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ISSNO0428-9345Asia Diagnostic Guide toAquatic Animal DiseasesFAOFISHERIESTECHNICALPAPER402/2NETWORK OFAQUACULTURECENTRESIN ASIA-PACIFICN AC AFoodandAgricultureOrganizationoftheUnitedNationsF A OFI ATI SP A N

ISSNO0428-9345Asia Diagnostic Guide toAquatic Animal DiseasesFAOFISHERIESTECHNICALPAPER402/2Edited byMelba G. Bondad-ReantasoNACA, Bangkok, Thailand(E-mail: Melba.Reantaso@enaca.org)Sharon E. McGladderyDFO-Canada, Moncton, New Brunswick(E-mail: McGladderyS@dfo-mpo.gc.ca)Iain EastAFFA, Canberra, Australia(E-mail: Iain.East@affa.gov.au)andRohana P. SubasingheFAO, Rome(E-mail: Rohana.Subasinghe@fao.org)NETWORK OFAQUACULTURECENTRESIN ASIA-PACIFICN AC AFoodandAgricultureOrganizationoftheUnitedNationsF A OFI ATI SP A N

The designations employed and the presentation of material in thispublication do not imply the expression of any opinion whatsoeveron the part of the Food and Agriculture Organization of the UnitedNations (FAO) or of the Network of Aquaculture Centres in Asia-Pacific(NACA) concerning the legal status of any country, territory, cityor area or of its authorities, or concerning the delimitation of its frontiersor boundaries.ISBN 92-5-104620-4All rights reserved. No part of this publication may be reproduced,stored in a retrieval system, or transmitted in any form or by any means,electronic, mechanical, photocopying or otherwise, without the priorpermission of the copyright owner. Applications for such permission,with a statement of the purpose and extent of the reproduction,should be addressed to the Co-ordinator, Network of AquacultureCentres in Asia-Pacific (NACA), Suraswadi Building, Department ofFisheries, Kasetsart University Campus, Ladyao, Jatujak, Bangkok10900, Thailand, or the Chief, Publishing and Multimedia Service,Information Division, FAO, Viale delle Terme di Caracalla, 00100 Rome,Italy or by e-mail to copyright@fao.org .© FAO and NACA 2001iii 4

PREPARATION OF THIS DOCUMENTThe Asia Diagnostic Guide to Aquatic Animal Diseases or ‘Asia Diagnostic Guide’ is a comprehensive,up-datable diagnostic guide in support of the implementation of the Asia RegionalTechnical Guidelines on Health Management for the Responsible Movement of Live AquaticAnimals or ‘Technical Guidelines’. It was developed from technical contributions of membersof the Regional Working Group (RWG) and Technical Support Services (TSS) and other aquaticanimal health scientists in the Asia-Pacific region and outside who supported the Asia-PacificRegional Aquatic Animal Health Management Programme. The Asia Diagnostic Guide is a third ofa series of FAO Fisheries Technical Papers developed as part of an FAO Technical Co-operationProject – Assistance for the Responsible Movement of Live Aquatic Animals – implementedby NACA, in collaboration with OIE and several other national and regional agencies and organizations.The Technical Guidelines and the associated Beijing Consensus and ImplementationStrategy (BCIS) was published as first (FAO Fisheries Technical Paper 402) of the series. TheManual of Procedures for the Implementation of the Asia Regional Technical Guidelines onHealth Management for the Responsible Movement of Live Aquatic Animals or ‘Manual ofProcedures’, which provides background material and detailed technical procedures to assistcountries and territories in the Asia-Pacific region in implementing the Technical Guidelines wasthe second of the series (FAO Fisheries Technical Paper 402, Supplement 1). The Asia DiagnosticGuide (FAO Fisheries Technical Paper 402, Supplement 2) is published as the third document ofthe series. All of the above-mentioned documents, developed in a highly consultative processover a period of three years (1998-2001) of consensus building and awareness raising, are inconcordance with the OIE International Aquatic Animal Code (Third Edition) and the OIE DiagnosticManual for Aquatic Animal Diseases (Third Edition) and the WTO’s Sanitary andPhytosanitary Agreement (SPS) and in support of relevant provisions of FAO’s Code of Conductfor Responsible Fisheries (CCRF).DistributionAquatic animal health personnelFAO Fishery Regional and Sub-Regional OfficersFAO Fisheries DepartmentNACACover page: Representation of relationship between host, pathogen and the environment indisease development.iv 5

PREFACEThe Food and Agriculture Organization of theUnited Nations (FAO) and the Network ofAquaculture Centres in Asia-Pacific (NACA) arepleased to present this document entitled AsiaDiagnostic Guide to Aquatic Animal Diseasesor ‘Asia Diagnostic Guide’. The Asia DiagnosticGuide is the third and last of a series of FAOFisheries Technical Papers (FAO Fish. Tech.Pap. No. 402 and 402 Supplement 1), whichwas developed by representatives from 21Asian governments, scientists and experts onaquatic animal health, as well as by representativesfrom several national, regional and internationalagencies and organizations. TheAsia Diagnostic Guide provides valuable diagnosticguidance for implementing the Asia RegionalTechnical Guidelines on Health Managementfor the Responsible Movement of LiveAquatic Animals and their associated implementationplan, the Beijing Consensus andImplementation Strategy (BCIS) (see FAO Fish.Tech. Pap. No. 402). It also complements theManual of Procedures for implementing theTechnical Guidelines (see FAO Fish. Tech. Pap.No. 402, Supplement 1). The entire series ismeant for assisting national and regional effortsin reducing the risks of diseases due totrans-boundary movement (introduction andtransfer) of live aquatic animals. The implementationof the Technical Guidelines will contributeto securing and increasing income ofaquaculturists in Asia by minimizing the diseaserisks associated with trans-boundarymovement of aquatic animal pathogens. Inmany countries in Asia, aquaculture and capturefisheries provide a mainstay of rural foodsecurity and livelihoods, and effective implementationof the Technical Guidelines will contributeto regional efforts to improve rural livelihoods,within the broader framework of responsiblemanagement, environmentalsustainability and protection of aquaticbiodiversity.An FAO Technical Co-operation Programme(TCP) Project (TCP/RAS 6714 (A) and 9065 (A)- “Assistance for the Responsible Movementof Live Aquatic Animals”) was launched byNACA in 1998, with the participation of 21countries from throughout the region. This programcomplemented FAO’s efforts in assistingmember countries to implement the relevantprovisions in Article 9 - Aquaculture Development- of the Code of Conduct for ResponsibleFisheries (CCRF), at both the national andregional levels. A set of Guiding Principles, formulatedby a group of aquatic animal healthexperts at the Regional Workshop held in 1996in Bangkok, formed the basis for an extensiveconsultative process, between 1998-2000, involvinginput from government-designated NationalCo-ordinators (NCs), NACA, FAO, OIE,and regional and international specialists.Based on reports from these workshops, aswell as inter-sessional activities co-ordinatedby FAO and NACA, the final Technical Guidelineswere presented and discussed at the FinalProject Workshop on Asia Regional HealthManagement for the Responsible Trans-boundaryMovement of Live Aquatic Animals, held inBeijing, China, 27 th -30 th June 2000.The Technical Guidelines were reviewed anddiscussed by the participants of this meeting,which included the NCs, FAO, NACA, OIE (Representativesof the Fish Disease Commissionand Regional Representation in Tokyo), andmany regional and international aquatic animalhealth management specialists. The NCs gaveunanimous agreement and endorsement of theTechnical Guidelines, in principle, as providingvaluable guidance for national and regional effortsin reducing the risks of disease due to thetrans-boundary movement of live aquatic animals.Recognizing the crucial importance of implementationof the Technical Guidelines, the participantsprepared a detailed implementationstrategy, the Beijing Consensus and ImplementationStrategy (BCIS), focussing on NationalStrategies and with support through regionaland international co-operation. This comprehensiveimplementation strategy was unanimouslyadopted by the workshop participants.The countries that participated in the developmentof the Technical Guidelines and BCIS, andthe associated Manual of Procedures and AsiaDiagnostic Guide are Australia, Bangladesh,Cambodia, China P.R., Hong Kong China, India,Indonesia, Iran, Japan, Korea (D.P.R.), Korea(R.O.), Lao (P.D.R.), Malaysia, Myanmar,Nepal, Pakistan, the Philippines, Singapore, SriLanka, Thailand and Vietnam.vi 7

PREFACEFAO and NACA extend special thanks to all thegovernments, agencies, and organizations thattook part in this significant, and sometimesdaunting endeavor, as well as to all the individualswho generously contributed time, effortand expertise to the compilation of thisdocument and other information producedduring the process.Ichiro NomuraAssistant Director GeneralFisheries DepartmentFood and Agriculture Organization of the UnitedNationsViale delle Terme di Caracalla00100 Rome, ItalyFax: + 39 06 570-53020E-mail: ichiro.nomura@fao.org orfi-enquires@fao.orgWebsite: http://www.fao.org/fi/default.aspPedro BuenoCo-ordinatorNetwork of Aquaculture Centres in Asia-Pacific(NACA)Department of Fisheries,Kasetsart University Campus, Ladyao, JatujakBangkok 10900, ThailandFax: (662) 561-1727E-mail: Pedro.Bueno@enaca.orgWebsite: http://www.enaca.orgvii 8

