990; Lio-Po et al., 1992;Pathiratne et al., 1994; Angka et al., 1995;Karunasagar et al., 1995; Thanpuran et al.,1995; Rahman et al., 1999).

Pathology. A. hydrophila is a free-living,mesophilic bacterium found in soil, fresh-water lakes, ponds, streams, bottom mud,domestic tap water and sewage. It is oftenassociated with the normal flora of fish.Thus, the bacterium has been isolated fromboth healthy and diseased fish (Lio-Po andDuremdez-Fernandez, 1986; Lio-Po et al.,1986, 1992; Torres, 1990). It causes infec-tions not only in aquatic animals but also inavian hosts, cows and humans.

Infected fish lose their appetite, becomelethargic and swim near the surface. Exter-nal signs may vary according to fish speciesbut are generally similar to clinical signsof other bacterial septicaemia infectionsin fish, i.e. exophthalmia and distendedabdomen. However, septicaemia in acuteMAS can be fatal with no clinical signs.Among milkfish reared in pens in a fresh-water lake in the Philippines, acute signs ofpetechial haemorrhage of the skin and finbases including dermal and caudal fin rotwere observed (Lio-Po et al., 1986). Yambot(1997) also isolated A. hydrophila fromcage-cultured tilapia in the Philippines withhaemorrhagic skin, ulceration, loss of scales,mouth sores, eye abnormalities, fungalgrowth and/or tail and fin rot. Fingerlingsto adult Nile tilapia can be infected byA. hydrophila (Yambot, 1997). A case of

natural infection in tilapia caused byA. hydrophila was associated with epider-mal lesions so severe that the vertebraewere exposed (Lightner et al., 1988). Thissevere condition is not uncommon amongEUS-affected fish (Roberts et al., 1994b).Hence, it is not surprising that the bacteriumhas been consistently isolated from EUS-affected fish (Llobrera and Gacutan,1987; Boonyaratpalin, 1989; Costa andWejeyaratne, 1989; Subasinghe et al., 1990;Torres, 1990; Lio-Po et al., 1992; Pathiratneet al., 1994; Angka et al., 1995; Karunasagaret al., 1995; Thanpuran et al., 1995; Rahmanet al., 1999).

In channel catfish, A. hydrophilainfection has three categories: (i) motileaeromonad septicaemia with external signs;(ii) cutaneous, manifesting lesions that arelimited to the skin and underlying muscle;and (iii) latent septicaemia with no externalsigns (Grizzle and Kiryu, 1993). Internalclinical signs include oedema, haemorrhageand necrosis. The disease is acute in veryyoung fish while adults generally developchronic infections (Plumb, 1994).

Motile aeromonad infections are predis-posed by stress from temperature shock, lowdissolved oxygen, high ammonia, handlingor hauling, and an ongoing primary infection(Plumb et al., 1978; Lio-Po et al., 1986).These predisposing conditions are possiblyimmunodepressive, and the virulence ofthe Aeromonas strain is an importantfactor in the development of MAS epizootics(Thune et al., 1993). Moreover, A.hydrophila is often reported in mixedinfections with Edwardsiella tarda, E.ictaluri, Flavobacterium columnare, Strep-tococcus spp. or with parasites (Kanai et al.,1977; Liu et al., 1990; Duarte et al., 1993). Incarp dropsy, A. hydrophila was a compo-nent in the pathology of the disease, which isattributed to a virus as prime aetiologicalagent (Roberts, 1993).

Experimental A. hydrophila infectionsmay be induced in milkfish with up to100% mortality in 2 days after immersionexposure of scarified fingerlings, but not infish with intact skin (Lio-Po and Duremdez-Fernandez, 1986). In addition, intraperi-toneal injection with the bacterium causes




mortalities within 12 h of injection of thepathogen (Lio-Po and Duremdez-Fernandez,1986). In walking catfish and snakeheads,A. hydrophila induced dermal lesions afterintramuscular injection of at least 105 cellsper fish, which eventually ulcerated (Fig.7.2) (Lio-Po et al., 1992). Compared withother bacteria associated with EUS lesions,such as Pseudomonas sp., Aquaspirillum sp.and Streptococcus sp., A. hydrophilainduced the most severe lesions upon intra-muscular injection of snakeheads (Lio-Poet al., 1998). Experimental infection of Niletilapia fingerlings by immersion yielded anLD50 of 1.5 × 106 colony-forming units (cfu)ml−1 with 100% mortality at 108 cfu ml−1 andno mortality at 103 cfu ml−1 (Yambot, 1997).Attempts to induce external gross lesions inwalking catfish by dermal cut, dermal scrap-ing, fish bite, oral feeding, gastric lavageand cohabitation with a golden snail carrier(Ampullarius sp.) were unsuccessful (Lio-Poet al., 1996). In contrast, in channelcatfish with mechanically abraded skin,A. hydrophila experimentally induced sys-temic infection in 80% of exposed fish whilecutaneous lesions developed in the remain-ing fish (Matsche and Grizzle, 1999).

Histopathologically, marked necrosis ofthe muscle fibrils occurred in snakeheadsintramuscularly injected with A. hydrophila(Lio-Po, 1998). Walking catfish injected

intraperitoneally with A. hydrophila dev-eloped focal necrosis in the liver, kidney,intestine and dorsal musculature (Angka,1990). The infection elicits an intenseinflammatory response, with massiveinfiltration of monocytic and granulocyticcells into infected tissues (Huizinga et al.,1979; Ventura and Grizzle, 1988). Infectedgoldfish are anaemic, e.g. low red bloodcell, haematocrit and haemoglobin counts(Brenden and Huizinga, 1986). In addition,there is a shift in the differential countsof lymphocytes to a predominance ofneutrophils.

Motile aeromonads secrete extracellularproducts (ECPs) and these include toxins,protease, cytotoxin, haemolysin, leuco-cidin, gelatinase, elastase, staphylolysin,caseinase, enterotoxin and a dermonecroticfactor (Hsu et al., 1981; Olivier et al., 1981;Kanai and Wakabayashi, 1984; Lallier et al.,1984; Krovacek, 1989; Yadav et al., 1992).Moreover, cytotoxin-producing strains wereassociated with EUS-affected fish (Yadavet al., 1992). Dermonecrotic strains of thisbacterium secrete haemolysin at 10 and 30°C(Olivier et al., 1981). However, correlationbetween virulence and ECP production wasnot consistent (Leaño et al., 1996). Recently,Cascon et al. (2000) described the molecularcharacteristic of an elastase secreted byA. hydrophila that is important in its


Fig. 7.2. Catfish (Clarias batrachus) showing ulcerative lesions 4 days post-intramuscular injection withAeromonas hydrophila.



pathogenicity. This protease has a highamino acid sequence similarity to proteasessecreted by Pseudomonas aeruginosa,Helicobacter pylori and Vibrio spp. Aninvestigation of 12 strains of A. hydrophilaisolated from fish showed that protease pro-duction varied among strains, with peak pro-tease production optimum at 27.6 ± 4.9°C.(Uddin et al., 1997). The protease levelsincreased during the late log phase to earlystationary phase.

Siderophore production is also descri-bed but this is not related to virulence of thebacterium (Santos et al., 1988; Leaño et al.,1995). Virulence of A. hydrophila varies,even among isolates from the same epizootic(Lio-Po et al., 1992). Also, Rahman et al.(1997) reported that A. hydrophila stored in0.60 and 0.85% NaCl solutions had highervirulence than the cultured bacterium wheninjected intraperitoneally into carp andgoldfish. Subsequent studies showed thata significantly higher number of starved A.hydrophila adhered to the skin of cruciancarp (Carassius carassius) than the culturedbacterium (Rahman and Kawai, 1999).Attachment of A. hydrophila to carp epithe-lial cells is attributed to a 43 kDa outermembrane protein adhesin because of anabundance of this particular receptor on thecell surface (Lee et al., 1997). Dooley et al.(1986) earlier described a crystalline surfacelayer, or S-layer, of 52 kDa protein, andfurther correlated this to strain virulence(Murray et al., 1988; Ford and Thune, 1991).

Diagnosis. Motile Aeromonas spp. are flag-ellated, Gram-negative, short rods. Theydo not produce pigments and are resistantto vibriostat 0/129 (2,4-diamino-6,7-diisopropylpteridine phosphate). The bacte-ria grow at a temperature range of 18–39°C(Uddin et al., 1997). In tryptic soy agar (TSA)or in brain heart infusion agar (BHIA) at25–30°C incubation for 24–48 h, Aeromonasspp. produce white to creamy, convex, moistcolonies. In Rimler–Shotts medium, thebacteria form orange–yellow colonies at35°C (Shotts and Rimler, 1973). The threeimportant Aeromonas spp. in fish canbe differentiated using biochemical tests(Lio-Po et al., 1992; Plumb, 1994). Joseph

and Carnahan (1994) have further classifiedthese into seven species (Table 7.2). Mol-ecular identification can be applied byribotyping of restriction genomic DNAs ofaeromonads using different fragments of the16S rDNA gene of Escherichia coli as a probe(Lucchini and Altwegg, 1992). Also, ampli-fied fragment length polymorphism (AFLP)as a high-resolution genotype tool for classi-fication of Aeromonas spp. and pulse-fieldgel electrophoresis as a rapid technique fortyping of A. hydrophila have been devel-oped (Huys et al., 1996; Talon et al., 1996).Igbal et al. (1998) furthur recommend theapplication of genetic identification usingDNA–DNA hybridization.

Both virulent and non-virulent strainshave been isolated from diseased fish(Torres, 1990; Lio-Po et al., 1992; Leañoet al., 1996). Definitive identification ofeither strain is a major difficulty and hasbeen the subject of a number of researchefforts. Cartwright et al. (1994) developedmonoclonal antibodies for detection of viru-lent strains using either ELISA or fluoresceinisothiocyanate (FITC) immunofluorescence.Virulent strains of A. hydrophila requireabout 30 min to induce CPE in EPC cells,while avirulent strains do not induce thispathological effect (Leung et al., 1996). APCR method that is reported to be rapid,sensitive and specific for the detection ofvirulence factors of Aeromonas spp. hasbeen developed (Bin Kingombe et al., 1999).Recently, identification of the genetic differ-ences and virulence genes among differentstrains of A. hydrophila using a suppressionsubtractive hybridization (SSH) techniquewas reportedly successful (Zhang et al.,2000).

Prevention and control. MAS outbreaks arecommon in eutrophic lakes and ponds.Outbreaks of A. hydrophila infections infish-pen-reared milkfish are usually relatedto transport and handling stress, adverseenvironmental conditions of low oxygenconcentration, low pH, and increased levelsof ammonia and carbon dioxide (Waltersand Plumb, 1980; Lio-Po, 1984; Lio-Po et al.,1986). Moreover, tilapia fingerlings duringseining can get caught between nets and




subsequently develop MAS causing morethan 25% mortality after stocking in cages(J.A. Plumb, personal communication).Therefore, prevention of these stressfulconditions will minimize MAS outbreaks.

Limited success has been achievedwith vaccination against MAS in milkfish(G.D. Lio-Po, unpublished data) but vaccina-tion is protective in tilapia (Ruangpan et al.,1985). Indian major carp vaccinated with A.hydrophila yielded increased agglutinatingantibody titre (Karunasagar et al., 1991).Vaccination by intraperitoneal injectionof formalin-killed A. hydrophila to catfish

(Clarias macrocephalus) is more effectivethan immersion or oral administration(Areechon et al., 1992). Biofilm vaccineat 1013 cfu g−1 of A. hydrophila in catla(Catla catla), rohu (Labeo rohita) and com-mon carp for 15–20 days elicited high serumantibody titre and protective responsefor 60 days (Azad et al., 1999). Bluegourami (Trichogaster trichopterus), whenintraperitoneally immunized with majoradhesin (43 kDa) in Freund’s completeadjuvant, developed protective immunity tochallenge by homologous and heterologousstrains of A. hydrophila and one virulent


Resulta for:

CharacteristicA. hydrophila

(n = 46)

A. veroniibv. sobria(n = 26)

A. veroniibv. veronii

(n = 8)A. caviae(n = 33)

A. schubertii(n = 6)

A. jandaei(n = 9)

A. trota(n = 13)

Esculin hydrolysisVoges–ProskauerreactionPyrazinamidaseactivitycAMP-like factor(aerobic only)Fermentation

ArabinoseMannitolSucrose

SusceptibilityAmpicillinCarbenicillinCephalothinColistinb

DecarboxylaseLysineOrnithine

Arbutin hydrolysisIndoleH2Sc

Glucose (gas)Haemolysis (TSAwith 5% sheeperythrocytes)

++

+

+

V++

RRRV

+–+++++

–+

–

+

–++

RRSS

+––++++

++

–

+

–++

RRSS

+++++++

+–

+

–

+++

RRRS

––++––V

–V

–

–

–––

RRSS

+–––––+

–+

–

V

–+–

RRRR

+––++++

––

–

–

–+–

SSRS

+–V+++V

a+, positive for > 70% of isolates; –, negative, i.e. positive for < 30% of isolates; V, variable; R, resistant;S, susceptible.bMIC (single dilution), 4 µg ml−1.cH2S from GCF medium.

Table 7.2. Comparison of distinguishing profiles of mesophilic clinical Aeromonas species (reprintedfrom Annual Review of Fish Diseases, Vol. 4, Joseph and Carnahan, 1994, with permission from ElsevierScience).