FOREWORDMovement of live aquatic animals is a necessityfor development of aquaculture on bothsubsistence and commercial levels. However,such movements increase the probability of introducingnew pathogens, which can have direconsequences on aquaculture, capture fisheriesand related resources, as well as the livelihoodswhich depend on them. In order to minimizeor avoid the risk of pathogen transfer viaaquatic animal movements, it is essential thatthe individuals and organizations involved insuch activities appreciate, and participate in,the overall health management process.The adverse social, economic and environmentalimpacts that have resulted from the irresponsibleor ill-considered movement of live aquaticanimals and their products have led to globalrecognition of the need for health managementprotocols to protect aquaculture, fisheries resourcesand the aquatic environment. In manycases, these impacts have been a direct resultof the absence of effective national and regionalhealth management strategies. However, formulationof effective quarantine measures,health certification and guidelines applicable onan international scale is complicated. A widerange of social, economic and environmentalcircumstances have to be considered, alongwith the range of aquatic animal species involvedand their pathogens and diseases. Inaddition, differing reasons for moving liveaquatic animals and products impose a furtherset of variables to the process. Nevertheless,the serious impacts of unrestricted regional andinternational movement of aquatic animals meritinternational recognition - a fact clearly reflectedin the International Aquatic Animal Health Codeand the Diagnostic Manual of Aquatic AnimalDiseases of the Office International desÉpizooties 1 , which provide guidelines and recommendationsfor reducing the risk of spreadingspecific pathogens considered relevant tointernational trade of aquatic animals.Since present international protocols are notalways applicable to the disease concerns ofaquatic food production and trade in the AsiaRegion, the need for effective health managementprotocols that focus on the species anddisease problems of this region has been recognizedfor many years. A regional, as opposedto national, approach is considered appropri-ate, since many countries in the region sharesocial, economic, industrial, environmental, biologicaland geographical characteristics. Manycountries also share waterbodies withneighbours and the watersheds of several majorAsian rivers transcend national boundaries.A regionally adopted health management programwill facilitate trade, and protect aquaticproduction (subsistence and commercial) andthe environment upon which they depend, frompreventable disease incursions.A joint FAO/NACA Asia-Regional Programmeon Aquatic Animal Health Management was undertakento review the need for better healthmanagement to support safe movement of liveaquatic animals and the applicability of existinginternational codes on aquatic animal healthmanagement, quarantine and health certification,including those of the OIE, the EuropeanInland Fisheries Advisory Commission (EIFAC),and the International Council for Exploration ofthe Sea (ICES) to Asian circumstances. Thisreview 2 highlighted the fact that the diseaserisks associated with pathogen transfer in theAsia Region can only be reduced through abroader approach to aquatic animal healthmanagement than currently outlined in diseasespecificcodes of practice (e.g., the OIE code)or in codes and protocols developed specificallyfor northern hemisphere countries (e.g.,the ICES and EIFAC codes). In addition, it underlinedthe need for pre-border (exporter), borderand post-border (importer) involvement inthe program, to ensure co-operative healthmanagement of aquatic animal movement. Withthe support of an FAO Technical Co-operationProgramme (TCP) implemented by NACA, theAsia Regional Technical Guidelines on HealthManagement for the Responsible Movement ofLive Aquatic Animals is a document that wascompiled by a group of aquatic animal healthexperts within and outside the region to assistthe development of effective health managementprocedures for safe movement of liveaquatic animals within and between countriesin the region. The first companion document,the Manual of Procedures for the Implementationof the Asia Regional Technical Guidelineson Health Management for the ResponsibleMovement of Live Aquatic Animals, providesbackground material and detailed technical proceduresto assist countries and territories in the1see OIE. 2000a. International Aquatic Animal Health Code. 3rd edn. Office International des Epizooties, Paris, 153 p.; and OIE.2000b. Diagnostic Manual for Aquatic Animal Diseases. 3rd edn, Office International des Epizooties, Paris, 237 p.2see Humphrey, J.D., J.R. Arthur, R.P. Subasinghe and M.J. Phillips. 1997. Aquatic Animal Quarantine and Health Certificationin Asia. Proceedings of the Regional Workshop on Health and Quarantine Guidelines for the Responsible Movement (Introductionand Transfer of Aquatic Organisms), Bangkok Thailand, 28 January 1996. FAO Fish. Techn. Pap. No. 373, 153 p.viii 9

FOREWORDAsia Region in implementing the TechnicalGuidelines. This second companion document,Asia Diagnostic Guide, provides valuable diagnosticguidance for implementing the TechnicalGuidelines and also complementary to theManual of Procedures.10 ix

ACKNOWLEDGEMENTSVery special thanks go to Dr. Michael J. Phillipsof NACA for his vision and constant encouragement;NACA Co-ordinators, Mr. HassanaiKongkeo (1996-2001) and Mr. Pedro Bueno(2001 to present) for their strong support to theAsia regional program on aquatic animal health;and the team from Multimedia Asia for their creativeideas and friendly cooperation and quickresponse to the sometimes untimely demandsto complete the Asia Diagnostic Guide.The Editors12 xi

TABLE OF CONTENTSTitle PageDisclaimer and Copyright StatementsPreparation of This DocumentAbstractPrefaceForewordAcknowledgementsTable of ContentsGlossaryAbbreviationsScientific and Common NamesSECTION 1- INTRODUCTIONI. INTRODUCTIONI.1 BackgroundI.2 Objectives and ScopeI.3 Guide for UsersI.4 Health and Aquatic AnimalsI.5 Role of Diagnostics in Aquatic Animal HealthI.6 Levels of DiagnosticsI.7 ReferencesBasic Anatomy of a Typical Bony FishSECTION 2 - FINFISH DISEASESF.1 GENERAL TECHNIQUESF.1.1F.1.1.1F.1.1.2F.1.1.2.1F.1.1.2.2F.1.1.2.3F.1.1.3F.1.1.3.1F.1.1.3.2F.1.2F.1.3F.1.3.1F.1.3.2F.1.3.3F.1.3.4F.1.3.5F.1.3.6F.1.3.7F.1.3.8F.1.4F.1.4.1F.1.4.2F.1.4.3F.1.5Gross ObservationsBehaviourSurface ObservationsSkin and FinsGillsBodyInternal ObservationsBody Cavity and MuscleOrgansEnvironmental ParametersGeneral ProceduresPre-Collection PreparationBackground InformationSample Collection for Health SurveillanceSample Collection for Disease DiagnosisLive Specimen Collection for ShippingDead or Tissue Specimen Collection for ShippingPreservation of Tissue SamplesShipping Preserved SamplesRecord-KeepingGross ObservationsEnvironmental ObservationsStocking RecordsReferencesiiiiiivvviviiix131733353940404042434346485050505050515252525253535354545455555656575757575713

TABLE OF CONTENTSVIRAL DISEASES OF FINFISHF.2 Epizootic Haematopoietic Necrosis (EHN)F.3 Infectious Haematopoietic Necrosis (IHN)F.4 Oncorhynchus masou Virus (OMV)F.5 Infectious Pancreatic Necrosis (IPN)F.6 Viral Encephalopathy and Retinopathy (VER)F.7 Spring Viraemia of Carp (SVC)F.8 Viral Haemorrhagic Septicaemia (VHS)F.9 LymphocystisBACTERIAL DISEASE OF FINFISHF.10 Bacterial Kidney Disease (BKD)FUNGUS ASSOCIATED DISEASE FINFISHF.11 Epizootic Ulcerative Syndrome (EUS)59626568727679828690F.AIF.AIIF.AIIIANNEXESOIE Reference Laboratories for Finfish DiseasesList of Regional Resource Experts for FinfishDiseases in Asia-PacificList of Useful Diagnostic Manuals/Guides toFinfish Diseases in Asia-Pacific9598105Basic Anatomy of an OysterSECTION 3 - MOLLUSCAN DISEASESM.1 GENERAL TECHNIQUESM.1.1M.1.1.1M.1.1.2M.1.1.3M.1.1.4M.1.2M.1.3M.1.3.1M.1.3.2M.1.3.3M.1.3.4M.1.3.5M.1.3.6M.1.3.7M.1.4M.1.4.1M.1.4.2M.1.4.3M.1.5Gross ObservationsBehaviourShell Surface ObservationsInner Shell ObservationsSoft-Tissue SurfacesEnvironmental ParametersGeneral ProceduresPre-Collection PreparationBackground InformationSample Collection for Health SurveillanceSample Collection for Disease DiagnosisLive Specimen Collection for ShippingPreservation of Tissue SamplesShipping Preserved SamplesRecord KeepingGross ObservationsEnvironmental ObservationsStocking RecordsReferencesDISEASES OF MOLLUSCSM.2 Bonamiosis (Bonamia sp., B. ostreae)M.3 Marteiliosis (Marteilia refringens, M. sydneyi)M.4 Mikrocytosis (Mikrocytos mackini, M. roughleyi)10811011111111111111411411611611611611611611711811811811911911912112512914

TABLE OF CONTENTSM.5 Perkinsosis (Perkinsus marinus, P. olseni)M.6 Haplosporidiosis (Haplosporidium costale,H. nelsoni)M.7 Marteilioidosis (Marteilioides chungmuensis,M. branchialis)M.8 Iridovirosis (Oyster Velar Virus Disease)133138144147M.AIM.AIIM.AIIIANNEXESOIE Reference Laboratories forMolluscan DiseasesList of Regional Resource Experts forMolluscan Diseases in Asia-PacificList of Useful Diagnostic Guides/Manuals toMolluscan Health149150152Internal and External Anatomy of a Penaeid ShrimpSECTION 4 - CRUSTACEAN DISEASESC.1 GENERAL TECHNIQUESC.1.1 Gross ObservationsC.1.1.1 BehaviourC.1.1.1.1 GeneralC.1.1.1.2 MortalitiesC.1.1.1.3 FeedingC.1.1.2 Surface ObservationsC.1.1.2.1 Colonisation and ErosionC.1.1.2.2 Cuticle Softening, Spots and DamageC.1.1.2.3 ColourC.1.1.2.4 Environmental ObservationsC.1.1.3 Soft-Tissue SurfacesC.1.2 Environmental ParametersC.1.3 General ProceduresC.1.3.1 Pre-collection PreparationC.1.3.2 Background InformationC.1.3.3 Sample Collection for Health SurveillanceC.1.3.4 Sample Collection for Disease DiagnosisC.1.3.5 Live Specimen Collection for ShippingC.1.3.6 Preservation of Tissue SamplesC.1.3.7 Shipping Preserved SamplesC.1.4 Record-KeepingC.1.4.1 Gross ObservationsC.1.4.2 Environmental ObservationsC.1.4.3 Stocking RecordsC.1.5 ReferencesVIRAL DISEASES OF SHRIMPC.2 Yellowhead Disease (YHD)C.3 Infectious Hepatopancreas and HaematopoieticNecrosis (IHHN)C.4 White Spot Disease (WSD)C.4a Bacterial White Spot Syndrome (BWSS)15615715715715715715815815815815816016016016016016216216216216416516516516516616616717317818315