250Z:\Customer\CABI\A4337 - Woo\A4419 - Woo Vouchers.vpTuesday, November 12, 2002 3:12:20 PM


strain of Vibrio anguillarum (Fang et al.,2000). Vaccination studies in carp immu-nized with crude lipopolysaccharideshowed that the mechanism of immunityis attributed to a sensitized thymocyte–macrophage system (Baba et al., 1988a,b).

Recently, immersion vaccination ofcarp with A. hydrophila bacterins showed adistinct increase of lysozyme level in fishmucus, with stronger bacteriolytic proper-ties 7 and 28 days after immunization(Kozinska, 2000). Moreover, a single intra-peritoneal injection of 20 mg β(1,3)-D-glucankg−1 into blue gourami enhanced theimmune response against A. hydrophila forup to 29 days (Samuel et al., 1996). Subse-quent studies showed that the use of fourglycans, namely Bar (glycan extracted frombarley), krestin, scleroglucan and zymosan,significantly increased survival rates oftilapia and grass carp after infection withA. hydrophila (Wang and Wang, 1997).

Prophylactic bath treatments with1–3% NaCl will help reduce post-handlinginfections. Likewise, bath treatments with2–4 mg potassium permanganate l−1 are alsoeffective for external lesions. Medicatedfeed with 2–4 g oxytetracycline kg−1 feed(50–100 mg kg−1 fish) for 14 days isrecommended (Plumb, 1994). However,drug-resistant strains of A. hydrophila mayevolve (Aoki, 1999).

Pseudomonas septicaemia

Pseudomonas spp. are ubiquitous in waterand are opportunistic pathogens. In fresh-water culture systems, Pseudomonasfluorescens has been implicated inepizootic outbreaks in Nile tilapia, grasscarp, silver carp and bighead carp (A.nobilis) (Miyashita, 1984; Lio-Po andSanvictores, 1987; Thune et al., 1993).

Pathology. The clinical signs in fishaffected with Pseudomonas septicaemia arevery similar to those with MAS. Gross signsinclude ascites, exophthalmia, septicaemiaand ulcers. The infection may be acuteor chronic, with the latter commonly

associated with skin lesions. Histopatho-logical findings in Nile tilapia include focalnecrosis, abscess and granulomas in theeyes, gills, liver, swim-bladder, kidneyand spleen (Miyashita et al., 1984). The bac-terium also causes mortalities in 2-week-old Nile tilapia fry (Lio-Po and Sanvictores,1987).

P. fluorescens is part of the normal floraof tilapia gut (Sugita and Kadota, 1980).It remains viable in fresh water for up to150 days (Duremdez and Lio-Po, 1984) andsecretes an extracellular proteinase (Li andFleming, 1967).

Diagnosis. As clinical signs of Pseudo-monas septicaemia resemble those of MAS,isolation and identification of the bacterialpathogen is required. P. fluorescens is aGram-negative rod with one to three polarflagella. It grows on nutrient agar, Pseudo-monas F agar and blood agar (Austin andAustin, 1987). For strains pathogenic tofish, the optimum growth temperature is20–25°C. These secrete oxidase, catalase andgelatinase but not amylase, galactosidase,urease or hydrogen sulphide. It is citrate-positive, oxidative for glucose and producesa fluorescent pigment (Plumb, 1994).

Prevention and control. Stress from lowdissolved oxygen concentrations, highstocking density, physical trauma and poornutrition are predisposing factors in thedevelopment of Pseudomonas septicaemia(Post, 1983). Therefore, avoidance of theseconditions is necessary in the preventionof its outbreak. Suggested bath treatmentsduring the early stage of the disease include1–2 mg benzalkonium chloride l−1 for 1 h,0.5–1 mg furanace l−1 for 5–10 min or1–5 mg malachite green l−1 for 1 h (Austinand Austin, 1987).

Edwardsiellosis and enteric septicaemia

E. tarda is synonymous to Paracolobactrumanguillimortiferum and to the E. anguilli-mortiferum described by Wakabayashiand Egusa (1973) and Kuo et al. (1987),




respectively. Edwardsiellosis has beenreported in 25 countries worldwide (Austinand Austin, 1987). The disease affects eels(Wakabayashi and Egusa, 1973), channelcatfish (Meyer and Bullock, 1973), mullet(Kusuda et al., 1976), tilapia (Lio-Po et al.,1982), carp (Sae-Oui et al., 1984) andstriped bass (Herman and Bullock, 1986).Although, there are no reports of cage-cultured fish affected by edwardsiellosis, itposes a health threat. E. tarda can also causeserious infections in humans.

Enteric septicaemia is attributed toE. ictaluri in freshwater fish. This is a majorpathogen of cage-reared channel catfishand accounts for about 30% losses in thesoutheastern USA. The estimated annualloss attributed to this pathogen is US$20–30million (Plumb and Vinitnantharat, 1993).The majority of cases occur in May and Juneand again in September and October whenwater temperatures are between 22 and 28°C(Plumb and Schwedler, 1982). A morbidityrate as high as 68% was observed in May1987, and an estimated 10–32% yearlymorbidity rate among the primary diagnosticcases of cage-cultured and pond-culturedchannel catfish in 1987–1991 (Duarte et al.,1993).

E. ictaluri has been reported inThailand, the USA and Australia. It ispathogenic to channel catfish but onlyvery slightly pathogenic to blue catfish.White catfish (Ictalurus melas) and brownbullhead (Ictalurus nebulosus) are occasion-ally infected, while natural infections ofwalking catfish have been reported inThailand (Plumb, 1994). Information onthe pathology, epizootiology, diagnosis,prevention and control of Edwardsiella isdetailed in Chapter 4 and in Plumb (1999).

Columnaris disease

Columnaris disease is an acute to chronicinfection of freshwater fish and a commonbacterial infection in the southeasternUSA (Duarte et al., 1993; Mitchell, 1997).Outbreaks are from March to Septemberwith peaks in June, and usually followoutbreaks of other diseases. The disease

occurs in Asia, the USA and Europe,affecting warmwater fish like channelcatfish and other ictalurids, cultured eels,common carp and tilapia (Plumb, 1994).The pathogen is Flavobacterium colum-nare, formerly called Flexibacter colum-naris, Cytophaga columnaris, Chondrococ-cus columnaris and Bacillus columnaris.

Pathology. Infection primarily begins at themouth, fins and gills. Clinical signs includefrayed fins with greyish to white margins,depigmented, necrotic skin lesions withyellowish or pale margins, which candevelop into shallow ulcers, yellowishmucoid material at the mouth and light todark brown gill discoloration. Gill lesionsinitiate at the distal end of the filaments,which extend to the base. Epithelialvacuolation, necrosis, congestion, oedema,fusion and degeneration of the secondarylamellae subsequently follow. Acute mortal-ity is usually associated with gill lesions.Internal pathology or host inflammatoryresponse may occur, and the pathogenmay be isolated from internal tissues(Thune et al., 1993; Plumb, 1994; Shotts andStarliper, 1999).

Farkas and Olah (1986) described thethree stages of gill necrosis. The first stage isinitiated and maintained by environmentalstress (probably ammonia, pH, temperatureor any toxins in the rearing water) butF. columnare is seldom detectable on gillsthat are pale or dark purple. The secondstage consists of bacterial invasion of thedamaged gill at water temperatures above20°C, causing gill necrosis, resulting ina grey-white coating of the gills. In thethird stage, the white coating of the gillsdisappears and the infected gills becomedistorted. Different stages of gill necrosismay be observed in the same fishpopulation.

Transmission of the bacterium isvia water. The disease is most commonlyassociated with stress from high tempera-tures, elevated organic loads, high stockingdensity, low dissolved oxygen and traumafrom excessive handling. In channel catfish,it occurs more often at temperatures between25 and 32°C with significant mortality.




Young fish are more susceptible than olderfish. It may occur as a primary infection or asa mixed infection with another bacterium,E. ictaluri or A. hydrophila, or in associationwith a parasite, e.g. Henneguya sp. or Ichthy-obodo sp. (Hawke and Thune, 1992; Duarteet al., 1993; Plumb, 1994). Columnaris dis-ease appears to follow outbreaks of otherdiseases (Duarte et al., 1993).

Survivors of columnaris disease releasethe pathogen into the water at rates of upto 5 × 103 cells ml−1 h−1 (Fujihara andNakatani, 1971), and surviving fish mayrelease the bacterium for up to 140 dayspost-infection. The severity of lesiondepends on the virulence of the strainand the ability of the pathogen to produceproteolytic enzymes. F. columnare producesan extracellular chondroitin AC lyase thatdegrades chondroitin and hyaluronic acidin fish connective tissue (Griffin, 1991).Bertolini and Rohovec (1992) also reportedfour extracellular proteases with molecularweights of 32, 34, 40 and 47 kDa. Newtonet al. (1997) further observed that moreprotease is secreted into a medium withlow nutrients and salt (Ordal’s medium)than into media with high concentrationsof nutrients or salt (TYES, Hsu–Shotts,modified Shieh’s media).

Gills or dermal/muscular capillariesof infected fish become congested anddegenerate (Plumb, 1994). Kuo et al. (1981)showed that survival of fish given0.35–1.4 mg iron 100 g−1 fish prior tochallenge with the pathogen was reducedfrom 3 days to 1 day. Furthermore, highlyvirulent strains of F. columnare adheredmore readily to the gills than low virulencestrains, and were enhanced in ion-richwater, in the presence of nitrite or organicmatter and at 28°C temperature (Decostereet al., 1999).

The bacterium can survive up to 16 daysat 25°C in hard, alkaline water with a highorganic load, but survival decreases at pH 7or less and in waters with less than 50 mgCaCO3 l−1 and with low organic matter(Fijan, 1968). In sterile mud at 25°C, theorganism survives for 16 days (Becker andFujihara, 1978).

Diagnosis. F. columnare is a slender,Gram-negative, non-flagellated rod (about0.5 × 4–12 µm) with gliding motility andforms ‘hay stacks’ or columns. Primaryisolation of the pathogen can be achievedon selective Cytophaga agar supplementedwith 5 µg neomycin ml−1 and 200 IUpolymyxin B ml−1 (Hawke and Thune,1992). F. columnare colonies are yellow toorange and rhizoid. This aerobic organismcannot tolerate more than 0.5% NaCl andit grows between 4 and 36°C, producinggelatinase, caseinase, catalase, oxidaseand chondroitin sulphatase (Song et al.,1988).

Diagnosis of the disease is dependent onthe appearance of typical lesions on the skin,fins and gills, including the detection of thefilamentous bacterial cells in wet mountsmade from lesions. Based on the genesequence of the 16S rRNA of the bacteria,Bader and Shotts (1998) designed primersfor its detection using PCR.

Prevention and control. Disease preventionis by maintenance of fish under optimalenvironment conditions, proper handlingof fish, prophylactic treatment and goodhealth management practices (Plumb,1994). Daily oral vaccination with heat-killed F. columnare for 4 weeks reportedlyreduced mortality of rainbow trout from48 to 8%, with protection correlated withantibody levels (Fujihara and Nakatani,1971). Moore et al. (1990), however, showedthat immunization of channel catfishusing formalin-inactivated F. columnarebacterin by immersion yielded inconsistentresults.

Potassium permanganate at 5 mg l−1

(depending on the organic load of the rearingwater) in combination with oxytetracyclineadded to feed at 50 mg kg−1 fish day−1 for10 days is effective in controlling outbreaksin cages. Potassium permanganate (based onthe cage volume) mixed with a few litres ofwater and then poured through a 7.5 cmdiameter polyvinyl chloride (PVC) pipe intothe cage and allowed to dissipate into thepond by diffusion is also effective (Duarteet al., 1993).




Streptococcalsepticaemia/meningoencephalitis

In freshwater cage-cultured Mozambiquetilapia (Oreochromis mossambicus), epizo-otics attributed to streptococcal septicaemiawere reported in Taiwan (Tung et al.,1985). Other outbreaks have included thedisease in Nile tilapia, hybrid tilapia(O. niloticus × O. aureus), rainbow trout(Oncorhynchus mykiss), striped bass(Morone saxatilis) and hybrid striped bass(Morone chrysops × M. saxatilis) in Israel,Japan and the USA (Kitao et al., 1981; Kitao,1993; Eldar et al., 1994; Perera et al., 1994;Baya et al., 1996; Stoffregen et al., 1996).Pathogenic species are Streptococcus iniae(phenotypically identical to Streptococcusshiloi) (Eldar et al., 1995b), Streptococcusdifficile and other Streptococcus spp.

Most reports on streptococcal infectionshave occurred among wild and culturedmarine chinook salmon (Oncorhynchustshawytscha), rabbitfish (Siganus canali-culatus) and barramundi (Lates calcarifer)in the USA, Singapore and Japan (Moring,1982; Foo et al., 1985; Bromage et al., 1999).S. iniae specifically causes infections inmarine finfishes, as discussed in Chapter 5.

Pathology. Among tilapia (15–20 cm inlength) cage-cultured in a dam, thisbacterium caused cumulative mortalityof 50–60% within 1 month (Tung et al.,1985). Clinical signs include unilateraland bilateral exophthalmia with or withoutconjunctival haemorrhage and corneal opac-ity. Petechiae occur on the underside ofthe operculum, around the anus, caudaland pectoral fins and mouth, with darkeningof the body and discoloration of the dorsaland lateral trunk and peduncle with nodularor abscess formation. Abdominal swellingwith ascites is common. Affected fish areanorexic, swim sluggishly in a circle, turn-ing laterally, and eventually die.