TABLE OF CONTENTSC.5 Baculoviral Midgut Gland Necrosis (BMN)C.6 Gill-Associated Virus (GAV)C.7 Spawner Mortality Syndrome("Midcrop mortality syndrome")C.8 Taura Syndrome (TS)C.9 Nuclear Polyhedrosis Baculovirosis (NPD)BACTERIAL DISEASE OF SHRIMPC.10 Necrotising Hepatopancreatitis (NH)FUNGAL DISEASE OF CRAYFISHC.11 Crayfish Plague186189192194201207211C.AIC.AIIC.AIIIANNEXESOIE Reference Laboratories forCrustacean DiseasesList of Regional Resource Experts for CrustaceanDiseases in the Asia-PacificList of Useful Manuals/Guide to CrustaceanDiseases in Asia-Pacific215216219List of National Coordinators(NCs)Members of Regional Working Group (RWG) andTechnical Support Services (TSS)List of Figures22122523016

GLOSSARY 1AbscessAbiotic factorsan aggregation of haemocytes (blood cells) associated with necrotic(decaying) host cells. Abscesses may or may not contain debris frominvasive organisms which have been killed by host defences. In advancedabscesses there is a decrease in cell definition (especially the nuclei)towards the centre of the lesion, compared to cells around the periphery.Abscesses frequently involve breakdown of epithelial linings and may besurrounded by phagocytic and/or fibrocytic haemocytes.physical factors which affect the development/survival of an organismAcquired immunity defence response developed following recovery from an infection (orvaccination) to a specific infectious agent (or group of agents)AcuteAdhesionAetiologic Agent(Etiologic)infection or clinical manifestation of disease which occurs over a shortperiod of time (cf 'Chronic')(Crustacea) binding of subcuticular tissues to the cuticle due to destructionof the cuticle by chitinolytic bacteria or fungi. This may impede moulting.the primary organism responsible for changes in host animal, leading todiseaseAetiology (Etiology) the study of the cause of disease, including the factors which enhancetransmission and infectivity of the aetiologic agent.AlevinsAnaemiaAnorexiaAntennal glandAntibody (Ab)AntigenAquatic animalsAquacultureAscitesAsepticfry of certain species of fish, particularly trout and salmonids that still havethe yolk-sac attached(Vertebrate) a deficiency in blood or of red blood cellsloss of appetite(Crustacea) excretory pores at the base of the antennae (also known askidney gland, excretory organ and green gland)a protein capable of cross-reacting with an antigen. In vertebrates,antibody is produced by lymphoid cells in response to antigens. Themechanism of antibody production in shellfish is not known.a substance or cell that elicits an immune reaction. An antigen may haveseveral epitopes (surface molecules) to which antibody can bind (cfMonoclonal and Polyclonal Antibodies).live fish, molluscs and crustaceans, including their reproductive products,fertilised eggs, embryos and juvenile stages, whether from aquaculturesites or from the wildcommonly termed "fish farming", it refers more broadly to the commercialhatching and rearing of marine and freshwater aquatic animals and plantsaccumulation of serous fluid in the abdominal cavity; dropsyfree from infection; sterile1Definitions of words with * were adopted from OIE International Aquatic Animal Health Code. 3rd Edition. 2000. All otherdefinitions were taken from the following references: FAO/NACA (2000); Dorland's Illustrated Medical Dictionary (27th Edition);"Virology Glossary" copyright 1995 by Carlton Hogan and University of Minnesota (permission to copy and distribute granted toindividuals and non-profit groups http://www.virology.net/ATVG;ossary.html); On-line Medical Dictionary athttp://www.graylab.ac.uk/omd/index.html.17

GLOSSARYAtrophyAutolysis(-lytic)AvirulentAxenic cultureBacteriologyBacteriophageBacteriumBasophilicBioassayBroodstock*CalcareousCannibalismCarrierCeroidChelating agentdecrease in amount of tissue, or size of an organ, after normal growth hasbeen achievedenzyme induced rupture of cell membranes, either as a normal function ofcell replacement or due to infectionan infection which causes negligible or no pathology (cf Virulent).culture containing cells of a single species (bacterial culture) or cell-type(tissue culture) (uncontaminated or purified)science that deals with the study of bacteria(abbreviation - Phage) any virus that infects bacteria(bacteria) unicellular prokaryotic (nuclear material not contained within anucleus) microorganisms that multiply by cell division (fission), typicallyhave a cell wall; may be aerobic or anaerobic, motile or non-motile, freeliving,saprophytic or pathogenicacidic cell and tissue components staining readily with basic dyes (i.e.hematoxylin); chromatin and some secretory products in stained cellsappear blue to purplea quantitative procedure that uses susceptible organisms to detect toxicsubstances or pathogens.sexually mature fish, molluscs or crustaceanspertaining to or containing lime or calciumthe eating of a species of animal by the same species of animalan individual who harbors the specific organisms of a disease withoutmanifest symptoms and is capable of transmitting the infection; thecondition of such an individual is referred to as carrier statenon-staining metabolic by-product found in many bivalves. Abnormallyhigh concentrations indicate possible environmental or pathogen-inducedphysiological stress.chemical agent used to decalcify calcium carbonate in mollusc shells orpearls, e.g., ethylenediaminetetracetic acid (EDTA)Chemotherapeutant chemical used to treat an infection or non-infectious disorderChitinChitinolytic(chitinoclastic)ChronicClinicalChromatinlinear polysaccharide in the exoskeletons of arthropods, cell walls of mostfungi and the cyst walls of ciliates(Mycology and Bacteriology) chitin degrading organisms with enzymescapable of breaking down the chitin component of arthropod exoskeletonslong-term infection which may or may not manifest clinical signspertaining to or founded on actual observationnucleoprotein complex containing genomic DNA and RNA in the nucleusof most eukaryotic cells18

GLOSSARYChromatophoresCiliostaticCloneCoagulationConchiolinConcretionsContagiousCrustaceans*CuticleCystCytologyCytopathic effectDecalcificationDecapitationDeoxyribovirusDFATDiapedesisDiseaseDisease agentDiagnosis*Disinfection*motile, pigment-containing epidermal cells responsible for colourexotoxin toxin secreted by some bacteria that inhibits ciliary functionsa population derived from a single organismclotting (adhesion of haemocytes)nitrogenous albuminoid substance, dark brown in colour, that forms theorganic base of molluscan shellsnon-staining inclusions in the tubule and kidney cells of scallops and pearloysters, produced during the digestive cycle. Similar inclusions are alsofound in the gut epithelia of other bivalves.a disease normally transmitted only by direct contact between infectedand uninfected organismsaquatic animals belonging to the phylum Arthropoda, a large class ofaquatic animals characterized by their chitinous exoskeleton and jointedappendages, e.g. crabs, lobsters, crayfish, shrimps, prawns, isopods,ostracods and amphipods(Crustacea) the protein structure of arthropods consisting of an outer layer(epicuticle), an underlying exocuticle (pigmented), endocuticle (calcified)and membranous uncalcified layer. Chitin is in all layers except theepicuticle.(a) a resilient dormant stage of a free-living or parasitic organism, or(b) a host-response walling off a tissue irritant or infectionthe study of cells, their origin, structure, function and pathologypertaining to or characterized by pathological changes in cellsthe process of removing calcareous mattercutting of the head portion(DNA-virus) virus with a deoxyribonucleic acid genome (cf Ribovirus)Direct Fluorescent Antibody Test/Technique; an immunoassay techniqueusing antibody labelled to indicate binding to a specific antigenmigration of haemocytes across any epithelium to remove metabolic byproduct,dead cells and microbial infectionsany deviation from or interruption of the normal structure or function of anypart, organ, or system (or combination thereof) of the body that ismanifested by a characteristic set of symptoms and signs and whoseaetiology, pathology and prognosis may be known or unknownan organism that causes or contributes to the development of a diseasedetermination of the nature of a diseasethe application, after thorough cleansing, of procedures intended todestroy the infectious or parasitic agents of diseases of aquatic animals;this applies to aquaculture establishments (i.e. hatcheries, fish farms,19

GLOSSARYobjects that may have been directly or indirectly contaminatedDNA (ssDNA,dsDNA)DNA probesDropsyEcdysal glandEctoparasiteELISAEmaciationEndemicEndothelialEndotheliumEndosymbiosisEnvelopeEnzooticEosinophilicEpibiontEpipoditeEpitopeEpizooticEpidemiologydeoxyribonucelic acid. Nucleic acid comprised of deoxyribonucleotidescontaining the bases adenine, guanine, cytosine and thymine.Single strand DNA (ssDNA) occurs in some viruses (usually as a closedcircle). In eukaryotes and many viruses, DNA is double-stranded (dsDNA).segments of DNA labelled to indicate detection of homologous segmentsof DNA in samples of tissues or cultures (see RNA probes)the abnormal accumulation of serous fluid in the cellular tissues or in abody cavity(Crustacea) see Y-organa parasite that lives on the outside of the body of the hostEnzyme Linked Immunosorbent Assay, used to detect antigen (antigencapture ELISA) or antibody (antibody capture ELISA)a wasted condition of the bodypresent or usually prevalent in a population or geographical area at alltimespertaining to or made up of endotheliumthe layer of epithelial cells that lines the cavities of the heart and of theblood and lymph vessels, and the serous cavities of the body originatingfrom the mesoderman association between two organisms (one living within the other) whereboth derive benefit or suffer no obvious adverse effect(Virology) lipoprotein membrane composed of host lipids and viralproteins (non-enveloped viruses are composed solely of the capsid andnucleoprotein core)present in a population at all times but, occurring only in small numbersof casesbasic cell and tissue components staining readily with acidic dyes (i.e.eosin); stained cells appear pink to redorganisms (bacteria, fungi, algae, etc.) which live on the surfaces (cffouling) of other living organisms(Crustacea) cuticular extension of the base (protopodite) of the walkinglegs (pereiopods)the component of an antigen which stimulates an immune response andwhich binds with antibodyaffecting many animals within a given are at the same time; widely diffusedand rapidly spreading (syn. Epidemic - used for human disease)science concerned with the study of the factors determining and influencing the frequency and distribution of disease or other health related events20