Internal signs include petechiae andhaemorrhage of the intestinal tract, liverand pyloric caeca. Systemic infection hasbeen observed with evidence of bacterialdissemination in the heart, liver, kidney,

stomach, small intestine, brain, eyesand musculature. Multiple necroses withgranuloma occur in the hepatic parenchyma.The spleen develops hyperplasia of thereticuloendothelial cells with necrotic foci.Degenerative changes in the renal tubules,catarrhal enteritis in the small intestine andstomach, bacterial meningitis and abscessformation in the muscles have been noted.

The disease was experimentally repro-duced in trout and tilapia using 107 and 108

cfu of S. shiloi and S. difficile, respectively,with virulence increased to 102 and 105 cfuafter in vivo passage (Eldar et al., 1995a).Streptococcus is also more pathogenic toNile tilapia than to channel catfish (Changand Plumb, 1996). In a mixed infectionexperiment with Streptococcus sp. and A.hydrophila as inocula, mortality was higheramong experimental fish inoculated withboth bacterial pathogens compared withthose inoculated with either Streptococcusor A. hydrophila (Liu et al., 1990). Infectionvia the nares is a potential route in Niletilapia and hybrid striped bass (Evanset al., 2000). Experimental transmissionoccurs by immersion, injection, orally or bycohabitation and is enhanced by injury tothe skin or stressful environment. Sources ofinfection are water, mud, contaminated feedor carrier fish (Plumb, 1994).

Environmental factors influenced thedevelopment of streptococcal disease inNile tilapia. Shoemaker et al. (2000) showedthat significantly higher mortality (about28.4%) developed in medium (11.2 g l−1),compared with 4.8% in low (5.6 g l−1) fishdensity treatments exposed to 2.5 × 107

cfu ml−1 S. iniae by immersion. Moreover,the infection could be transmitted bycohabitation with S. iniae-infected Niletilapia for 48 h. In another study, Bunchand Bejerano (1997) demonstrated that lowoxygen and high nitrite levels increasedmortality in hybrid tilapia exposed toStreptococcus sp. However, these factorshad no additive effect. Furthermore,streptococcal infection in Nile tilapiafingerlings may occur in association withTrichodina infestation (J.A. Plumb, personalcommunication).




Diagnosis. Streptococcal organisms can beisolated in culture from the brain, kidney,heart, spleen and exophthalmia in Todd–Hewitt (TH) broth (DIFCO), nutrient agarsupplemented with sheep or goat’s blood,brain heart infusion agar or TSA for 24–48 hat 20–30°C (Kitao et al., 1981). ModifiedHucker’s Gram-staining showing small, Gram-positive cocci, approximately 0.3–0.5 µm indiameter, most often occurring in chains, is apresumptive diagnosis. These organisms arenon-motile and encapsulated. Plumb (1994)divided Streptococci associated with fishepizootics into four major groups: (i) groupB, which is non-haemolytic; (ii) group Dalpha- and group D beta-haemolytic; (iii)alpha-haemolytic strains that do not reactwith Lancefield antisera; and (iv) otherStreptococci from freshwater and marinefish. The pathogen does not grow in 40%bile, 6.5% saline, 0.1% methylene blue milkor at 10 or 45°C (Kusuda and Salati, 1999).Details on the classification of Streptococcispp. based on biochemical and serologicaltests are in Kitao (1993) and Plumb (1994).All isolates from freshwater fish are beta-haemolytic (Kitao et al., 1981; Tung et al.,1985).

Prevention and control. Avoidance of stressdue to adverse or poor water quality,rough handling, high stocking density,non-removal of infected or dead fishand overfeeding should be followed.Formalin-killed S. difficile vaccine injectedintraperitoneally protects tilapia (Eldaret al., 1995c). Recently, Klesius et al. (2000)showed that intramuscular injection of acombined vaccine prepared from two strainsof S. iniae obtained from Nile tilapiaprovided relative percentage survivals of63.1 and 87.3% when challenged with itshomologous pathogens. Medicated feedwith enteroflaxin at 5 mg kg−1 body weightfor 10 days (Stoffregen et al., 1996) orwith erythromycin–doxycycline mixture at100 mg and 70 mg kg−1 body weight for6 days are also effective (Tung et al., 1985).Formalin treatment was used for theassociated Trichodina (J.A. Plumb, personalcommunication).

Pseudofungal Diseases

Stramenopiles are pseudofungal organismspreviously classified as mycotic microbes(Alexopoulos et al., 1996). Infectionsinduced by the stramenopiles (FamilySaprolegniaceae, Class Oomycetes) arecommonly called ‘water mould infections’,cotton tuft disease or saprolegniasis. Bran-chiomycosis and mycotic granulomatosisalso occur in cultured fish in fresh waters.The EUS is associated with a rhabdovirus,the bacterium Aeromonas hydrophilaand/or the stramenopile, Aphanomycesinvadans (see section on Diseases ofComplex Infectious Aetiology, p. 246).

Saprolegniasis

The Oomycetes are distributed worldwideand affect warmwater fish in ponds, lakes,dams and rivers. In India, Achlya spp.,Aphanomyces, Dictyuchus, Saprolegniaand Pythium were isolated from rohu, grasscarp, common carp, catla, banded gourami(Colisa fasciatus), Labeo bata, climbingperch (Anabas testudineus) and giantsnakehead (Channa micropeltes); Aphano-myces spp. from rohu and Puntius ticto;and Saprolegnia spp. from dwarf gourami(Colisa lalia), banded gourami, Nandusnandus, Heteropneustis fossilis andNotopterus notopterus (Srivastava, 1980;Bisht et al., 1996). Saprolegniasis was alsoreported in Nile tilapia, mango tilapia andcommon carp in Taiwan, Egypt, Nigeriaand Hungary (Chien, 1981; Okaeme et al.,1989; ElSharouny and Badran, 1995; Jeneyand Jeney, 1995).

Pathology. Aphanomyces piscicida causesmycotic granulomatosis in ayu (Plecoglos-sus altivelis) and dwarf gourami. Externalclinical signs include red spots on the bodysurface due to fungal growth, swelling, ero-sion and ulcers. Histologically, fungal-likehyphae and granulomas are seen in the inter-nal organs and musculature. It is also highlypathogenic to goldfish (Carassius auratus),




Rhodeus ocellatus, bluegill (Lepomismacrochirus) and crucian carp (Hataiand Egusa, 1977; Hatai et al., 1994).In experimentally infected ayu, typicalmycotic granulomatosis occurred, while incommon carp no inflammatory responsewas observed (Wada et al., 1996).

Fungal-like Aphanomyces spp.,Achlya, Allomyces and Saprolegnia arealso associated with EUS in snakeheads(Roberts et al., 1993; Paclibare et al.,1994; Willoughby et al., 1995). However,only Aphanomyces has been experimen-tally shown to induce lesions in naivesnakeheads (Chinabut et al., 1995; Lilleyand Roberts, 1997). Bruno and Wood(1999) provided a recent review onsaprolegniasis, which is also discussed indetail in Chapter 4.

Branchiomycosis

Another fungal-like pathogen, Branchio-myces, has also been implicated as a causeof loss of 85% of juvenile red tilapia hybrid(O. niloticus × O. mossambicus) and greentilapia hybrid (O. niloticus × O. aureus) inIsrael (Paperna and Smirnova, 1997). Carpare also susceptible (Post, 1983).

Pathology. Affected fish are lethargic withragged or corroded gills, which are eitherbright red or white to brown depending onthe degree of necrosis. Histological examina-tion of the gill filaments of infected fishdemonstrates the proliferation of hyphaeof up to 11 µm in diameter. At the onsetof sporulation, the hyphae contain multi-nucleated plasmodia, which develop intodaughter plasmodia. The final stage of celldivision yields a sporont filled with spores.Spores are released from the necrotic gillsand remain suspended in the water or fall tothe bottom.

In severe infection, some filamentsundergo complete degeneration with necro-tic residues of the pseudofungus. As a result,the pseudofungi reduce the blood supply tothe gills, causing necrosis and sloughingaway of the gill tissue. Hence, the disease

is commonly named ‘gill rot’ (Post, 1983).Secondary bacterial invasion of the filamentedges follows.

The presence of organic matter, algalblooms, dissolved fertilizer, low dissolvedoxygen, pH between 5.8 and 6.5, highstocking density and temperatures between25 and 32°C are predisposing factors. Underfavourable conditions, the disease maydevelop in 2–4 days although in vitro cultureof the pathogen produced spores on day14 of culture (Post, 1983).

Diagnosis. Two species have beendescribed: Branchiomyces sanguinis andB. demigrans. Squash preparations of thegills examined using light microscopy canbe used to differentiate the two species.B. sanguinis has a thin hyphal wall (0.2 µm),spores of 5–9 µm diameter and affects thegill filaments and gill lamellar capillaries.B. demigrans has a thicker hyphal wall(0.5–0.7 µm), spores of 12–17 µm diameterand infects the parenchyma of the gills (Post,1983).

Prevention and control. Affected fish shouldbe burned and/or buried. Survivors of theepizootic are carriers of the pathogen andshould not be cultured with naive fishor transported into Branchiomyces-freegeographical areas.

Diseases of Complex InfectiousAetiology

Epizootic ulcerative syndrome (EUS)

EUS affects wild and cultured snakeheads,catfish (Clarias spp.), Mastacembelusarmatus, Puntius spp., giant snakehead,Oxyeleotris marmoratus, Glossogobiusgiurus, blue gourami, snakeskin gourami(Trichogaster pectoralis), Trichopsis vittata,Siamese fighting fish (Betta splendens),swamp eels (Monopterus albus) and severalwild fish species (Lilley et al., 1998). Majoroutbreaks occurred in Malaysia in 1979,in Indonesia in late 1980, in Thailand in1981, in Kampuchea, Myanmar and Lao




PDR in 1984, in the Philippines in 1985, inSri Lanka in 1987, in Bangladesh and Indiain 1988, and in Bhutan and Nepal in 1989(Tonguthai, 1985; Lilley et al., 1998; Lio-Po,1998). In addition, EUS was observed inVietnam, Singapore and Pakistan. EUS-likelesions on fish were also reported in 1972 inAustralia (Rodgers and Burke, 1977) whereinfected fish included mullet (Liza spp.,Mugil sp.), sand whiting (Sillago ciliata),Acanthopagrus australis and Arrhamphussclerolepis. The disease was then called redspot disease (RSD). Similarly, in Papua NewGuinea, Toxotes chatareus, Kurtus gulliveri,Bunaka spp., goby, freshwater anchovyand spotted scat (Scatophagus argus) wereseverely affected in 1975 (Haines, 1983).Fish with EUS were found in all types offreshwater systems, including lakes, rivers,streams, culture ponds, rice paddies, irriga-tion canals and reservoirs. Cage-culturedsnakeheads in the Philippines are verysusceptible to the disease (Lio-Po et al.,1992). Similarly, it was reported amongcage-cultured P. gonionotus and L. hoevenii(Christensen, 1989).

Pathology. Lesions associated with EUS arecharacterized by severe, ulcerative, dermalnecrosis with extensive erosion/sloughingof the underlying musculature (Fig. 7.3). Thenecrotic muscular tissue emits a foul odour.Fish have frank ulcers that consist of erodeddermal layer, exposing the underlyingmusculature, which may be haemorrhagic.

In less severe infections, there is scale losswith erosion of the skin surface with orwithout haemorrhagic signs. To date, EUS isdefined as a seasonal epizootic condition offreshwater and estuarine warmwater fish ofcomplex infectious aetiology characterizedby the presence of invasive Aphanomycesand necrotizing ulcerative lesions typicallyleading to a granulomatous response(Roberts et al., 1994a).

In general, EUS outbreaks show aseasonal pattern (Phillips and Keddie,1990). In Laguna de Bay, the Philippines, theEUS morbidity rate among snakeheads wasestimated to be 59% in January, 1986 (Minesand Baluyot, 1986). Outbreaks are morecommon from September to March, whichcorrelates with the period when the watertemperature in the region is at its lowestrange of below 25°C. Such low temperaturesreduce the immune response of fish (Catapand Munday, 1998).

The spreading pattern of outbreaks ofEUS in Southeast and East Asia stronglyindicates the infectious nature of theaetiological agent. The actual pathogen ofthis disease has been in dispute for years.A rhabdovirus, Aeromonas hydrophilaand Aphanomyces invadans have beenassociated with EUS-affected fish(Frerichs et al., 1986; Llobrera and Gacutan,1987; Boonyaratpalin, 1989; Costa andWejeyaratne, 1989; Lio-Po et al., 1992, 2000;Pathiratne et al., 1994; Chinabut et al., 1995;Karunasagar et al., 1995; Thanpuran et al.,


Fig. 7.3. Snakehead (Channa striata) affected with epizootic ulcerative syndrome.



1995; Kanchanakhan, 1996; Lilley andRoberts, 1997; Lilley et al., 1998). Saitanuet al. (1986) also detected a virus associatedwith EUS.