GLOSSARYand their causes in a defined population for the purpose of establishingprograms to prevent and control their development and spreadEpizootiologyEpitheliumEpitopeErosionEukaryoteanExoenzymeExopthalmiaExoskeletonExudateEuthanasiaFiltrationFinfish*FryFingerlingFixationFixativeForeign bodiesFormalinFoulingFungusthe study of factors influencing infection by a pathogenic agentthe layer of cells covering the surface of the body and all gastrointestinallinings. Epithelia are usually one cell thick and supported by a basalmembrane.structural component of an antigen which stimulates an immune responseand which binds with antibody.destruction of the surface of a tissue, material or structureorganism that contains the chromosones within a membrane-boundnucleus (cf Prokaryote)extracellular enzyme released by a cell or microorganismabnormal protrusion of the eyeballs(Crustacea) the chitin and calcified outer covering of crustaceans (andother arthropods) which protects the soft-inner tissuesmaterial, such as fluid, cells, or cellular debris, which has escaped fromblood vessels and has been deposited in tissues or on tissue surfaces,usually as a result of inflammationan easy or painless deathpassage of a liquid through a filter, accomplished by gravity, pressure orvacuum (suction)fresh or saltwater fish of any agenewly hatched fish larvaea young or small fishpreservation of tissues in a liquid that prevents protein and lipidbreakdown and necrosis; the specimen is hardened to withstand furtherprocessing; and the cellular and sub-cellular contents are preserved in amanner close to that of the living statea fluid (e.g. aldehyde or ethanol-based solutions)) that prevents denaturation and autolysis by cross-linking of proteinsany organism or abiotic particle not formed from host tissuea 37% solution of formaldehyde gasthe mass colonisation of hard substrates by free-living organisms. Extremefouling of living organisms, such as molluscs or shrimp, can impede theirnormal body-functions leading to weakening and deathany member of the Kingdom Fungi, comprising single-celled or multinucleate organisms that live by decomposing and absorbing the organicmaterial in which they growoyster farms, shrimp farms, nurseries), vehicles, and different equipment/21

GLOSSARYGapingGram's StainGranulomasGranulomatosisGranulosis virusGross signsHaematopoieticHaematopoietictissueHaemocytesHaemolymphHaemocyteinfiltrationHaemocytopeniaHaemocytosisHaemorrhageHatcheries*HepatopancreasHistologyHistolysisHistopathologyweakened molluscs that cannot close their shells when removed fromwater; this rapidly lead to desiccation or predation of the soft-tissues andis indicative of molluscs in poor condition (including possible infection)stain used to differentiate bacteria with permeable cells walls (Gramnegative)and less permeable cell walls (Gram-positive)any small nodular delimited aggregation of granular haemocytes, ormodified macrophages resembling epithelial cells (epithelioid cells)any condition characterized by the formation of multiple granulomasBaculoviridae belonging to subgroup (B), characterised by a singlenucleocapsid within an envelope. Granulosis viruses form intra-nuclearellipsoid or rounded occlusion bodies (granules or capsules) containingone or two virions.signs of disease visible to the naked eyepertaining to or effecting the formation of blood cells(Decapoda) a sheet of tissue composed of small lobulessurrounded by fibrous connective tissue which lies along the dorso-lateralsurfaces of the posterior portion of the cardiac stomach (Brachyura) orsurrounding the lateral arterial vessels, secondary maxillipeds andepigastric tissues (Penaeidae and Nephropidae); (Bivalves) unknown;(Vertebrates) spleenblood-cellscell-free fraction of the blood containing a solution of protein and nonproteinaceousdefensive moleculesaccumulation of haemocytes around damaged or infected tissues; sincethe type of haemocytes most commonly responsible for phagocytosis aregranulocytes, focal infiltration is often referred to as a "granuloma"a reduction in the number of cells in the circulatory system, usuallyassociated with a reduction in blood-clotting capabilitysystemic destruction of blood cells (syn. Haemolysis)(Vertebrate) escape of blood from the vessels; bleeding(Invertebrate) uncontrolled loss of haemocytes due to tissue trauma,epithelial rupture, chronic diapedesisaquaculture establishments raising aquatic animals from fertilized eggsdigestive organ composed of ciliated ducts and blind-ending tubules,which secrete digestive enzymes for uptake across the digestive tubuleepithelium; also responsible for release of metabolic by-products and othermolecular or microbial wastes (cf Metaplasia, Diapedesis)the study that deals with the minute structure, composition and function oftissuesbreakdown of tissue by disintegration of the plasma membranesstructural and functional changes in tissues and organs of the body which22

GLOSSARYcause or are caused by a disease seen in samples processed byhistologyHomogenate tissue ground into a liquid state in which all cell structure is disintegratedHost individual organism infected by another organismHusbandry management of captive animals to enhance reproduction, growth andhealthHyperplasia abnormal increase in size of a tissue or organ due to an increase innumber of cellsHypertrophy abnormal enlargement of cells due to irritation or infection by anintracellu lar organism.Hyphae (Mycology) tubular cells of filamentous fungi; may be divided by crosswalls(septae) into multicellular hyphae, may be branched. Interconnectinghyphae are called mycelia.Icosahedral shape of viruses with a 5-3-2 symmetry and 20, approximatelyequilateral, triangular facesIFAT Indirect Fluorescent Antibody Test/Technique; a technique usingunlabelled antibody and a labelled anti-immunoglobulin to form a'sandwich' with any antigen-bound antibodyImmunity protection against infectious disease conferred either by the immuneresponse generated by immunization or previous infection or by othernon-immunologic factorsImmunization protection against disease by deliberate exposure to pathogenantigens to induce defence system recognition and enhance subsequent responses to exposure to the same antigens (syn Vaccination)Immunoassay any technique using the antigen-antibody reaction to detect andquantify the antigens, antibodies or related substances (see ELISA,IFAT, DFAT)ImmunodepressionImmunofluorescencedecrease in immune system response to antigens due to an infection(same or different agent) or exposure to an immunosupressantchemical.(syn. Immunosupression)any immuno-histochemical method using antibody labeled with afluorescent dyeDirect - if a specific antibody or antiserum with a fluorochrome andused as a specific fluorescent stainIndirect - if the fluorochrome is attached to an antiglobulin, and atissue constituent is stained using an unlabeled specific antibody andthe labeled antiglobulin, which binds the unlabeled antibodyImmunoglobulin (Ig)family of proteins constructed of light and heavy molecular weightchains linked by disulphide bonds; usually produced in response toantigenic stimulationImmunohistochemistry application of antigen-antibody interactions to histochemical tech23

GLOSSARYniques, as in the use of immunofluorescenceImmunologybranch of biomedical science concerned with the response of theorganisms to antigenic challenge, the recognition of self and not self, andall the biological (in vivo), serological (in vitro), and physical chemicalaspects of immune phenomenaImmunostimulation enhancement of defense responses, e.g., with vaccinationImmunizationInclusion bodyInfectiousInfectionInfiltrationInflammationinduction of immunitynon-specific discrete bodies found within the cytoplasm or nucleus of acell. Frequently viral (cf Cowdry body, Polyhedrin Inclusion /OcclusionBodies), or bacterial microcolonies (cf RLOs) (syn. Inclusions)capable of being transmitted or of causing infectioninvasion and multiplication of an infectious organism within host tissues.May be clinically benign (cf sub-clinical or 'carrier') or result in cell or tissuedamage. The infection may remain localized, subclinical, and temporary ifthe host defensive mechanisms are effective or it may spread an acute,sub-acute or chronic clinical infection (disease).(Invertebrates) haemocyte migration to a site of tissue damage or infectionby a foreign body/organism ('inflammation'). Infiltration may also occur forroutine absorption and transport of nutrients and disposal of wasteproducts.(Vertebrate) initial response to tissue injury characterised by the release ofamines which cause vasodilation, infiltration of blood cells, proteins andredness that may be associated with heat generation(Invertebrates) infiltration response to tissue damage or a foreign body. Theinfiltration may be focal, diffuse or systemic (syn. Infiltration).Innate immunityIntensity ofinfectionIntercellularInterstitial tissueIntracellularIntrapallialKaryolysisKaryorrhexicLesionhost defence mechanism that does not require prior exposure to thepathogenthe number of infectious agents in an individual organism or specimen;"mean" intensity is the average number of infectious agents present in allinfected individuals in a samplesituated or occurring between the cells in a tissuetissue or cells between epithelial bound organ systems; also known as (cells)Leydig tissue (molluscs) or connective tissuesituated or occurring within a cell(Bivalves) space between the mantle, gills and other soft-tissues; thespace between the mantle and inner shell is the extrapallial spacea form of necrosis where the chromatin leaches out of the nucleus withoutdisrupting the nuclear membrane, leaving an 'empty' appearing nucleusrupture of the nucleus and nuclear membrane, releasing chromatingranules into the cytoplasmany pathological or traumatic change in tissue form or function24

GLOSSARYLethargyLiquefactionLuminescentLymphoid organLymphoid organabnormal drowsiness or stupor (response only to vigorous stimulation); acondition of indifferenceconversion of a tissue into a semi-solid or fluid mass due to necrosismarine or euryhaline bacteria which contain luciferase (a fluorescent, bacteriaenzyme)e.g., Vibrio harveyi and V. splendidus(Crustacea) an organ situated between the anterior and posterior stomachchambers which connects the sub-gastric artery to the anterior aorta, via amass of interconnected tubulesspherical cellular masses composed of presumed phagocytic haemocytes,spheres which sequester Taura Syndrome Virus (TSV) and aggregatewithin intertubular spaces of the lymphoid organsMacrophages (Vertebrates) large (10-20 mm) amoeboid blood cells, responsible for phagocytosis, inflammation, antibody and cytotoxin production.Mandibular organMantle retraction/recessionMelaninMelanisationMelanophoresMetaplasiaMicrocoloniesMicroorganismMolecular probesMolluscs*Monoclonalantibody(MaB)MoribundMortality(Crustacea) large glandular organ close to the ventral epidermis betweenthe mandibles; believed to be related to the moulting cycle, although itdoes not produce a known moult-inducing hormoneduring periods of no growth in molluscs, the mantle retracts away fromthe edge of the shell. Prolonged mantle retraction leaves the inner shelledge open to erosion and fouling.dark brown-black polymer (pigment) of indole quinone which has enzymeinhibiting properties. It forms part of the primary defence mechanismagainst cuticle and epidermal damage in many crustaceansabnormal deposits of dark pigment in various organs or tissues(Crustacea) dermal cells containing melanin (syn. melanocytes)the change in shape of any epithelial cell, e.g., from columnar to cuboidalor squamous (flattened)membrane-bound populations of Chlamydia bacteria or non-membranebound Rickettsial colonies (cf Inclusion bodies)principally, viruses, bacteria and fungi (microscopic species, and taxonomically-related macroscopic species). Microscopic protistans (Protozoa)and algae may also be referred to as microoorganisms.see DNA probesaquatic organism belonging to the Phylum Mollusca in the KingdomMetazoa characterized by soft unsegmented bodies. Most forms areenclosed in a calcareous shell. The different developmental stages ofmolluscs are termed larvae, postlarvae, spat, juvenile and adult.identical antibody molecules produced by clonage of the antibodyproducing cell and responsive to a single antigen epitope (cf Epitope)diseased; near deathdeath25

GLOSSARYMoultingMucousMucusMultiple aetiologyMycelial coloniesMyceliumMycologyMycosisMyodegenerationMysis larvaeNacreNauplius(-plii)NecrosisNotifiableDiseases*NuclearPolyhedrosisVirus (NPV)NucleocapsidOcclusionOcclusion body(Crustacea) the shedding of the exoskeleton to permit growth (increase insize) of internal soft-tissues (syn. Ecdysis)pertaining or relating to, or resembling mucusthe free slime of the mucous membrane, composed of secretion of theglands, along with various inorganic salts, desquamated cells andleukocytesdisease associated with more than one infectious agent; may be directlyattributed to one or more infectious organism (cf Syndrome)(Bacteriology) colony growth of Gram-positive Actinomycete bacteria withbranched mycelia which may fragment into rods or coccoid forms(Mycology) network formed by interconnecting hyphae (syn. Mycelialnetwork)the study of fungi (Mycota)any disease resulting from infection by a fungusbreakdown of muscle fibres(Crustacea) pelagic larval stage between protozoea (zoeal) and post larvainner layer of molluscan shells; may have an iridescent crystal matrix(mother-of-pearl)(Crustacea) earliest larval stage; with three pairs of appendages,uniramous first antennae, biramous second antennae and mandiblessum of the morphological changes indicative of cell death and caused bythe progressive and irreversible degradative action of enzymes; it mayaffect groups of cells or part of a structure or an organ; necrosis may takedifferent forms and be associated with saprobionts (bacterial, fungal orprotistan) proliferation.'diseases notifiable to the OIE' means the list of transmissible diseasesthat are considered to be of socio-economic and/or public health importance within countries and that are significant in the international trade inaquatic animals and aquatic animal products (see also OIE 1997, OIE2000a, b)Baculoviruses (Type A) which produce intranuclear polyhedral proteinmatrices (see Polyhedral Occlusion/Inclusion Bodies)protein-nucleic acid complex which may form the core, capsid and/orhelical nucleoprotein of the virion(vascular) filling or blocking of vascular sinuses by haemocytes; (perivascular) infiltration of haemocytes, several cells deep into the tissues surrounding vascular sinuses; (luminal) filling or blocking of gonoducts, renal ducts,digestive tubules or ducts by haemocytes or other cell debris(see Polyhedrin Inclusion/Occlusion Body)26

GLOSSARYOedema (edema)OpportunisticOsmoregulationOther SignificantDiseases*OutbreakOvertParasiteParasitologyPassagePatent infectionPathogenPathogenicityPathognomonicPathologyPCRPereiopodsPeriostracumPhagesPhagocytosispresence of abnormally large amounts of fluid in the intercellular spaces ofthe bodyorganism capable of causing disease only when a host's resistance ispathogen lowered by other factors (another disease, adverse growingconditions, drugs, etc.)maintenance of osmolarity by a simple organism or body cell with respectto the surrounding mediumdiseases that are of current or potential international significance inaquaculture, but that have not been included in the list of diseasesnotifiable to the OIE because they are less important than the 'notifiablediseases', or because their geographical distribution is limited, or is toowide for notification to be meaningful,or it is not yet sufficiently defined, orbecause the aetiology of the disease is not well enough understood, arapproved diagnostic methods are not available (see also OIE 1997, OIE2000a, b)the sudden onset of disease in epizootic proportionsopen to view; not concealedan organism which lives upon or within another living organism (host) atwhose expense it obtains some advantage, generally nourishmentscience that deals with the study of parasites(Virology) the successive transfer of a virus or other infectious agentthrough a series of experimental animals, tissue culture, or syntheticmedia with growth occurring in each mediumperiod when clinical signs and/or the infectious organism can be detected(cf Prepatent)an infectious agent capable of causing diseasethe ability to produce pathologic changes or diseasesign or symptom that is distinctive for a specific disease or pathologic conditiondeals with the essential nature of disease, especially of the structural andfunctional changes in tissues and organs of the body which cause or arecaused by a diseasePolymerase Chain Reaction, a process by which nucleic acid sequencescan be replicated ('nucleic acid amplification')(Crustacea) thoracic appendages ('walking legs') (cf Pleopods andUropods)(Molluscs) calcareous layers of shell which may contain quinine-tannedprotein(see Bacteriophage)uptake by a cell of material from the environment by invagination of itsplasma membrane27

GLOSSARYPlasma membranetrilaminar membrane enclosing the cytoplasm and organelles of a cellPleiopod small legs of some crustaceansPleomorphic organism demonstrating more than one body form within a life-cyclePolyadenalated messenger RNA (mRNA) which has a polyadenylate sequence bound toRNA the 3' end of the molecule. This is common in most eukaryote mRNAand is present in some riboviruses. The function of this addition isunknown.Polyclonal (more correctly, but rarely, termed 'Polyclonal antiserum') anantibodies antiserum prepared from an organism exposed to an antigen. The PAb(PAb) contains several different antibodies, each specific to a different epitopeof the same antigen. (see Monoclonal antibody).Polyhedral Inclusion/ protein-based crystalline matrix made up ofOcclusion Body Polyhedrin (Baculovirus group A - Nuclear Polyhedrosis Viruses(POB, PIB) (NPV)) or Granulin (Baculovirus group B - Granulosis Viruses (GV)).Baculovirus group C do not form occlusion bodies.Polymorphic (a) capability of molecules, such as enzymes, to exist in several forms;(b) ability of nuclei of certain cells (e.g., haemocytes) to change shape;and (c) ability of microorganisms to change shape (e.g., in different hostspecies or tissues)Pop-eye abnormal protrusion of the eyes from the eye socketsPostlarvae the stage following metamorphosis from larvae to juvenile in the(PL) life cycle of Crustacea. In penaeid shrimp, this is commonly counted indays after appearance of postlarval features, e.g., PL12 indicates apost-larvae that has lived 12 days since its metamorphosis from thezoea stage of development.Predator an organism that derives elements essential for its existence fromorganisms of other species, which it consumes and destroysPredispose to make susceptible to a disease which may be activated by certainconditions, as by stressPreening (Crustacea) cleaning surface tissues or eggs exposed to fouling (cfEpibionts and Fouling); some crustaceans have modified appendages toenhance preening (e.g., the gill-rakers of Brachyura)Prepatent period period between infection and the manifestation of clinical or detectablesigns of diseasePrevalence percentage of individuals in a sample infected by a specific disease,parasite or other organismProkaryote (syn. Bacteria) cellular micro-organisms in which the chromsones are notenclosed within a nucleusProphylactic (-axis):action or chemotherpeutant administered to healthy animals in order toprevent infection (see Treatment)Pustule a sub-epidermal swelling containing necrotic cell debris as a result ofinflammation (haemocyte infiltration) in response to a focal infection28

GLOSSARYPutativePyknosis/PyknoticQuarantinesignifies that which is commonly thought, reputed or believedcontraction of nuclear contents to a deep staining (basophilic) irregularmass, sign of death cell (cf Karryorhexis and Karyolysis)holding or rearing of aquatic animals under conditions which prevent theirescape, and the escape of any pathogens they may be carrying, into thesurrounding environment. This usually involves sterelisation/disinfection ofall effluent and quarantine materials.Quarantine measures are measures developed as a result of risk analysisto prevent the transfer of disease agents with live aquatic animal movements, with pre-border, border and post-border health managementprocesses, however, such activities are equally applicable to intra-nationalmovements of live aquatic animal.RepairReservoirResistanceResistanceRibosomesRibovirus(RNA-virus)RiskRNARNA probesrRNASaprobiontsSchizontsprocess to re-establish anatomical and functional integrity of tissues afteran injury or infection(host or infection) an alternate or passive host or carrier that harborspathogenic organisms, without injury to itself, and serves as a source fromwhich other individuals can be infected(to Disease) (cf Acquired immunity and Innate immunity) the capacity of anorganism to control the pathogenic effects of an infection. Resistance doesnot necessarily negate infection ('Refraction') and varying degrees oftolerance to the infection may be manifest. Heavy sub-clinical infectionsare indicative of resistance (syn. Tolerance; opp. Susceptible)(Antibiotic or 'drug' resistance) the capability of a microbe to evadedestruction by an antibiotic. This may arise from changes in the antigenicproperties of the microbe. Survival and multiplication leads to developmentof drug resistant strains of the pathogen. This may confer resistance torelated (heteroresistance) or non-related antibiotics (multiple drugresistance).intracytoplasmic granules which are rich in RNA and function in proteinsynthesisvirus with a ribonucleic acid (see RNA) genome (see Deoxyribovirus)the probability of negative impact(s) on aquatic animal health, environmental biodiversity and habitat and/or socio-economic investment(s)ribonucleic acid consisting of ribonucleotides made up of the bases (ssRNA,dsRNA) adenine, guanine, cytosine and uracilsegments of RNA which are labelled to detect homologous segments ofRNA or DNA in tissue or culture samples (cf DNA probes)(Ribosomal RNA) RNA component of the ribonucleoprotein organelleresponsible for protein synthesis within a cell(syn. Saprotroph) organisms which obtain nutrition from dead organicmatterthe multinucleated stage or form of development during schizogony29