The association of a rhabdovirus withEUS in Thailand and in the Philippineswas first reported by Frerichs et al. (1986)and by Lio-Po et al. (2000). The virus is bul-let-shaped, typical of the rhabdovirus genus(Family: Rhabdoviridae) and induces a CPEin BF-2, SSN-1, CFS, CCO and SHS cells,producing virus titres in the latter cells of 106

TCID50 ml−1 at 25°C in 2–3 days (Lilley andFrerichs, 1994; Lio-Po et al., 2000). Opti-mum replication in SHS cells is at 15–25°C.Characterization and serological comparisonof the virus with other fish rhabdovirusesassociated with EUS-affected fish in Thai-land showed that the Philippine virus iso-late is morphologically similar and slightlyantigenically related to the ulcerativedermal rhabdovirus (UDRV) (Lio-Po et al.,2000). Earlier experiments on the pathoge-nicity of rhabdovirus from EUS fish were notdemonstrated (Frerichs et al., 1993). How-ever, subsequent studies experimentallyinduced lesion development and mortalityin virus-injected snakeheads reared at20–22.5°C but not at 28–32°C (Lio-Po et al.,2001). Similarly, Kanchanakhan (1996)reported that rhabdoviruses can experimen-tally cause skin damage in juvenile snake-heads at ~20°C. This lower temperaturerange corresponds to the water temperatureduring the cooler months of Decemberthrough to February when outbreaks of EUSamong freshwater fish occur in the Philip-pines and in other EUS-affected countries.

A. hydrophila has been consistentlyisolated from lesions of EUS-affected fish(Llobrera and Gacutan, 1987; Boonyarat-palin, 1989; Costa and Wejeyaratne, 1989;Subasinghe et al., 1990; Torres, 1990; Lio-Poet al., 1992; Pathiratne et al., 1994; Angkaet al., 1995; Karunasagar et al., 1995;Thanpuran et al., 1995; Rahman et al., 1999).Pure cultures of the bacterium inoculatedintramuscularly induced dermonecroticlesions in healthy catfish and snakeheads(Lio-Po et al., 1992, 1996, 1998; Pathiratneet al., 1994; Angka et al., 1995; Karunasagaret al., 1995). This bacterium grows at a

temperature range of 18–39°C and secretesa dermonecrotic factor at temperatures of10 and 30°C (Olivier et al., 1981; Uddinet al., 1997). Moreover, cytotoxin-producingstrains were associated with EUS-affectedfish and hypothesized to play an importantrole in the pathogenesis of the disease(Yadav et al., 1992).

The pseudofungi Aphanomyces spp.,Achlya, Allomyces and Saprolegnia havealso been reported in EUS-affected snake-heads (Roberts et al., 1993; Paclibare et al.,1994; Willoughby et al., 1995). Isolates ofA. invadans were experimentally shown toinduce lesions in most test snakeheads orsand whiting (Roberts et al., 1993; Chinabutet al., 1995; Catap and Munday, 1998). Otherstudies have reported that the pseudofungigrow invasively through the fish musclecausing severe myonecrosis (Callinan et al.,1995; Chinabut et al., 1995; Lilley andRoberts, 1997). Granuloma developmentwas observed at 26°C or above, while fish atlower temperatures showed acute inflam-mation (Chinabut et al., 1995). In addition,Catap and Munday (1998) observed thatsand whiting injected with zoospores ofAphanomyces sp. at 26°C developed highlyinflamed, haemorrhagic external lesions,while similarly treated fish held at 17°C hadslightly inflamed injection sites. The tem-perature-related growth rate of this pathogenappears to correlate with the findings thatAphanomyces isolates from EUS-affectedfish generally thrive better at 26–30°C thanat lower temperatures (Lilley and Roberts,1997).

Diagnosis. The virus is typical of the bullet-shaped rhabdoviruses with an estimatedsize of 65 × 175 nm (Lio-Po et al., 2000).Filtrates derived from the visceral organsof EUS-affected fish can induce a CPE wheninoculated into susceptible cells. The bacte-ria, A. hydrophila, and the pseudofungus,Aphanomyces sp., can be isolated fromulcers and muscles of EUS-affected fish bymethods described in the section on motileAeromonas septicaemia (Chapter 4) and inLilley et al. (1998). Histopathology ofmuscular lesions of affected fish shows thedevelopment of a necrotic granulomatous




mycosis, which may eventually invadethe abdominal viscera (Lilley et al., 1998).Bacterial colonies are also histologicallydemonstrated in EUS-affected snakeheads(Lacierda, 1995).

Prevention and control. Quarantine andrestricted movement of EUS-susceptible fishfrom endemic areas to non-endemic sitesshould be practised. Prophylactic treatmentwith 5 ppm Coptrol (a chelated copper com-pound) was reported to prevent inductionof EUS lesions while a proprietary mixture,CIFAX, may be curative (Lilley et al., 1998).Moreover, recent studies showed that fishfed with the immunostimulant Salar-becsurvived better when challenged with A.invadans (Miles et al., 2001).

Parasitic Diseases

Although there is information on parasiticdiseases of fish in tropical aquaculture(Kabata, 1985; Lim, 1991d, 1992; Paperna,1991, 1996; Arthur, 1992; Arthur andLumalan-Mayo, 1997), there is little orno information dealing specifically withparasitic diseases in cage culture systems.This paucity of information on diseasepathogens and control measures and thelack of regulations concerning movementof diseased fish and mandatory reportingof diseases and mortalities in developingcountries, coupled with the diverse speciescultured, have made management of para-sitic diseases in warmwater cage culture adifficult task. The diseases and specificidentity of the parasites infecting warmfreshwater cultured fish (in particular cagecultured fish) are seldom known and atbest only the genera are recorded (Paperna,1991). Overall, there is also a lack of knowl-edge about the actual disease patterns, thepathology and prevailing factors predispos-ing fish to the disease (Christensen, 1989;Dharma et al., 1992; Nasution et al., 1992;Alawi and Rusliadi, 1993). The lack ofcomprehensive investigations into thediseases encountered in cage culturesystems has resulted in the abandonmentof some lucrative projects such as the

culture of Oxyeleotris marmorata in cagesin Thailand (ADB/NACA, 1991). Thus,subsequent details of the diseases encoun-tered in cage culture systems are discussedunder generic and other taxonomic group-ings, rather under the specific pathogens inquestion.

Generally, wild/feral fish have greaterparasite species diversity but lower popula-tion abundance and the converse is truefor cultured fish but further studies arerequired (L.H.S. Lim, personal observation;Lerssutthichawal, 1999). Personal observa-tions and discussions with tropical fisheryscientists and the current literature indicatethat not all parasites known from other formsof culture systems have a similar impact oncage-cultured fish.

Diseases caused by protistans

The protozoan or protistan parasites thatcause disease in fish belong to several phylaand these include the Ciliophora, Myxozoa,Microspora, Sarcomastigophora and Api-complexa (Dickerson and Dawe, 1995;Dykova, 1995; Lom, 1995; Lom and Dykova,1995; Molnar, 1995; Noga and Levy, 1995;Woo and Poynton, 1995). The commonlyreported pathogenic protistans in or on fishreared in cages in warm waters include themyxosporeans, trichodinids and the dino-flagellates (Christensen, 1989; T.T. Dung,personal communication; F. Shaharom,personal communication). Leptobarbushoevenii cultured in cages in Indonesia areinfected with myxosporeans (Christensen,1989). In Vietnam, fish in cage culture areplagued by Trichodina, Balantidium (inthe intestines of catfish) and Glossatella(T.T. Dung, personal communication).The oodinid dinoflagellate Piscinoodinumsp. infects grass carp, bighead carp andP. gonionotus in pond culture, as wellas catfish and tilapia in cage culture(Shaharom-Harrison et al., 1991; F.Shaharom, personal communication).Various other protistan parasites have alsobeen recorded but their prevalence is notknown. For example, Ichthyobodo (Costia)and Oodinium are known to affect hybrids




of Clarias in tropical warm freshwaters(Paperna, 1991), resulting in pale gills andexcessive mucus secretions, causing thefish to gasp for air. The lack of reports onprotistan diseases in warmwater cage cul-ture systems could be due to lack of exper-tise in diagnosing the disease and/or theabsence of reporting procedures, rather thanthe absence of the disease agents.

Movement of fish for culture hascontributed to the worldwide distribution ofmany of their parasites, especially parasiticprotistans. For example, Eimeria cheni andEimeria sinensis, originally found in farmedcarp in China, are now found in Europe(Molnar, 1976). Nile tilapia imported intoThailand from Egypt were also infectedwith Eimeria vanasi (Paperna, 1991), whilecichlid fish farmed in Israel (Landsberg andPaperna, 1985) were infected by E. vanasiand Gousia cichlidarum. A few protozoandiseases found in cold waters could beregarded as emerging disease problems incage culture in warm waters since thesecould be carried with their host species.

Myxosporean diseases

Myxosporeans are observed as cysts, infect-ing the skin and subcutaneous layer,muscle, gills, central nervous system aswell as visceral organs. These causeextensive lesions as cysts break, and mortal-ity occurs in cultured as well as feral fish(Lom and Dykova, 1995). In most cases inSoutheast Asia, the specific myxosporideanpathogens are not known and at bestthe identification is at generic level.Thelohanellus (Myxobolidae), Myxobolus(Myxobolidae) and Myxosoma (Myxidiidae)have been reported from exotic carpand indigenous cyprinids in the Indiancontinent, Southeast Asia and China(ADB/NACA, 1991; Paperna, 1991).Thelohanellus has been reported on P.gonionotus, common carp and Clarias spp.in Peninsular Malaysia (Paperna, 1991;ADB/NACA, 1991). Myxosporeans are amajor problem in Central Java (Indonesia),infecting L. hoevenii and P. gonionotusreared in ponds. However, in cages, theparasite was only found on L. hoevenii

(Christensen, 1989). Myxobolus koi hasbeen found on the gills of common carp andgoldfish in Japan (Egusa, 1992) and on fishfarms in Israel, Indonesia and the Indiancontinent, causing high mortality amongthe younger fish (ADB/NACA, 1991;Paperna, 1991), while Myxobolus artusis found on common carp in East andSoutheast Asian countries (Lom andDykova, 1995).

Pathology. M. koi infections on the gills ofcommon carp and goldfish result in manysmall white to large pinkish to red cysts inthe gill tissue (Paperna, 1991; Egusa, 1992).Large cysts are enclosed in the host connec-tive tissues, which turn dark red due tohaemorrhaging, leading to congestion anddegeneration of the gill capillaries. Themovement of the opercula and respiratoryprocesses are further affected by increasedmucus production and epithelial prolifera-tion. Spores of M. koi were also observedin the heart, liver, kidney and intestine(Hoshina, 1952). According to Lom andDykova (1995), Thelohanellus pyriformisforms large plasmodia in the subcutaneoustissue and muscle of cyprinids causing fatalepizootics in Indonesia. Little is knownabout the pathology caused by the othermyxosporeans.

Diagnosis. One characteristic sign of myxo-sporean infections is small white and/orlarge cysts on the gills. For example, M. koiare observed as small white cysts and largepinkish to reddish cysts in the gill tissuesof common carp and goldfish (Paperna,1991; Egusa, 1992). Opercular movements ofinfected fish are hampered and respiration isaffected by the increased mucus secretionand epithelial proliferation (Hoshina, 1952).Some myxosporeans are confined to thebody and these occur as white cystsunder the scales, often near the tail or fins,resulting in sores or ulcers on the skin(Christensen, 1989). Identification of themyxosporeans is based on the morpho-logical characteristics of the spores. Cystson the skin or gills are removed and gentlybroken to release the spores (preferably) on




glass slides. The multicellular spores areunique in possessing nematocyst-like polarcapsules (Lom and Dykova, 1995). They areusually oval–pear to round shaped, anteriorend pointed, posterior end rounded, 1–2polar capsules with polar filaments, sporo-plasm with or without iodophilic vacuoleand with or without posterior processes(Shulman and Shtein, 1962; Lom andDykova, 1995).

The spores of Myxobolus are oval topear-shaped with two polar capsules attheir pointed anterior; the posterior end isrounded and lacks processes. The spores ofHenneguya are round, oval or fusiform withtwo anterior polar capsules and valves withtwo caudal processes from the posterior end.The oval to round spores of Myxosoma aredifferent in having two polar capsules atone end and lack processes and iodophilicvacuoles, while Thelohanellus has oval toround spores with smooth valves withoutprocesses and one medially displaced polarcapsule.

Prevention and control. There is no effec-tive treatment and the best method is toremove and destroy heavily infected fishfrom cages (Christensen, 1989). In light(early) infections, the cysts should becarefully removed and destroyed. Treatmentwith saline (0.23–5.0%), copper sulphate(0.025–0.05%), potassium permanganate,formalin, methylene blue, glacial acetic acidor phenol is not effective (Hoshina, 1952) asmyxosporean spores are highly resistant tochemicals. The inclusion of certain drugs(such as Proguanil and furazolidone) in thefish feed has been shown to reduce sporeproduction and alleviate lesions (Lom andDykova, 1995). Although the life cycles ofsome species of myxosporeans are knownto involve intermediate hosts such asoligochaetes (Lom and Dykova, 1995), forthe majority of cases, the life cycles have notbeen elucidated and the actual intermediatehosts not identified. Hence, control of myxo-sporeans via eradication of intermediatehosts (oligochaetes) is not a viable option atthe present time. Eradication of heavilyinfected hosts appears to be the most viableoption for the moment.

Diseases caused by ciliates

The ciliates (Phylum Ciliophora) arecommon ectoparasites of fish, especially inhatcheries and on young fish in grow-outponds. Ichthyophthirius multifiliis is themost well known pathogenic ciliate and isrelated to the marine pathogen, Crypto-caryon irritans. Others include the tricho-dinids and Chilodonella. However, in themajority of reported cases in tropicalaquaculture, the specific identities of theseciliates are not known. Besides the knownobligatory parasitic (pathogenic) ciliates,there are also facultative parasites (Tetra-hymena, for example), which are opportu-nistic organisms.