GLOSSARYSecondarySepticaemiaSerologySerumShipment*SporangiumSporangiumSporeSporogenesisSterilizationStressSub-clinicalSurveillance*SusceptibleSyndromeSynergisticSystemicSystemic infectionTail rotTelsoninfection infection resulting from a reduction in the host's resistance as aconsequence of an earlier infectionsystemic disease associated with the presence and persistence ofpathogenic microorganisms or their toxins in the blood; blood poisoningterm now used to refer to the use of such reactions to measure serumantibody titers in infectious disease (serologic tests), to the clinicalcorrelations of the antibody titer (the 'serology' of a disease) and the use ofserologic reactions to detect antigensfluid component of coagulated haemolympha group of aquatic animals or products thereof destined for transportation(Mycology) hyphal swelling which contains motile or non-motile zoospores;release is via a pore or breakdown of the sporangial wall. (syn. Zoosporangium)(Bacteriology) the cell, or part of a cell, which subsequently develops intoan endospore (intracellularly formed spore)infective stage of an organism that is usually protected from the environment by one or more protective membranes (syn. Zoospores)formation of or reproduction by spores; sporulationany process (physical or chemical) which kills or destroys all contaminatingorganisms, irrespective of type; a sterile environment (aquatic or solid) isfree of any living organismthe sum of biological reactions to any adverse stimuli (physical, internal orexternal) that disturb the organism's optimum operating status(asymptomatic) an infection with no evident symptoms or clinical signs ofdisease, or a period of infection preceding the onset of clinical signs (cfPrepatent)a systematic series of investigations of a given population of aquaticanimals to detect the occurrence of disease for control purposes, andwhich may involve testing of samples of a populationan organism which has no immunity or resistance to infection by a anotherorganisman assembly of clinical signs which when manifest together are indicativeof a distinct disease or abnormality (syn. Pathognomic/ Pathognomonic)(infection) pathology increased by two or more infections by differentagents, compared with the effect from individual effects (opp. to 'antagonistic' or 'suppressive', where one infection counteracts the other)pertaining to or affecting the body as a wholean infection involving the whole bodydisintegration of tail and fin tissue(Crustacea) terminal segment of the abdomen which overlies the uropods30

GLOSSARYTomontTransmissionthe non-feeding, dividing stage or form in the life cycle of certain protozoathat typically encysts and produces tomites by fissiontransfer of an infectious agent from one organism to anotherHorizontal - direct from environment (e.g., via ingestion, skin and gills)Vertical - prenatal transmission (i.e., passed from parent to egg); may beeither inside the egg (intra-ovum) or through external exposure to pathogens from the parent generationTransportTraumaTreatmentactionTrophozoitesTumourUbiquitousUlcerUropodsVaccineVacuolatedVeligerVelum (Velar)ViableVirionmovement of stocks between locations by human influencean effect of physical shock or injurytaken to eradicate an infection (cf Prophylaxis)the active, motile, feeding stage of a protozoan organism, as contrastedwith the non-motile encysted stageabnormal growth as a result of uncontrolled cell division of a localisedgroup of cellsexisting or being everywhereexcavation of the surface of an organ or tissue, involving sloughing ofnecrotic inflammatory tissue.(Crustacea) the terminal appendages underlying the telson that form the'tail fan' (see Pereiopods and Pleopods)an antigen preparation from whole or extracted parts of an infectiousorganism, which is used to enhance the specific immune response of asusceptible hostcontaining spaces or cavities within the cytoplasm of a cell(Mollusc) ciliated planktonic larval stage(Mollusc) ciliated feeding surface of veliger larvaecapable of living or causing a diseaseindividual viral particlecontaining nucleic acid (the nucleoid), DNA or RNA(but not both) and a protein shell, or capsidVirogenic stroma(e) site of viral replication or assembly (syn. Viroplasm)VirogensisVirologyVirulenceproduction of virionsbranch of microbiology which is concerned with the study of viruses andviral diseasesthe degree of pathogenicity caused by an infectious organism, as indicatedby the severity of the disease produced and its ability to invade the tissuesof the host; the competence of any infectious agent to producepathologic effects; virulence is measured experimentally by the medianlethal dose (LD 50) or median infective dose (ID 50)31

GLOSSARYVirusone of a group of minute infectious agents, characterized by a lack ofindependent metabolism and by the ability to replicate only within livinghost cellsY-organ (Crustacea) (syn. Ecdysal gland) gland resonsible for production of the moultinghormone ecdysone. Production of the moulting hormone is controlled by amoult inhibiting hormone synthesised in the eye-stalkZoea larvaeZoospores(Crustacea) stage following metamorphosis from the nauplius larva,characterised by four pairs of thoracic appendages; may be referred to asprotozoea where differentiation between the nauplius and mysis orpostlarva stage of development is difficultmotile, flagellated and asexual spores32

ABBREVIATIONSBF-2 Bluegill-Fin 2BKDBacterial kidney diseaseBMNBaculoviral Midgut Gland NecrosisBMNVBaculoviral Midgut Gland Necrosis VirusBPBaculovirus penaeiBWSSBacterial white spot syndromeCAIsCowdry type A inclusion bodiesCHSE-214Chinook salmon embryo-214CPECytopathic effectCSHVCoho Salmon HerpesvirusCSTVCoho Salmon Tumour VirusCTABcetyltrimethylammonium bromideDFATDirect fluorescent antibody testDNADeoxyribonucleic aciddddouble distilleddsDNAdouble stranded DNADTABdodecyltrimethylammonium bromideEHNEpizootic Haematopoietic NecrosisEHNVEpizootic Haematopoeitic Necrosis VirusELISAEnzyme-linked Immunosorbent AssayEPCEpithelioma papulosum cyprinaeERAEUS-related AphanomycesEUSEpizootic Ulcerative SyndromeFBSfetal bovine serumFEVFish Encephalitis VirusFHMFathead MinnowGAVGill Associated VirusGPglucose peptoneGPYglucose peptone yeastH&EHaematoxylin & EosinHHNBVBaculoviral Hypodermal and Haematopoietic Necrosis1G4F1% Glutaraldehyde : 4% FormaldehydeICTVInternational Committee on Taxonomy of VirusesIFATIndirect Fluorescent Antibody TestIgGprimary antibody (IgG)IHHNInfectious Hypodermal and Hematopoietic NecrosisIHHNVInfectious Hypodermal and Hematopoietic Necrosis VirusIHNInfectious Haematopoietic NecrosisIHNVInfectious Haematopoietic Necrosis VirusIPNInfectious Pancreatic NecrosisIPNVInfectious Pancreatic Necrosis VirusISHin situ hybridizationkDakilodaltonKDM2Kidney Disease MediumKDMCKidney Disease Medium CharcoalLDVLymphocystis Disease VirusLOS‘lymphoid organ spheroids’LOVLymphoid Organ VirusLOVVLymphoid Organ Vacuolisation VirusLPVLymphoidal Parvo-like VirusMabMonoclonal antibodyMCMSMid-crop Mortality SyndromeMEMMinimal Essential MediumMGMycotic Granuloma-fungus“MSX”multinucleate sphere XNeVTANerka virus Towada Lake, Akita and Amori prefectureNHPNecrotising HepatopancreatitisNPBNuclear Polyhedrosis BaculovirosisOKVOncorhynchus kisutch virus33

ABBREVIATIONSOTCOMVOVVDPCRPBSPKDPLPNHPRDSRHVRKVRNARSDRTG-2RT-PCRRV-PJRVCSDS-PAGESEEDSEMBVSJNNVSKDMSMVSMVDSPFssDNAssRNA“SSO”SSN-1SVCSVCVTEMTNHPTPMSTSTSVUVVERVHSVHSVVIMSVNNYBVYHDYHVYHDBVYHDLVYTVWSBVWSDWSSWSSVoxytetracyclineOncorhynchus masou virusOyster Velar Virus DiseasePolymerase Chain ReactionPhosphate Buffered SalineProliferative Kidney DiseasePostlarvaePeru Necrotizing Hepatopancreatitis“runt deformity syndrome”Rainbow Trout HerpesvirusRainbow Trout Kidney VirusRibonucleic AcidRed spot diseaseRainbow Trout Gonad-2Reverse Transcriptase-Polymerase Chain ReactionRod-shaped Nuclear Virus of Penaeus japonicusRhabdovirus carpioSodium dodecyl sulfate polyacrylamide gel electophoresisShrimp Explosive Epidemic DiseaseSystemic Ectodermal and Mesodermal BaculovirusStriped Jack Nervous Necrosis VirusSelective Kidney Disease MediumSpawner-isolated Mortality VirusSpawner-isolated Mortality Virus DiseaseSpecific Pathogen Freesingle stranded DNAsingle stranded RNAseaside organismStriped Snakehead (Channa striatus) cell-lineSpring Viraemia of CarpSpring Viraemia of Carp VirusTransmission Electron MicroscopyTexas Necrotizing HepatopancreatitisTexas Pond Mortality SyndromeTaura SyndromeTaura Syndrome VirusultravioletViral Encephalopathy and RetinopathyViral Haemorrhagic SepticaemiaViral Haemorrhagic Septicaemia VirusVirginia Institute of Marine ScienceViral Nervous Necrosis)Yellowhead BaculovirusYellowhead diseaseYellowhead VirusYellowhead Disease BaculovirusYellow-Head-Disease-Like virusYamame tumour virusWhite Spot BaculovirusWhite Spot DiseaseWhite Spot SyndromeWhite Spot Syndrome Virus34

SCIENTIFIC AND COMMON NAMESA. FINFISH (Hosts)Scientific NameArgentina sphyraenaAristichthys nobilisBidyanus bidyanusCarassius auratusCarassius carassiusChanna striatusChanos chanosClupea harengusClupea pallasiCoregonus spp.Ctenopharyngodon idellusCyprinus carpioDicentrarchus labraxEpinepheles akaaraEpinephelus malabaricusEpinephelus moaraEsox luciusGadus macrocephalusGadus morhuaGalaxias olidusGambussia affinisHippoglossus hippoglossusHypophthalmichthys molitrixIctalurus melasLabroides dimidatusLates calcariferMacquaria australasicaMelanogrammus aeglefinusMerlangius merlangiusMicromesistius poutassouMugil cephalusOncorhynchus ketaOncorhynchus kisutchOncorhynchus masouOncorhynchus mykissOncorhynchus nerkaOncorhynchus rhodurusOncorhynchus tshawytschaOplegnathus fasciatusOplegnathus punctatusOreochromis spp.Paralichthys olivaceusPerca fluviatilisPlecoglossus altivelisPoecilia reticulataPseudocaranx dentexRhinonemus cimbriusSalmo salarSalmo truttaSalvelinus fontinalisScophthalmus maximusSeriola quinqueradiataSilurus glanisSparus aurataCommon Namelesser argentinebighead carpsilver perchgoldfishcrucian carpstriped snakeheadmilkfishherringPacific herringwhite fishgrass carpcommon carpEuropean sea bassred-spotted grouperbrown spotted grouperkelp grouperpikePacific codAtlantic codmountain galaxiasmosquito fishhalibutsilver carpcatfishdoctor fishsea bass, Australian barramundiMacquarie perchhaddockwhitingblue whitinggrey mulletchum salmoncoho salmonsockeye salmon/Yamame salmon/masousalmonrainbow or steelhead troutKokanee (non-anadromous sockeye) salmonamago salmonchinook salmonJapanese parrotfishrock porgyTilapiaJapanese flounderredfin perchayuguppystriped jackrocklingAtlantic salmonbrown troutbrook troutturbotJapanese yellowtail floundersheatfishgilt-head sea bream35