Trichodinid diseases

Pathogenic trichodinids include Chilodo-nella, Trichodina, Tripartiella and Tricho-denella. A large number of trichodinids areassociated with the goldfish, common carp,grass carp, silver carp and bighead carp andthese were introduced into Israel andSoutheast Asia from China (Chen, 1955;Paperna, 1991). The trichodinids (Tricho-dina acuta, Trichodina centrostrigeata andTrichodina heterodentata) from Africancichlids have also been introduced intoSoutheast Asia (Albaladejo and Arthur,1989; Bondad-Reantaso and Arthur, 1989).

Trichodinids can be found on snake-heads and Pangasius conchophilus culturedin cages in Vietnam (T.T. Dung, personalcommunication). Although chilodonellosisoccurs mainly in cold waters, Chilodonellahexasticha has also been found on thebighead carp in Malaysia (Shariff, 1984).

Trichodinids commonly cause mortal-ity in hatcheries and these may continue tobe a problem after fish are transferred to cageculture systems. Trichodinids are prevalenton young clariid hybrids of African catfish(Clarias gariepinus) and Clarias sp. in cages.These are also found on silver carp, bigheadcarp and grass carp in hatcheries in Chinaand Vietnam, and are also on pangasiidsand Catla sp. in cage culture. In Nepal,trichodinids cause mortality among the fryduring spring and autumn. Although there




are many species of trichodinids, only a feware known to be pathogenic (Lom, 1995).

Pathology. Pathological effects are depend-ent on the host’s response, the intensityof infections and environmental conditions,since stressful conditions can compromisethe host’s ability to counteract infections(Paperna, 1996). Some trichodinids livespecifically on the body surface or on thegills, while others are found both on theskin and the gills (Paperna, 1996). In skininfections, the preferred sites are the basesof fins. These parasites damage epithelialtissue through adhesion and crawlingactions (Paperna, 1996). They feed onthe epithelial cells causing abrasion andsome trichodinids may suck out cellularcontents, damaging cells, which degenerateand disintegrate resulting in erosion anddesquamation of the epidermis (Paperna,1996). The host responds to the infectionby increased mucus secretion and epi-thelial hyperplasia, cellular destructionand inflammation. The damaged gills andepidermal tissues are targets for bacterialinvasion. The infected epidermis thickens,becomes turbid with mucus and sloughedepithelial cells, and the fish becomesemaciated. When the gills are infected,excessive mucus is produced, with massivedestruction of the gills, and proliferationof epithelial cells causing difficulty inrespiration. Trichodinids are usually foundin association with monogenean and otherprotozoan infections. Massive infectionscausing damage in the epidermis asdescribed above result in mortality due todisruption in the respiratory functions ofthe gills (Paperna, 1996). Young fish in over-crowded and confined stressful habitats areusually heavily infected with trichodinids,while older fish have fewer but more host-specific species (Paperna, 1996).

Diagnosis. Trichodinids are easily obser-ved microscopically from skin and gillscrapings (Paperna, 1996). Taxonomy of thetrichodinids is based on the structure ofthe buccal ciliature, the morphology of theadhesive disc and the number and size of its

components (Lom, 1995). Trichodinids areessentially flat discs, with somatic ciliatureconsisting of 3–4 ciliary wreaths aroundthe aboral surface of the body, which istransformed into an adhesive disc. The discis a proteinaceous skeleton, composed ofa ring of hollow conical denticles. Thedenticles consist of blades (centrifugal flatprojections) and horns (rod-like centripetalprojections), connected to each other byradial pins (Fig. 7.4).

There are five genera of fish tricho-dinids (Lom, 1995). In warm freshwater cageculture systems, only Trichodina spp. hasbeen identified. Trichodina is characterizedby denticles with massive central conicalparts, flat semi-circular blades, straightthorns and a diameter of 50–100 µm. Forgeneral identification, skin and gill smearscontaining trichodinids should be air-dried,fixed in Bouins for 20 min, washed in 70%ethanol, rehydrated and stained in a haema-toxylin stain, dehydrated and mounted. Forspecific identification of the trichodinids,the adhesive disc is studied using a silverimpregnation method (Welborn, 1967;Paperna, 1996). Air-dried smears should befixed in 2% silver nitrate for 7–9 min in thedark, washed in distilled water and exposedto sunlight or UV light for 5–10 min.

Prevention and control. In most cases, anoutbreak of trichodinid infections is theresult of adverse environmental conditions,which are common in intensive culturesystems. The best preventative measure isto ensure that good quality environmentalconditions are maintained. To eliminatetrichodinids from aquaculture systems,several chemicals have been recommended(Lom, 1995): saline solution (0.1–0.2% as adip for 1–2 days), formalin (150–250 ppm asa dip for 30–60 min), acriflavine (indefi-nitely in water at 10–20 ppm) and potassiumpermanganate (0.1% as a dip for 30–45 min).Formalin has been used effectively to con-trol trichodinids in warm waters. The effi-cacy of formalin in controlling trichodinidsdepends on water quality (pH, salinity andambient temperature) and species of fishtreated. Van As et al. (1984) showed that




25 ppm for 24 h was effective in cleaninginfected carp, while 45 ppm for 24 h wasneeded to clean tilapia.

Ich or white spot disease

Ichthyophthirius multifiliis is a pathogenicciliate infecting freshwater fish causing ich-thyophthiriosis (also known as ich or whitespot). This pathogen was first reported fromChina (Dickerson and Dawe, 1995), but isnow a cosmopolitan pathogen in temperateand tropical warmwater fish (ADB/NACA,1991). It is predicted to spread with theincrease in aquaculture activities and alsovia the aquarium trade (Paperna, 1996). Theoutbreaks of ich are dependent on watertemperature and, as temperature increases,the life cycle of this parasite is completed ina shorter time (Dickerson and Dawe, 1995),making them a potential danger to cageculture systems in tropical warmwaters.This parasite is maintained within the fish

as a low subclinical (enzootic) infection andas encysted tomonts. It persists in the envi-ronment, becoming epizootic clinical infec-tions when fish are stressed as a result ofpoor management practices (e.g. poor feed,overcrowding and poor sanitation). Thepathogen is not host-specific and recoveryfrom the disease confers resistance to rein-fection (Paperna, 1996).

Pathology. The feeding or trophont stage islocated within the epidermis (gills or skin)of the fish (feeding on the basal layer ofthe epidermis). The matured tomonts leavethe fish and damage the epidermis causingdetachment from its basal membrane; theysecrete a gelatinous cyst wall and divideasexually to form tomites, which differenti-ate into infective theronts and are releasedinto the water. The tomites develop intoinfective theronts, which penetrate theepidermis of the fish becoming establishedin the basal layer of the epithelium just


Fig. 7.4. Trichodina acuta from the skin of Ctenopharyngodon idellus (Klein’s silver impregnation)(courtesy of Dr Richard Arthur, Canada).



above the basal membrane, and feed onepithelial cells. The rate of developmentof these stages is dependent on water tem-perature (see above). Intense and prolongedinfections cause epithelial proliferation,haemorrhagic inflammation and subsequentdisintegration of the integument.

Diagnosis. Clinical signs include anorexiaand lethargy, and the disease is character-ized by white spots on the skin and gills(Dickerson and Dawe, 1995). Skin and gillscrapings examined under the microscopereveal the ciliate (1 mm in diameter) witha small cytostome. Ichthyophthirius fixedand stained with Giemsa or haematoxylinreveals a large crescent-shaped macro-nucleus and small micronucleus.

Prevention and control. This pathogen isparticularly difficult to control. An inte-grated approach incorporating appropriateculture practices (locating cages in areaswhere water movement is continuous andstocking of clean and healthy fish), immuni-zation and chemotherapy in cases of heavyinfestations are probably the most effectivemeans of disease control (Dickerson andDawe, 1995; Paperna, 1996).

The chemicals recommended for treat-ment include sodium chloride, malachitegreen, formalin and potassium permanga-nate (Dickerson and Dawe, 1995; Paperna,1996). The efficacy of these chemicals isdependent on a number of factors such asenvironmental conditions, the fish speciesin question and the different developmentalstages of the parasites (see below). Forexample, encysted tomonts in the environ-ment are resistant to antiparasitic chemicals(Paperna, 1996). The stages of the parasitethat can be destroyed are the dividingtomonts and the newly released tomites.Several chemicals have been listed foruse against this pathogen, and the cost-effective chemicals suitable for large-scale farming systems are malachitegreen (0.05–0.15 ppm used continuously for3–4 days) and a mixture of formalin andmalachite green (50 and 0.05 ppm) (Paperna,1996). The fish species has to be taken into

consideration when chemicals are usedsince some species, especially catfish, do notrespond well to malachite green (Paperna,1996). Potassium permanganate has beenused successfully in ponds to control ich butits effectiveness is affected by the amountof organic matter in the water (Dickersonand Dawe, 1995). Malachite green in a non-water-soluble formulation in feed has beenreported to be effective against trophonts(Schmahl et al., 1992). Immersion of fishinfected with ich in Toltrazuril or triazinone(10 µg ml−1) for 4 h (repeated daily for 3days) has been shown to be effectiveagainst trophonts (Dickerson and Dawe,1995). However, malachite green has beenreported to be carcinogenic and its useis limited to aquarium fish, and shouldnot be used in fish cultured for humanconsumption (Dickerson and Dawe, 1995).

Studies have shown that fish that haverecovered from ich infections developimmunity against the parasite (Dickersonand Dawe, 1995). Immunization and vac-cination offer another way to protect fishagainst ich. Experimental immunizationusing killed vaccines, intraperitoneal inoc-ulation with live theronts and controlledexposure to infective tomites have beenused (Paperna, 1996; Sin et al., 1996). Anexperimental recombinant vaccine (from a316 bp gene fragment of the immobilizingantigens, or i-antigens, of I. multifiliis andexpressed in Escherichia coli) has been dev-eloped for ichthyophthiriosis (Woo, 1998).Goldfish inoculated with the recombinantprotein vaccine in Freund’s adjuvant sur-vived a parasite challenge (He et al., 1997).

Diseases caused by dinoflagellates

There are five genera of parasitic oodiniddinoflagellates: Amyloodinium, Piscino-odinium, Crepidoodinium, Ochthyodiniumand Oodinioides on fish (Noga and Levy,1995). The ichthyotoxins produced by dino-flagellates cause massive mortality in cul-tured and feral fish (Steindinger and Baden,1984). The important freshwater pathogenicdinoflagellate in fish is Piscinoodinium,




which is closely related to the marinedinoflagellate pathogen, Amyloodinium.Piscinoodinium is not host-specific and hasbeen reported on feral, aquarium andcultured food fish species from diversefamilies in warm waters (Lom andSchubert, 1983; Paperna, 1991, 1996;Shaharom-Harrison et al., 1991).

Velvet or rust disease

Fish with excessive mucus covering thebody together with a rust-coloured appear-ance of the skin are infected with Piscino-odinium pillulare, the causative agent forvelvet rust diseases, gold dust disease,pillularis disease and freshwater Oodiniumdisease (Shaharom-Harrison et al., 1991).Piscinoodinium, like its marine relativeAmyloodinium, is found on a wide range ofhost species and is known to cause mortal-ity in warmwater fish (Paperna, 1996).

P. pillulare has been reported from 14tropical ornamental fish species as well ascultured carp and cyprinids (Shaharom-Harrison et al., 1991; Noga and Levy, 1995).In Peninsular Malaysia, P. pillulare occurson aquarium fish, cultured grass carp,bighead carp, P. gonionotus and L. hoevenii,causing mortality in the latter (Shaharom-Harrison et al., 1991). This pathogen alsocauses disease in cage-cultured Hemibagrusnemurus in the Trengganu River and inTilapia cultured in Kenyir dam, Malaysia(F. Shaharom, personal communication),although not to the same extent as that foundon pond cultured fish.

Pathology. Histopathological changes ofgill structure occur with a massive prolifera-tion of the gill epithelium, fusion of adjacentlamellae and separation of the gill res-piratory epithelium resulting in a severehyperplasia of the entire gill filament(Shaharom-Harrison et al., 1991). Thetrophonts of P. pillulare penetrate the hostcells by nail-like extensions resulting indegeneration and collapse of the cells,leading to focal erosion and proliferation ofthe epithelium and obliteration of the gilllamellae. The inner strata of the epitheliumbecome spongious and may undergo

complete lysis (Lom and Schubert, 1983;Paperna, 1991).

Diagnosis. Initial diagnosis can be based onclinical signs and confirmed by microscopicexamination of the trophont stage. Piscino-odinium infects skin and gills with clinicalsigns similar to amyloodiniosis. Infectedfish have a yellow to rust-coloured (velvety)skin, dense covering of mucus resulting indarkening of the skin, dyspnoea, anorexiaand skin ulcers (Shaharom-Harrison et al.,1991).

All oodinids have a parasitic trophontstage and a sessile, stalked, sac-like tropho-zoite stage, which feeds on the skin and gillepithelia. The trophont has a prominentstalk, which anchors the parasite to the host.It probably uses the stalk to absorb nutrients.After feeding, the trophont detaches, with-draws the stalk and forms an encystedtomont (reproductive cysts). The tomontdivides asexually forming dinospores, themobile infective stages. The trophonts andtomonts are important for definitive diagno-sis, and microscopic identification of thesestages is necessary. Trophonts are oval withsmooth walls, usually visible to the nakedeye as white spots (80–100 µm) and inLugol’s iodine turn dark blue.

Piscinoodinium is distinguished fromother oodinid dinoflagellates on the basis ofthe morphology of the trophont, especiallythe type of host attachment and modeof nutrition (Lom, 1981). Fish should beexamined live or immediately after death,and snips of the gills can be removed fromlive or recently dead fish and examined.Trophonts are removed by brushing the fishgently in a dish of water and the sedimentis examined under the microscope. Thetrophont of Piscinoodinium is a yellow-green, pyriform or sac-like cell, almostround, 12 × 29 µm, with a rudimentarysulcus and a short stalk with an attachmentdisc extending from its base and thin hold-fasts (rhizocysts) radiating from the stalk(Lom and Schubert, 1983). Head parts of therhizocysts are inverted in separate compart-ments (rhizothecas) in the sole of the disc,while their shafts are firmly embedded inthe host cell cytoplasm. The theca covers




the entire cell except for the area of theattachment disc.

Prevention and control. Outbreaks ofoodinid infections result from stress dueto poor environmental conditions. Hence,environmental manipulation is probably aviable approach to control outbreaks of Pis-cinoodinium. Formalin detaches trophonts,but does not inhibit division (Paperna,1996). A copper ion concentration of about0.15 ppm (mixture of 5-hydrate coppersulphate with citric acid monohydrate) inwater is effective in controlling Piscinoodin-ium (Paperna, 1996). A salt dip for 1–3 mindislodges the trophonts, while immersionfor 3–5 days in a combination of 7 g salt l−1

and 40 mg potassium permanganate l−1

is also effective. However, freshwater fishcannot tolerate high salt concentration andpotassium permanganate higher than 2 mgl−1 (van Dujin, 1973; Plumb, 1979).

Diseases caused by monogeneans

Monogeneans are among the most com-monly reported parasitic agents of fish(ADB/NACA, 1991). They are mainlyectoparasitic on the gills, buccal cavity,body surface and fins of freshwater fishalthough some are endoparasitic (Gussevand Fernando, 1973; Euzet and Combes,1998). Monogeneans are oviparous with theexception of the viviparous gyrodactylids.Although they rarely cause disease in wildfish, apart from the benedenids (Paperna,1975), they are important pathogens inintensive fish culture (Paperna et al., 1984).Their direct life cycle results in rapid andcontinuous recruitment, especially in warmwaters; this makes monogeneans especiallydangerous in intensive culture. Diseasecaused by monogeneans is normally moredebilitating than fatal, and subsequent mor-tality is usually attributed to viral or bacte-rial infection. Monogeneans stress the fishhosts by destroying the epidermal integrityof the fish, thus predisposing their hosts toother pathogens. Cone (1995) suggested thatmonogeneans could be the mechanical

vectors of bacterial and viral diseases, butfurther confirmation is needed. In intensiveculture systems, where intensity of infec-tion can be high on the gills, monogeneanscan cause death directly by inhibiting respi-ration through physical damage to the gills.Fish mortality from monogenean infectionsmay result from damage to gill tissues andskin caused by attachment organs, and byfeeding on the integument, which stimu-lates cell proliferation and secretion ofcopious amounts of mucus (Paperna, 1991).Cage culture in tropical areas is usuallyconducive to the perpetuation of parasiticdiseases with high stocking density. Thenets trap eggs, infective larvae and fooddebris around the cages, which attractcarrier/reservoir feral fish.

Most monogenean genera are specific toa group of related host species. Dactylogyrusis found on cyprinids and catfish harbourThaparocleidus. Although at species levelmost species are specific to a particular hostspecies, some species, like Thaparocleiduscaecus, are found on a number of pangasiids(Lerssutthichawal, 1999).

Many of the monogenean species onwarm freshwater cultured fish have not beenidentified or are incorrectly classified. Forexample, Dactylogyrus spp. have also beenincorrectly implicated as being pathogenicto snakeheads, tilapia and clariids culturedin Southeast Asia (Kabata, 1985). These fishpossess their own unique and specificmonogeneans (Lim and Furtado, 1983,1986; Lim, 1986, 1991a). Trianchoratusis found on snakeheads other thangiant snakehead, which is infected bySundanochus spp., while Cichlidogyrusspp. infect the tilapias and Quadriacanthusspp. and Bychowskyella spp. infect theSoutheast Asian clariids. The gyrodactylids,on the other hand, are ubiquitous, althoughat species level they might be host-specific.

The most commonly reported monogen-eans on warm freshwater cultured fish arethe Dactylogyrus spp. on carp, Cichlidogyrusspp. on cichlids, Bychowslyella spp. andQuadriacanthus spp. on clariids, Trian-choratus spp. on snakehead, Pseudo-dactylogyroides spp. on O. marmorata, andThaparocleidus spp. on catfish other than




clariids and Gyrodactylus spp. (see below).Pseudodactylogyrus spp. have beenrecorded from eels (Anguilla spp.) in warmwaters of Indonesia (K. Buchmann, personalcommunication), and it should be noted thatPseudodactylogyrus infections caused massmortalities of cultured eels, especiallyAnguilla japonicus, in Europe in the 1980s(Buchmann et al., 1987; Buchmann, 1997).

In the majority of cases, the specificidentity of the pathogenic monogeneans,signs and pathology of the infection, diseasemechanism and control and preventativemeasures have not been specificallyelucidated and documented. For instance,it is known that Thaparocleidus siamensisoccurs in greater intensity than Thaparo-cleidus caecus on cultured P. hypophthal-mus in Peninsular Malaysia and Thailand(Lim, 1990, 1996; Lerssutthichawal, 1999),but it is not known which of the two speciesis pathogenic. The translocation of mono-geneans, along with their hosts, has beenwell documented for the various Dacty-logyrus spp. on imported Chinese carp,Cichlidogyrus spp. on tilapia and recentlyfor Quadriacanthus clariadis on the C. garie-pinus imported into Thailand (Paperna,1991; Lerssutthichawal, 1999).

There is information on the signs,pathology and control measures for somespecies of Dactylogyrus, Gyrodactylus andCichlidogyrus, but not for other monogeneanpathogens. In most cases, the informationis derived from pond culture systems andnot from cage culture systems. It should bealso noted that habitat can affect parasiticinfections, as indicated by the infestationof tilapia by Neobenedenia spp. instead ofCichlidogyrus spp. when farmed in cages inestuarine waters (see Chapter 5).

Diseases caused by Dactylogyrus species

Dactylogyrus species are specific to theCyprinidae although they are also found onHemiramphidae (L.H.S. Lim, unpublisheddata) and one species on a catfish (Gussev,1976). This genus is frequently listed as adisease-causing agent since cyprinids arethe most cultured fish group. Cyprinidscultured in cages and pens include the

common carp, grass carp, bighead carp,silver carp and Catla spp., as well as otherSoutheast Asian carp such as P. gonionotusand L. hoevenii (see Chapter 1).

The four important species of Dactylo-gyrus that cause disease in cultured commoncarp in Israel are Dactylogyrus anchoratus,Dactylogyrus extensus, Dactylogyrus min-utus and Dactylogyrus vastator (Paperna,1991). These have different temperaturepreferences: for example, D. extensus flour-ishes at low water temperatures (optimumtemperatures of 16–17°C), while D. vastatorprefers warmer waters (20–24°C). Currently,D. minutus can be found on common carp inTaiwan (Paperna, 1991). The grass carp areinfested with Dactylogyrus lamellatus andDactylogyrus ctenopharyngodonis, silvercarp with Dactylogyrus hypophthalmich-thys, Dactylogyrus suchengtaii and Dacty-logyrus scriabini and bighead carp withDactylogyrus aristichthys and Dactylogyrusnobilis (Paperna, 1991). In PeninsularMalaysia, D. nobilis and D. aristichthys arefound on cultured bighead carp and D. lam-ellatus on grass carp (Shaharom, 1988). P.gonionotus and L. hoevenii are infested withDactylogyrus leptobarbus and Dactylogyruslampam (Mizelle and Price, 1964; Lim andFurtado, 1986; Lim, 1991b), respectively, inPeninsular Malaysia. In Thailand, however,there are seven species of Dactylogyruson feral P. gonionotus (Chinabut and Lim,1993). Dactylogyrus has also been shown tocause mass mortality of fry, small fish andbroodfish (Paperna, 1991).

Pathology. The pathology caused by Dacty-logyrus spp. on exotic carp has been reportedin studies done in Europe, but not for thespecies infecting the indigenous cyprinidsof Southeast Asia. Feeding on epithelialcells and anchorage (attachment) by themonogeneans cause severe destruction ofthe gills resulting in haemorrhage and meta-plasia of the gill tissue. Secondary bacterialinfections usually occur and result in deathof the fish. The pathologies caused byD. vastator and D. lamellatus are similar(Molnar, 1972; Paperna, 1991). D. vastatorinfestations cause severe hyperplasia ofthe epithelium of gill filaments. Extensive




proliferation of the respiratory epithelium ofthe gills interferes with respiratory functionsand may be a direct cause of death. The sitesof proliferation are dependent on the pre-ferred sites of the monogenean species. D.vastator prefers the tips of the gill filamentsand causes mass mortality in young fish butseldom on fish greater than 32–35 mm sincethe functions of the remaining gill filamentsare not affected. Massive infestations ofD. extensus can cause mortality in 4–7 kgbroodfish (Paperna, 1991).

Diagnosis. Fish infected with Dactylogyrusspp. are lethargic and usually found swim-ming on the surface of the water. Fishheavily infected with Dactylogyrus havepale to greyish gills, swollen at the edges,and the opercula appear to open widerthan normal and secrete excessive amount ofmucus (Christensen, 1989). Heavily infectedfish are also anorexic and are usually foundgasping for air and exhibiting abnormalbehaviour such as jumping out of the water.

Dactylogyrus spp. are usually found onthe gills, although in massive infections theycan also be found on the buccal cavity.Dactylogyrus spp. can kill directly bydamaging gill structures and affecting respi-ration. In warm eutrophic waters with lowoxygen, this becomes serious. Dactylogyrusinfections usually result in secondarybacterial infections with subsequent mortal-ity. At present, Dactylogyrus infections areconfirmed by examination of the gillsand infected fins for presence of themonogeneans.

The signs and pathology of monogeneaninfections are neither generic- nor species-specific. Hence, diagnosis of monogeneaninfection is based on the identificationof the pathogen itself. Correct diagnosis ofmonogeneans requires proper preparation ofthe parasite specimens. Gills can either becompletely removed or gill clippings can betaken from the infected fish. Each parasite isremoved carefully from the gills under a dis-secting microscope, placed on a slide andcovered with a coverslip. Excess water isremoved and the corners of the coverslipsealed with nail polish to prevent it frommoving (Lim, 1991c). Ammonium picrate is

added underneath the coverslip to clearand fix the specimens, which are examinedusing a phase contrast microscope. Mono-genean species are usually identified onthe basis of the sclerotized reproductiveand haptoral armaments on the cleared andflattened specimens. The Dactylogyrus areoviparous monogeneans with or withoutfour eye-spots, 14 marginal hooks, twoanchors, one to two connective bars and twoneedle-like structures and spindle-shapeddactylogyrid-type seminal vesicles. Thedescriptions for the various Dactylogyrus arefound in Gussev (1985) for imported carp, inLim and Furtado (1986) and Chinabut andLim (1993) for P. gonionotus and in Mizelleand Price (1964) for L. hoevenii. Presently,other diagnostic techniques (such as immu-nological) are not known.

Prevention and control. The main methodfor control of monogeneans is the applica-tion of chemicals. Chemotherapeutic treat-ments include dips or baths in salt, formalinor organophosphates (Dylox, Dipterex,Neguvon, Chlorophos), Bromex-50 andpotassium permanganate (Paperna, 1996;T.S. Thana, personal communication; T.T.Dung, personal communication). The rec-ommended doses and concentrations varyaccording to host and parasite species, aswell as physico-chemical properties of thewaters. A 1 h bath with formalin at 1:4000(< 10°C), 1:5000 (10–15°C) or 1:6000 (15°C)and a bath with 3–5 ppm potassium per-manganate for 1–2 h (Hoffman and Meyer,1974) has been recommended. Trichlorfon(Dylox) may be added in the food (50 mg kg−1

fish) four times at 3 day intervals each monthduring the critical periods. Lime and otherchemicals have been recommended for pondapplications: 0.4–0.5 ppm of trichlorphon(0.0-dimethyl-2,2,2 trichloro-1-hydroxy-ethyl phosphanate) has been used in Japanand 0.2 ppm dimethyl-1,2 dibromo-2,2dichloroethyl phosphate (Bromex) in Israel(Egusa, 1992). These chemicals will be effec-tive if the cages are in ponds. However, theywill not be effective for large bodies of waterand rivers where cages are usually located.The above chemotherapeutic formulationsare for specific regions, and effective doses




have to be tested for different waters. Ifchemicals prove ineffective, most farmerswill simply destroy their heavily infectedfish (personal communication with farm-ers). Eradication of feral reservoir fish fromponds is possible but not when the cages arein rivers or large lakes. The best alternativemanagement strategy includes good hus-bandry based on knowledge of the reproduc-tive biology and ecological requirements ofthe parasites such as temperature depend-ency. Using healthy fish fry from reliablehatcheries, limiting stocking density of fish,providing good quality feed and sanitationof nets will help to keep infestation at alow level. Some fish are able to acquireimmunity against monogenean infections(Paperna, 1964, 1991) and more studiesshould be done to see if this could be used inthe control of monogenean infections.