SCIENTIFIC AND COMMON NAMESSprattus sprattusTakifugu rubripesTinca tincaThymallus thymallusTrisopterus esmarkiiUmbrina cirrosasprattiger puffertenchgraylingNorway poutshi drumB. MOLLUSCS (Hosts)Scientific NameAcanthogobius flavimanusArca sp.Argopecten gibbusAustrovenus stutchburyiBarbatia novae-zelandiae (Family Arcidae)Cerastoderma (= Cardium) eduleCrassostrea angulataCrassostrea ariakensisCrassostrea commercialisCrassostrea gigasCrassostrea virginicaCrassostrea angulataHaliotis cyclobatesHaliotis laevigataHaliotis roeiHaliotis rubraHaliotis scalarisMacomona liliana (Family Tellinidae)Mercenaria mercenariaMytilus edulisMytilus galloprovincialisOstrea angasiOstrea conchaphila (O. lurida)Ostrea edulisOstrea lutaria (Tiostrea lutaria)Ostrea puelchanaPatinopecten yessoensisPinctada albicansPinctada maximaPteria penguinRuditapes decussatusRuditapes philippinarumSaccostrea commercialisSaccostrea (Crassostrea) cucullataSaccostrea echinataSaccostrea glomerataScrobicularia planaTiostrea chilensis (Ostrea chilensis)Tiostrea lutariaTridacna maximaCommon NameJapanese yellow gobyclamsCalico scallopNew Zealand cockles(not available)Common European cocklePortuguese oystersAriake cupped oysterSydney rock oysterPacific oysterAmerican oystersPortugese oystersabalonegreenlip abaloneabaloneblacklip abaloneabalone(bivalve, not available)hard shell clamedible musseledible musselflat oyster (southern mud oyster)Olympia oysterEuropean oysterNew Zealand oyster(not available)Japanese (Yesso) scallopspearl oysterMother of pearlwinged pearl oysterEuropean clamManila clamSydney rock oysterMangrove oysterNorthern black lip oysterSydney rock oystersPeppery furrow shellSouth American oyster(not available)giant clamC. CRUSTACEANS (Hosts)Scientific NameAcetes spp. (Crustacea:Sergestidae)Cherax quadricarinatusEuphausia spp.Common Namekrill, small shrimpfreshwater crayfish, red clawkrill36

SCIENTIFIC AND COMMON NAMESMarsupenaeus (Penaeus) japonicusMetapenaeus ensisPalaemon styliferusPenaeus aztecusPenaeus californiensisPenaeus chinensisPenaeus duodarumPenaeus esculentusPenaeus indicusPenaeus japonicusPenaeus marginatusPenaeus merguiensisPenaeus monodonPenaeus occidentalisPenaeus paulensisPenaeus penicillatusPenaeus plebejusPenaeus schmittiPenaeus semisulcatusPenaeus setiferusPenaeus stylirostrisPenaeus subtilisPenaeus vannameiKuruma prawnred endeavour or greasy back shrimp/prawn(not available)Northern brown shrimpyellowleg shrimpChinese white shrimpcaged pink shrimpbrown tiger shrimp/prawnIndian or red legged banana shrimp/prawnJapanese king or Kuruma shrimp/prawnAloha prawncommon or Gulf banana shrimp/prawngiant black tiger shrimp/prawnWestern white shrimppink shrimpredtail prawn, beige colored shrimpEastern king shrimp/prawnwhite shrimpgrooved tiger or green tiger shrimp/prawnNative white shrimpblue shrimpSouthern brown shrimpwhite shrimpD. Pathogens/Disease AgentsAeromonas hydrophilaArgulus foliaceusArgulus sppAphanomyces astaciAphanomyces invadansAphanomyces invaderisAphanomyces piscicidaBaculovirus penaeiBonamia ostreaeBonamia spDermocystidium marinumHaplosporidium costaleHaplosporidium. NelsoniHerpervirusHexamita inflataHexamita salmonisMytilicola sp.Labyrinthomyxa marinusLerneae cyprinaceaMarteilia mauriniMarteilia refringensMarteilia sydneyiMarteilioides branchialisMarteilioides christenseniMarteilioides chungmuensisMarteilioides lengehiMikrocytos mackiniMikrocytos roughleyiMinchinia costaleMinchinia nelsoniMyxobolus artusLigula sp.Perkinsus atlanticus37

SCIENTIFIC AND COMMON NAMESPerkinsus marinusPerkinsus olseniPerkinsus qugwadiPiscicola geometraPolydora sp.Posthodiplostomum cuticolaRanavirusRenibacterium salmoninarumRhabdovirus carpioSalmincola salmoneusStaphylococcus aureusVibrio harveyiVibrio splendidusVibrio spp38

SECTION 1- INTRODUCTIONI. INTRODUCTIONI.1 BackgroundI.2 Objectives and ScopeI.3 Guide for UsersI.4 Health and Aquatic AnimalsI.5 Role of Diagnostics in Aquatic Animal HealthI.6 Levels of DiagnosticsI.7 References3940404042434346I. INTRODUCTIONI.1 BackgroundThe FAO Regional Technical CooperationProgramme (TCP) Project “Assistance for ResponsibleMovement of Live Aquatic Animals”(TCP/RAS/6714-A and 9605-A), wasimplemented in January 1998 by NACA , in cooperationwith the OIE 1 , regional and internationalagencies (e.g. AAHRI 2 , AusAID/APEC 3 ,AFFA 4 , and others), representatives (designatedNational Coordinators and focal points for diseasereporting) of 21 governments/territoriesin the Asia-Pacific region (Australia,Bangladesh, Cambodia, China PR, Hong KongSAR China, India, Indonesia, Iran, Japan, Korea(DPR), Korea (RO), Lao PDR, Malaysia,Myanmar, Nepal, Pakistan, Philippines,Singapore, Sri Lanka, Thailand and Vietnam)and many regional and international aquaticanimal disease experts. The over-all objectiveof the program was to provide guidance tocountries in undertaking responsible movement(introductions and transfers) of live aquatic animalsthrough appropriate strategies that minimizepotential health risks associated with liveaquatic animal movements. The program tookinto account the need for concordance with existinginternational agreements/treaties (e.g.WTO’s SPS Agreement and OIE health standards)along with the need for the strategies tobe practically applicable to the Asia region andin support for FAO’s Code of Conduct for ResponsibleFisheries (CCRF). This TCP becamethe focal point for the development of a strong,multidisciplinary Asia-Pacific RegionalProgramme on Aquatic Animal Health Managementwhich is a major element of NACA’s FiveYear Work Programme (2001-2005). The “AsiaRegional Technical Guidelines on HealthManagement for the Responsible Movementof Live Aquatic Animals and the Beijing Consensusand Implementation Strategy(TGBCIS)” or ‘Technical Guidelines’ (FAO/NACA 2000) and the corresponding “Manualof Procedures (MOP)” (FAO/NACA 2001) weredeveloped over a period of three years (from1998-2001) of awareness and consensus buildingin consultation (through various nationallevel and regional workshops, FAO/NACA/OIE1998) with government representatives, representativesof collaborating organizations andaquatic animal health experts. The ‘TechnicalGuidelines’ was finally adopted in principle duringa Final Workshop of the TCP held in Beijing,China PR in June 2000 (FAO/NACA 2000). TheAsia Diagnostic Guide to Aquatic Animal Diseasesor ‘Asia Diagnostic Guide’ is a third ofa series of documents produced under the TCPthat will support the implementation of the‘Technical Guidelines’ particularly with respectto the component on disease diagnosis, surveillanceand reporting.The ‘Asia Diagnostic Guide’ is a comprehensivediagnostic manual for the pathogens anddiseases listed in the NACA/FAO/OIE QuarterlyAquatic Animal Disease Reporting System 5 . Itwas developed from technical contributionsfrom members of the Regional Working Group(RWG) and Technical Support Services (TSS)of the TCP and other aquatic animal health scientistsin the Asia-Pacific region and outsidewho supported the regional programme.Many useful aquatic animal health diagnosticguides and manuals and others in CD-ROM formatalready exist in the literature. Some are in1Office International des Epizooties2Aquatic Animal Health Research Institute of the Thai Department of Fisheries3Australian Agency for International Development/Asia-Pacific Economic Cooperation4Agriculture, Fisheries and Forestry of Australia5The quarterly reporting system was developed as one of the four major components of the TCP, developed based on the OIEInternational Aquatic Animal Health Code – 1997, in cooperation with the OIE Regional Representation for Asia and the Pacific.39