Diseases caused by Cichlidogyrus species

Cichlids are cultured in warm fresh-water cages as well as in warm estuarinewaters. Tilapias cultured in freshwaterare affected by Cichlidogyrus spp., whilein marine waters they are infected bythe marine monogenean, Neobenedeniamelleni (see Chapter 5). Tilapia (cichlids)is cultured in cages in freshwater inIndonesia, Vietnam and the Philippinesas well as Malaysia. Cichlids are hosts tospecies of Cichlidogyrus, Onchobdella andEnterogyrus (an endoparasitic monogeneanpresent on Sri Lankan cichlids). Sev-eral species of Cichlidogyrus and E.cichlidarium have been introduced withtheir fish hosts into the Philippines,Indonesia and Peninsular Malaysia (L.H.S.Lim, unpublished data; Shaharom, 1985).The Cichlidogyrus species on tilapia inIndonesia have been incorrectly identifiedas Dactylogyrus spp. (ADB/NACA, 1991).Cichlidogyrus spp. are also found on tilapiain cages in Vietnam (T.T. Dung, personalcommunication).

As noted by Paperna (1980) and Papernaet al. (1984), no reports of mortality dueto Cichlidogyrus spp. have been recorded,but Cichlidogyrus sclerosus was found

to cause severe gill damage in tilapiascultured in the Philippines (Kabata, 1985).Neobenedenia spp. found on tilapia in cagesin estuarine waters are more pathogenic thanCichlidogyrus spp. (see Chapter 5).

Diagnosis. The behaviour of the fish canindicate the presence of parasites and this issimilar to Dactylogyrus infection. However,accurate diagnosis requires removing thegills or gill clippings; the monogeneansare collected and prepared as stated abovefor Dactylogyrus. Cichlidogyrus can bedistinguished from other monogeneans byhaving a haptor with four anchors, with twobars, one of which is V-shaped and theother made up of three parts. To identifythe different Cichlidogyrus species, consultPaperna (1980).

Prevention and control. As for Dactylogyrusinfections.

Trianchoratus and Sundanonchus infections

Monogenean species belonging to these twogenera are found to infect the channids.Sundanonchus spp. are restricted to C.micropeltes while Trianchoratus spp. arefound on the other channids. Althoughthese monogenean species are found on andknown to plague cultured snakeheads, thereis no report of mortality due to thesemonogeneans.

Diagnosis. Methods for collecting andpreparation of monogenean species fordiagnosis are the same as for the Dactylo-gyrus spp. above. Trianchoratus spp. havefour anchors, of which one pair is vestigial,no connective bars, 14 marginal hooksand a dactylogyrid-type seminal vesicle(Lim, 1986), while Sundanonchus spp.,infecting giant snakeheads, can be differ-entiated from the other monogeneans inhaving four anchors, with two connectivebars (dorsal bar may be split into two), 16marginal hooks, a dactylogyrid-type seminalvesicle and an X-shaped vitelline duct(Lim and Furtado, 1985; Kritsky and Lim,1995).





Diseases caused by Pseudodactylogyroidesmarmoratae

Pseudodactylogyroides marmoratae hasbeen found on cage-cultured O. marmorata,a highly priced fish in Malaysia and Viet-nam (Leong and Wong, 1998; T.T. Dung,personal communication). O. marmorata iscultured in cages in Peninsular Malaysia,Indo-China and Thailand. However, thisfish is no longer cultured in Thailandbecause of disease problems (ADB/NACA,1991). Other than the fact that this parasitecauses disease, practically nothing isknown about the signs, its pathology or howto control this pathogen.

Diagnosis. The monogenean species are col-lected and prepared as given for the Dactylo-gyrus species above. Morphologically, Pseu-dodactylogyroides spp. (Fig. 7.5) possess fouranchors, of which one pair is usually under-developed and small and the larger pair has apatch-like inner root, two connective bars,14 marginal hooks and a dactylogyrid-typeseminal vesicle (Lim, 1995).


Diseases caused by Thaparocleidus species

Species belonging to this monogeneangenus are found on cultured pangasiids andbagrids in Southeast Asia (Lim, 1990;Lerssutthichawal, 1999; A. Pariselle, per-sonal communication). As with Pseudo-dactylogyroides, little is known aboutthe pathology caused by this group ofmonogeneans, or its level of pathogenicity.

Diagnosis. Monogenean species frompangasiids are collected and prepared asfor Dactylogyrus infection above. Thaparo-cleidus spp. (Fig. 7.6) have four anchors, twoconnective bars, one of which may be wholeor separated into two, 14 marginal hooks anda sac-like seminal vesicle (Lim, 1996).


Diseases caused by Gyrodactylus species

The gyrodactylids are easily differentiatedfrom the dactylogyrids since they are vivi-parous with developing embryos in theuterus. The young gyrodactylids do notneed to search for a host. It has beenpostulated that gyrodactylids are able todisengage and reattach to new hosts, espe-cially under intensive culture where fishare in close proximity to each other.

Some Gyrodactylus spp. have widehost specificity and cause fish mortality.Gyrodactylids are easily translocated via thelive fish trade, for example Gyrodactylusturnbulli is spread via the aquarium trade toEngland, New England States, Nova Scotiaand Peru from Singapore (Cone, 1995).Although the gyrodactylids are importantpathogens in warmwater culture systems,there is a paucity of information on thisgroup. Studies on the pathogenicity of theGyrodactylus spp. are mostly from temper-ate countries (Paperna, 1991; Cone, 1995).Gyrodactylus infection is common onClarias spp. such as C. batrachus, C. macro-cephalus, C. gariepinus and the hybrid ofC. macrocephalus and C. gariepinus rearedin cages in Thailand (Aqua Farm News,1993). Paperna (1991) reported Gyrodact-ylus rysavyi and Macrogyrodactylus on C.gariepinus in Africa. These parasites mayhave an impact on the future of cage cultureof C. gariepinus in the Ivory Coast as well asthe Clarias culture in Thailand, the Philip-pines and Indonesia. Gyrodactylus fuscushas been found on Clarias fuscus in NorthVietnam. Unidentified Gyrodactylus spp.infect cage-cultured L. hoevenii and P.gonionotus in Indonesia (Christensen,1989). Gyrodactylus is found on tilapiacultured in freshwater and brackish waters(Natividad et al., 1986).

Pathology. Gyrodactylus spp. are usuallyfound on the skin and fins, although thereare species that can be found on the gills.They are also found in conjunction withprotozoan and bacterial infections. Mucus





Fig. 7.5. Pseudodactylogyroides marmoratae from the gills of Oxyeleotris marmorata (reproduced withpermission from Systematic Parasitology).




Fig. 7.6. Thaparocleidus caecus from the gillsof Pangasius hypophthalmus (reproduced withpermission from The Raffles Bulletin of Zoology).

secretion is increased during heavy infec-tions, fins become frayed, skin ulcerated andgills damaged by the feeding and attachmentprocesses of the worm. Fish infected with

Gyrodactylus exhibit abnormal behaviour,i.e. rubbing against the net, anorexia, hyper-production of skin mucus, haemorrhagiculcers on the body sides, fin rot (mainly analand caudal fins) and thickening and opacityof the eye cornea. At this stage, it is easy todetect parasites on the eyes, skin and fins.The skin usually appears whitish. At the laterstage of infections, reddish inflamed areasdevelop on the skin and the eyes may becomeopaque and blind (Christensen, 1989).

Diagnosis. Initial diagnosis can be based onclinical signs with confirmation by examina-tion of the parasites. The monogeneans arecollected from skin and gill scrapings andprepared as for Dactylogyrus spp. (see above).The anterior region of the gyrodactylid isdivided into two lobes with two sets of headglands. Its haptor is armed with 16 hinged,marginal hooks, two anchors and two con-nective bars. Gyrodactylus spp. are difficultto identify (Paperna, 1991). The body size,excretory systems, dimensions and morpho-logy of the sclerotized parts (reproductivespines, anchors, marginal hooks, connectivebars) are important criteria for species differ-entiation (Malmberg, 1970). Gyrodactylusspp. in the tropical regions are poorly stud-ied and more investigations are required.

Prevention and control. A formalin bathusing 20–25 ml of 40% formalin in 100 lof well-aerated, clean water for 30 min isused. Other formulations include formalinat 1:2000 for 10 min and ammonia solutionat 1.5 ml ammonia 1−1. The latter twomethods reduce infections but do noteradicate them. Trichlorphon (0.25 ppm) isalso effective (Meyer, 1968).

Diseases caused by other helminths

Although there are pathogenic trematodes,nematodes, cestodes and acanthocephalansin tropical aquaculture (Paperna, 1996),the pathogenic species causing disease intropical cage culture systems are unknown.For instance, trematodes and cestodes havebeen found in cage-cultured Pangasius



bocourti and in snakeheads in Vietnam buttheir identities are unknown (T.T. Dung,personal communication). With the intro-duction of more exotic species into tropicalwaters there will probably be more reportsof helminthic parasites in aquaculture inthe future. The impact of helminthic infec-tions in aquaculture is not known: althoughthe intestines of several feral giant snake-heads infected with the acanthocephalanspecies Gorgorhynchus ophicephali wereobserved with perforations (personal obser-vation), its impact on cultured giantsnakeheads is not known. Since there areno reports of massive mortalities causedby cestodes, trematodes or nematodes, thefollowing sections will briefly deal withknown helminth infections in fish speciesthat are cultured in cages in warm waters.

Trematode infection

Sanguinicola (blood fluke) has beenrecorded on exotic cultured grass carp andbighead carp (Anderson and Shaharom-Harrison, 1986). Thus far, no Sanguinicolahas been reported on clariids of SoutheastAsia although Sanguinicola dentata isfound on Clarias lazera (now known as C.gariepinus) from Africa (Paperna, 1996) andthis species has been imported into Thai-land for culture purposes. Metacercariaecausing ‘black spots’ in cichlids and clariidsin Africa (Paperna, 1996) could spread toother tropical waters. Kabata (1985) notedthe presence of clinostomatids and hetero-phyids in farmed fish in the warm waters,but thus far none have been reported amongcage-cultured fish.

Nematode infection

Nematodes are common on feral as wellas food fish (L.H.S. Lim, unpublished data;Kabata, 1985). The nematode Anguillicolacrassa could become important since itshost, A. japonicus, is cultured in Taiwanand on a smaller scale in Indonesia. Theother nematode of importance is Philomet-roides cyprini in common carp (Paperna,1996). Camallanids are common on feralcatfish (L.H.S. Lim, unpublished data;

Kabata, 1985) but little is known about theireffect in cage culture.

Cestode infection

The adult Asian tapeworm, Bothriocephalusacheilognathii, causes mortality in heavilyinfected grass carp in Europe (Paperna,1996). The Asian tapeworm is not confinedto cultured imported carp but has spread tonative fish in warm waters of Asia (Peninsu-lar Malaysia) and Israel with the importedgrass carp, silver carp and bighead carp (Sha-harom, 1985; ADB/NACA, 1991; Paperna,1991, 1996). Paperna (1996) has provided adetailed account of the disease caused bythis cestode species. Cestodes are also pres-ent in cultured and wild fish in warmwaters: Lytocestus spp. are found in cul-tured and wild C. batrachus, while Sengaspp. are found in cultured and wild snake-heads, Channa spp. (Furtado, 1963; Furtadoand Lau, 1971; Furtado and Tan, 1973).

Cestode infections in fish and resultingmortality are sporadic. Fish infested withintestinal (adult) cestodes have retardedgrowth, erratic swimming behaviour anddistended abdomen, become emaciated,cease to feed, develop a haemorrhagic enteri-tis caused by the destruction of the intestinalepithelium, and heavily infected fish havevarying degrees of aseptic dropsy (Paperna,1996). The cyclopoid copepod is the inter-mediate host, and the cestodes could be animportant pathogen in cage culture systemssince fish are in intimate contact with theenvironment.

Diseases caused by parasitic arthropods

Lernaea and Ergasilus spp. (Copepoda),Argulus (Branchiura) and Alitropus(Isopoda) have been recorded on a widerange of cultured fish species (Kabata, 1985;ADB/NACA, 1991). Lernaea and Arguluscause the most problems in warmwateraquaculture in Southeast Asia and theIndian continent. They were introducedinto these countries via fish importation(ADB/NACA, 1991) and will be discussedin greater detail below. The isopod




Alitropus is another common arthropod inaquaculture systems (Fig. 7.7) and is associ-ated with poor fish growth and increasedfish mortality (L.H.S. Lim, unpublisheddata; Lester and Roubal, 1995). It could be apotential pathogen in warm freshwater cagecultures. However, nothing much is knownabout its impact in aquaculture. Infestationby the copepods Ergasilus, besides causingcage-cultured fish to lose weight and appearunsightly, causes gill damage and, in heavyinfestations, results in gill dysfunction(Kabata, 1985). Ergasilus was recorded tocause fish mortality in Indonesia, especiallyin young fish (ADB/NACA, 1991). Ergasilusis a common crustacean parasite of fishand a potential pathogen in cage culturesystems; however, little is known about itsecology or pathology.

Lernaea infections

Lernaea or anchor worm causes the mostdamage in warm freshwater fish and isusually associated with high mortality.Lernaea spp. seem to prefer warm watersof 26–30°C (Shields and Tidd, 1974).Although it is known that this parasitecauses disease in cage-cultured fish inSoutheast Asia, the extent of its impact anddamage to aquaculture has not been esti-mated (Kabata, 1985).

Pathology. Lernaea cyprinacea is distrib-uted widely with the global translocation ofcarp and is now recorded in 45 species ofcyprinids as well as in other orders of fish,especially the siluriforms (Lester and Roubal,1995). Lernaea is found in India, Nepal,Bangladesh, Thailand, Indonesia, Pen-insular Malaysia, Vietnam, China and Japan(ADB/NACA, 1991). Lernaeosis occurs inChina on silver carp, bighead carp, grasscarp and black carp, in India and Bangladeshon all the major carp, in Vietnam on bigheadcarp, grass carp, silver carp, common carp,crucian carp and snakehead, and in Indone-sia on common carp, P. gonionotus, spottedgourami, mudfish and catfish. In 1976, theseparasites reached epizootic levels, destroy-ing about 30% of fish in over 7500 ha ofponds, ricefields and open waters in West

Java and North Sumatra (ADB/NACA, 1991).In Southeast Asia, Lernaea polymorpha isfound on bighead carp and silver carp(Shariff and Sommerville, 1986).

Haemorrhaging and gross lesions occurat the site of Lernaea infections and are asso-ciated with bacterial and other secondaryinfections. There are relatively few studieson the effects of anchor worm infection onthe fish hosts in warm waters. Some authorssuggest that the attached females feed onhost blood, while others suggest that theyprobably ingest host cells and absorb tissuefluids (Egusa, 1992). Lester and Roubal (1995)provided detailed information on the othersigns associated with Lernaea infections, andthese included blindness, epidermal anddermal necrosis and haemorrhage and encap-sulation of the embedded horns of Lernaea.Copepodids of Lernaea may cause disrup-tion and necrosis of the gill epithelia, and


Fig. 7.7. Alitropus species (Isopoda) found on theskin of Channa micropeltes (picture by K.S. Liew).



large numbers of larvae on the gills maycause fish mortality. Lesions caused by pen-etration of metamorphosing females are gen-erally associated with punctuate haemor-rhage, and muscle necrosis is evident at thepoint of penetration of parasites (Khalifa andPost, 1976). Penetrating female L. polymor-pha cause punctuate haemorrhage in big-head carp resulting in mortality in heavilyinfected fish (Shariff and Sommerville, 1986).L. cyprinacea in the eyes cause blindness.

Diagnosis. There are over 40 species ofpathogenic Lernaea (Kabata, 1983), butin most outbreaks, the specific identity ofthe parasite is unknown. Lernaea spp. aremacroscopic and easily seen with the nakedeye on the surface of fish. Only femalesof Lernaea are parasitic and are highlymodified so they do not resemble free-livingcopepods. Adult Lernaea females havetheir anterior end embedded into the bodymusculature of their host, while their longrod-shaped body with two egg sacs pro-trudes outside the host tissue. The anteriorhead region is modified as a small hemi-spherical cephalothorax, which containsthe mouth, with a well-developed holdfast,bifurcate dorsal process and simple ventralprocess (anchor). The anterior region mayeven penetrate into the body cavity andembed into visceral organs. Lernaea spp. aredistinguished by the shape of the anterioranchors, which may be modified by bone orother structures encountered during devel-opment in their host tissue. Ergasilus, on theother hand, is recognizably a copepod with asecond antenna modified for attachment anda pair of multiseriate egg sacs arising fromthe genital segment.

Prevention and control. Several chemicalsare recommended but their efficacy requiresfurther careful testing (Kabata, 1985; Egusa,1992; Paperna, 1996). Kasahara (1962)effectively used Dipterex (organophosphatetrichlorphon) to control and eradicate thelarval stages of L. cyprinacea in the watercolumn. At temperatures of 20–27°C, con-centrations of 0.5 and 0.2 ppm killthe nauplii in 1 and 2 days, respectively.

Copepodid stages are killed in 24–36 hat 0.2 ppm and in 12–18 h at 0.5 ppm at20°C, but Dipterex is not effective on adultfemales. However, in cages located in riversor large lake systems, the use of chemicals isineffective and dipping fish in chemicalsseem to be insufficient to get rid of all thecopepodid stages (Lester and Roubal, 1995).

Argulus infections

The majority of the branchiurans are fresh-water parasites (about 75% of the 120species of Argulus), with few estuarine ormarine species (Kabata, 1985). Argulus orfish louse (Fig. 7.8) is macroscopic and eas-ily observable on the skin and fins and alsoin the oral cavity. Infected skin becomesopaque with frayed fins. This ectoparasiticcrustacean feeds on the mucus layer, fleshand blood of the fish. The prolonged feed-ing and strong attachment of Argulus byits suckers on to the host result in directmechanical damage to the skin and dis-ruption of epithelial structure, resulting inlesions and subsequent invasions by oppor-tunistic pathogens such as pseudofungi(Singhal et al., 1986; van der Salm et al.,2000). There are at least four species ofargulids (Branchiura, Argullidae) that areeconomically important as parasites of fishin warm freshwater aquaculture, i.e. Arg-ulus japonica, Argulus foliaceus, Argulusindicus and Argulus siamensis, and thesehave been introduced along with theircyprinid hosts and are now reportedfrom both local indigenous cyprinids and


Fig. 7.8. Argulus species (Branchiuran) foundon the skin of Channa micropeltes (picture byK.S. Liew).



non-cyprinid hosts in the introduced areas(Paperna, 1991). A. japonica is in Israel,while A. foliceaus is in Thailand, Penin-sular Malaysia and Sri Lanka on carp andnative cyprinid species (Kabata, 1985).A. indicus, an Asiatic species, is on anaban-tids, chaniids, tilapias and native cyprinidsin Indonesia, Thailand and India. A. sia-mensis is reported in Thailand from ana-bantids and Cirrhina spp. (Gopalakrishnan,1968; Kabata, 1985) and in India on asnakehead species (Channa gachua) (Rama-krishna, 1951). A. japonica is an importantparasite of warm freshwater fish, whileArgulus coregoni parasitizes cold fresh-water fish.

Pathology. This parasite is not host-specificand is found on a wide range of fish speciesfrom cyprinids to siluriforms and perciforms(see above). The life cycle of the parasiteis direct and the eggs hatch into free-swimming larvae, which must find a hostwithin 2–3 days. It is reported to cause mas-sive mortality of fish in Bangladesh, and inthe majority of cases the outbreaks were sea-sonal, usually in the colder months (Kabata,1985). Argulus usually infects the young fishfrom spring until early summer. The parasiteis also common in India, affecting the majorIndian carp, especially Rohu spp. In Penin-sular Malaysia, argulids have been foundon wild fish such as C. micropeltes (L.H.S.Lim, unpublished data), the imported fryof bighead carp and grass carp (Shaharom,1988). Argulus spp. are found on the sandgoby and snakeheads in cages in Vietnam(T.T. Dung, personal communication).Heavily infected fish are lethargic, listless,cease to feed and rub themselves on the sub-strate in an attempt to dislodge the parasite.

The lesion or wound made by the feed-ing Argulus may be restricted to the epider-mis or may penetrate through to the stratumspongiosum of the dermis and even thestratum compactum turning the dermisoedematous (Lester and Roubal, 1995). Thearea may become necrotic with secondarybacterial and fungal infections. Mortalitymay be associated with changes in the ionicand osmotic homeostasis, anorexia and

secondary infections. Kabata (1970),Paperna and Zwener (1976) and Paperna(1980) noted that lytic and toxic substanceswere secreted into the dermal area, whilefeeding caused acute haemorrhagic,inflamed wounds. Argulus feeding on bloodcauses fish to become anaemic and its pierc-ing proboscis stylet causes haemorrhagicspots on the epidermis. The spots are formedby epidermal hyperplasia. Bacterial infec-tions occur around the site of infection.Argulus may also be a vector of viral infec-tions. Ahne (1985) showed that springviraemia of carp (SVC) was transmitted by A.foliaceus, and in Israel, carp pox (carppapilloma) occurred in conjunction with A.japonica infestation (Sarig, 1971).

Diagnosis. The parasite is oval to round,dorso-ventrally flattened (about 4–8 mmin diameter), with a pair of modifiedsucker-like first maxillae. Its proboscis orfeeding organ is for inserting into the epider-mis and the underlying tissue of the fishhosts to feed on blood (Fig. 7.8).

Prevention and control. Several chemicals,especially organophosphate insecticides,formalin, chlorine, sodium chloride andeven antimalarial drugs, are recommended(Kabata, 1985; Egusa, 1992; Lester andRoubal, 1995; Paperna, 1996), but theirefficacy in different types of water bodiesis not known. Studies carried out in warmwaters of Israel and Africa show that some(see below) of the insecticides are effectivein killing argulids within the safety marginfor fish (Paperna, 1996). Lindane has beenused to clean fish of argulids prior to market-ing (Paperna, 1996). The chemicals in useare gemmexane (this is toxic to fish andman), Pyrethrum (not tested on a largescale yet) (Paperna, 1996), Dipterex, tricho-lorphon, Neguvon, malathion, formalinand antimalarial drugs such as quininehydrochloride (13 ppm) (Puffer and Beal,1981; Kabata, 1985; Singhal et al., 1986).However, not all the chemicals are equallyeffective for the different developmentalstages of argulids. For example, Dipterex(0.2–0.3 ppm) is effective against the adults




and larvae causing them to fall off the fishand die but has no effect on the eggs (Egusa,1992). The water chemistry and temperatureare important factors in the use of thesechemicals. The occurrence of these para-sites, despite the amount of chemicalsused, indicates that the eggs are still in thesystem and that the chemicals used are noteffective in destroying the eggs. The strategyof not stocking the ponds until the larvalstages have died could be effective. Othermethods of control include the use ofsubstrates such as wooden slats to trap eggs,filtering incoming water to remove larvalstages, stocking clean fish, quarantiningincoming fish with treatment if necessarybefore stocking, and stocking with argulid-predatory fish (Kabata, 1985).

Conclusions and Recommendationsfor Future Research

Microorganisms and parasites are normalflora and fauna inhabiting the skin, fins,gills and the gastrointestinal tract of fish.Under normal conditions, many of theseorganisms do not induce disease in theirfish host. However, man-made pollutantsand/or intensification of fish cultureresult in increased environmental changes,which may be stressful to fish. Bacterialmultiplication, for instance, is enhancedwith increasing organic matter fromuneaten feeds. The stress predisposes fishto invasion by opportunistic pathogens,with subsequent morbidity and mortality.Stress is also associated with handling,stocking, grading and shipping of fish.

Often fish mortality can be attributed toseveral factors (e.g. fish condition, patho-gens and environment) and it is difficult todetermine the significance of any one ofthese factors (Mitchell, 1997). Parasites suchas the monogeneans may not have a directeffect on fish mortality but they debilitatethe fish, making it more susceptible to otherpathogens. Their organs of attachment usu-ally create portals of entry for viral, bacterialand pseudofungal pathogens of fish. In addi-tion, some parasites are reservoirs of viralpathogens.

Despite the long history of aquaculturein the tropics and the importance of diseasein aquaculture, there have been few con-certed efforts to document and investigatethe diseases of fish cultured in cages andponds. This may be due to lack of trainedmanpower and institutional support. Thediversity of fish cultured in warm watersdoes not help to alleviate this problem.The usual approach to disease and healthmanagement is to use chemicals (usuallyindiscriminately) or, if this does not work, todiscard the fish species and start afresh withanother species. Also, there is no mandatoryrequirement in many of the developing andunderdeveloped countries to report fishdeath, and until recently fish were usuallyimported and stocked without quarantine.It must be taken into consideration thatthe movement of fish, especially acrossinternational boundaries, may transfer fishpathogens as well (Hedrick, 1996). In thisregard, the provisions of the Office Inter-national des Epizooties (OIE) (1995) Inter-national Aquatic Animal Health Codeshould be adhered to.

The lack of institutional support resultsin reduced research on pathogens and conse-quently an inability to control and preventdiseases. There is also a lack of trained per-sonnel in disease management and littlereliable information on the specific identityof pathogens. A related issue is the lack oflegislation and guidelines pertaining to theuse of drugs and chemicals in aquaculture.Currently, drugs and chemicals are usedindiscriminately (usually the aetiologicalagents are not identified), and without aspecific withdrawal period prior to the saleof the fish. Trained competent fish diseasemanagers, who are able to diagnose patho-gens and are capable of dispensing properprevention and control measures, are impor-tant to sustain aquaculture.

Acknowledgements

We would like to thank and acknowledgethe information provided by Dr RichardArthur (Canada), Dr F. Shaharom(Universiti Putra Malaysia, Trengganu




Branch, Malaysia), Dr S. Chinabut (AAHRI,Bangkok, Thailand), Dr T.S. Thana (Depart-ment of Fisheries, Phnom Penh, Cambodia),Dr T.T. Dung (Department of FreshwaterAquaculture, College of Agriculture, CanthoUniversity, Cantho, Vietnam) and Dr E.M.Leaño (SEAFDEC, Iloilo, Philippines). DrJ.A. Plumb (Department of Fisheries andAllied Aquacultures, College of Agricul-ture, Auburn University, Alabama, USA) isalso gratefully acknowledged for sharinginformation and for his critical review.

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