INTRODUCTIONthe language of individual countries. In the Asia-Pacific region, more recent ones includeIndonesia’s Manual for Fish Disease Diagnosis- II (Koesharyani et al. 2001, GRIM 6 /JICA 7 ); thePhilippines’ Diseases of Penaeid Shrimps in thePhilippines (Lavilla-Pitogo et al. 2000,SEAFDEC-AQD 8 ); Thailand’s (a) Diagnostic Proceduresfor Finfish Diseases (Tonguthai et al.1999, AAHRI), (b) Health Management in ShrimpPonds, Third Edition (Chanratchakool et al.1998, AAHRI), and (c) Epizootic Ulcerative Syndrome(EUS) Technical Handbook (Lilley et al.1998, ACIAR 9 /DFID 10 /AAHRI/NSW 11 -Fisheries/NACA);Australia’s Australian Aquatic AnimalDisease – Identification Field Guide (Herfortand Rawlin 1999, AFFA); and a CD-ROM onDiagnosis of Shrimp Diseases (Alday deGraindorge and Flegel 1999). Some more arelisted and appear as an Annex in the differentsections of the Asia Diagnostic Guide.The ‘Asia Diagnostic Guide’ supplementsthese existing manuals/guides and providesrelevant information on diseases in the NACA/FAO and OIE Asia-Pacific Quarterly AquaticAnimal Disease Reporting System, whichcommenced during 3 rd quarter of 1998 (NACA/FAO 1999, OIE 1999). The information in theAsia Diagnostic Guide is presented in a formatthat spans from gross observations at the pondor farm site (Level 1), to guidance for informationon technologically advanced molecular orultrastructural diagnostics and laboratory analyses(Levels II and III, and OIE 2000a, b), thus,taking into account international, regional, andnational variations in disease concerns, as wellas varying levels of diagnostic capability betweencountries of the Asia-Pacific region.I.2 Objectives and ScopeThe objective of the “Asia Diagnostic Guide”is to produce a manual/guide of specific usefor both farm and laboratory level aquatic animaldisease diagnostics in the Asia region thatcomplements the ‘Manual of Procedures’ andthat which will serve as a supplement to theimplementation of the ‘Technical Guidelines’.The Asia Diagnostic Guide is aimed at providinga tool that can be used to expand nationaland regional aquatic animal diagnostic capacitiesand the infrastructure required to meet theOIE aquatic animal health standards (OIE2000a, b). This guide aims to improve aquaticanimal health awareness as well as provideknowledge on how to access the diagnosticresources required to help prevent or controldisease impacts.The Asia Diagnostic Guide focuses on theNACA/FAO and OIE listed diseases, but alsoincludes some which are significant in parts ofthe Asia-Pacific region.I.3 Guide for UsersThe Asia Diagnostic Guide is divided into foursections: Section 1 on Introduction, Background,Scope and Purpose, Guide for Users,Health and Aquatic Animals, Role of Diagnosticsand Levels of Diagnostics; Sections 2 to4, divided into host groups, i.e. Finfish Diseases(Section 2), Molluscan Diseases (Section3) and Crustacean Diseases (Section 4),each commences with a chapter on “GeneralTechniques” which covers the essential “startingpoints” that will enable prompt and effectiveresponse(s) to disease situations in aquaticanimal production. This chapter is not diseasespecific,providing information applicable to awide range of both infectious and noninfectiousdisease situations. It emphasizes the importanceof gross observations (Level 1), and howand when they should be made. It also describesenvironmental parameters worth recording,general procedures for sampling andfixation and the importance of record-keeping.Each General Techniques section is dividedas follows:Gross ObservationsBehaviourSurface ObservationsEnvironmental ParametersGeneral ProceduresPre-collection PreparationBackground InformationSample Collection for Health ScreeningSample Collection for Disease DiagnosisLive Specimen Collection and ShippingDead or Tissue Specimen Collection andShippingPreservation of TissuesShipping Preserved Specimens6Gondol Research Institute for Mariculture of the Central Research Institute for Sea Exploration and Fisheries, Indonesia’sDepartment of Marine Affairs and Fisheries7Japan International Cooperation Agency8Aquaculture Department of the Southeast Asian Fisheries Development Center9Australian Centre for International Agricultural Research10Department for International Development of the United Kingdom11New South Wales (Australia)40

INTRODUCTIONTable I.2.1. NACA/FAO and OIE listed diseases and other diseases 12 covered in the Asia DiagnosticGuide.DISEASES PREVALENT IN SOME PARTS OF THE REGIONFinfish diseases1. Epizootic haematopoietic necrosis* (EHN)2. Infectious haematopoietic necrosis* (IHN)3. Oncorhynchus masou virus disease* (OMVD)4. Infectious pancreatic necrosis (IPN)5. Viral encephalopathy and retinopathy (VER)6. Epizootic ulcerative syndrome (EUS)7. Bacterial kidney disease (BKD)Mollusc diseases1. Bonamiosis * (Bonamia sp., B. ostreae)2. Marteiliosis * (Marteilia refringens, M. sydneyi)3. Microcytosis * (Mikrocytos mackini, M. roughleyi)4. Perkinsosis * (Perkinsus marinus, P. olseni)Crustacean diseases1. Yellowhead disease (YHD)2. Infectious hypodermal and haematopoietic necrosis (IHHN)3. White spot disease (WSD)4. Baculoviral midgut gland necrosis (BMN)5. Gill associated virus (GAV)6. Spawner mortality syndrome (‘Midcrop mortality syndrome’) (SMVD)DISEASES PRESUMED EXOTIC TO THE REGION, BUT REPORTABLE TO THE OIEFinfish diseases1. Spring viraemia of carp* (SVC)2. Viral haemorrhagic septicaemia* (VHS)Mollusc diseases1. Haplosporidiosis* (Haplosporidium costale, H. nelsoni)OTHER DISEASES WITHIN THE REGION, NOT CURRENTLY LISTEDFinfish diseases1. LymphocystisMollusc diseases1. Marteilioidosis (Marteilioides chungmuensis, M. branchialis)2. Iridovirosis (Oyster velar virus disease)Crustacean diseases (the following diseases are so far presumed, but not proven,to be exotic to this region)1. Taura Syndrome (TS)2. Nuclear Polyhedrosis Baculovirosis (Baculovirus penaei)3. Necrotising hepatopancreatitis4. Crayfish plague*OIE Notifiable Diseases (OIE 1997)12The diseases listed in the Asia-Pacific Quarterly Aquatic Animal Disease Reporting System were agreed through a process ofconsultation with the National Coordinators, members of the Regional Working Group (RWG) and Technical Support Services(TSS), FAO, NACA and OIE based on the OIE International Aquatic Animal Health Code – 1997, including some diseases whichare deemed important to the Asia-Pacific region.41

INTRODUCTIONRecord-keepingGross ObservationsEnvironmental ObservationsStocking RecordsReferencesThe “General Techniques” section is followed,under each host group, by a number of chaptersaimed at specific diseases (e.g. viral, bacterial,fungal) listed on the current NACA/FAOand OIE quarterly reporting list (Table 1.2.1).These are recognised of being of regional importance,as well as of international trade significance.Those diseases listed as “Notifiable”or “Other Significant Diseases” by the OIE arecross-referenced to the most up-to-date versionof the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000b, also available athttp://www.oie.int). The diagnostic techniquesdescribed in the Asia Diagnostic Guide are consistentwith those recommended by OIE. Sinceit is recognised that disease diagnostics is adynamic field, it is highly recommended thatanyone using this manual for the purpose ofhealth certification for international transfersof live aquatic organisms refer to the OIEdiagnostic manual prior to performing diagnostics(screening) for this purpose. In addition,other diseases/infectious agents not presentlyincluded on the Regional Quarterly ReportingList are included in the Asia DiagnosticGuide, since they are of interest to the regionand infect commercially significant species.Each chapter on specific diseases is presentedwith information on the following:• Causative Agent(s)- an introductory paragraphon the causative agent(s) responsiblefor the disease.• Host Range – the range of hosts that can beinfected (both naturally and experimentally).• Geographic Distribution - known/recordedgeographic range of the disease (updated,where applicable, using the Asia-PacificQuarterly Aquatic Animal Disease Reports for1999-2000 (OIE, NACA/FAO quarterly reports).• Clinical Aspects - the effects of the diseaseare described, ranging from gross observationsand behavioural changes, lesions andother external clues, to gross and microscopicinternal pathology.• Screening Methods – are the examinationmethods applied to check healthy appearingaquatic animals to determine whether ornot they are infected by a potentially significantinfectious agent.• Diagnostic Methods – are the examinationprocedures used to try and determine thecause of a disease or clinical infection.Screening and diagnostic methods aredivided into two types:Presumptive Diagnosis – preliminary diagnosisbased on gross observations and circumstantialevidence. Where more than one infectiousagent may be responsible, confirmatorydiagnosis (usually laboratory Level II and/or III)may be required; andConfirmatory Diagnosis – positive identificationof the causative agent, with a high degreeof diagnostic confidence.• Modes of Transmission - deals with theknown modes of transmission (spread) of thedisease and the factors associated with itsspread (environmental, handling, life-historystage, reservoirs of infection, etc.). This areaof diagnostics is known as epidemiology (orepizootiology) and available observationsfrom this field of study are also included.• Control Measures – describes any controlmeasures which are known to work, shouldthe disease appear.• References – recent relevant publicationsabout the disease.The chapters for each host group also includethree Annexes providing information of the (a)list of OIE Reference Laboratories, (b) a list ofregional disease experts who can provide informationand valuable health advise, and (c)useful guides/manuals. A Glossary is also included.I.4 Health and Aquatic AnimalsUnlike other farm and harvesting situations,where the animals and plants are visible, aquaticanimals require more attention in order to monitortheir health. They are not readily visible, exceptunder tank-holding conditions, and theylive in a complex and dynamic environment.Likewise, feed consumption and mortalitiesmay be equally well hidden under water. Unlikethe livestock sector, aquaculture has a widerange of diversity in species cultured, farmingenvironment, nature of containment, intensityof practice and culture system used. The rangeof diseases found in aquaculture is also varied,some with low or unknown host specificitiesand many with non-specific symptoms. Diseaseis now recognized as one of the most importantchallenges facing the aquaculture sector.The complexity of the aquatic ecosystemmakes the distinction between health, sub-optimalperformance and disease obscure. Diseasesin aquaculture are not caused by a single42

INTRODUCTIONevent but are the end result of a series of linkedevents involving the interactions between thehost (including physiological, reproductive anddevelopmental stage conditions), the environmentand the presence of a pathogen (Snieszko1974). Under aquaculture conditions, three factorsare particularly important affecting host’ssusceptibility: stocking density, innate susceptibilityand immunity (natural/acquired). Environmentincludes not only the water and its components(such as oxygen, pH, temperature, toxins,wastes) but also the kind of managementpractices (e.g. handling, drug treatments, transportprocedures, etc.). Pathogens may includeviruses, bacteria, parasites and fungi; diseasesmay be caused by a single species or a mixtureof different pathogens. The introduction ofinfectious diseases is another major concernin aquaculture. As in livestock, the aquacultureand fisheries sector will continue to face increasingglobal exposure to disease agents asit intensifies trade in live aquatic organisms andtheir products (Subasinghe et al. 2001).The first and most important defenses againstpreventable disease losses under such complexsituations are:monitoring as regularly as possible andappropriate action at the first sign(s) of suspiciousbehaviour, lesions or mortalities.These fundamental approaches – despite havingbeen long instilled in human and agriculturalproduction – still require reinforcement inmany aquatic animal production sectors. Somefarmers and harvesters still hesitate to act atthe first sign of health problems, due to concernthat it may reflect on their production capability,or that it will result in failure in the competitivemarket place. Hiding or denying healthproblems, however, can be as destructive toaquatic animals as it is elsewhere. It is importantto recognise that disease is a challengethat everyone has to face, and having the resourcesthat can effectively deal with it, are theprimary weapons against misplaced ignoranceand fear.I.5 Role of Diagnostics in Aquatic AnimalHealth Management and DiseaseControlDiagnostics play two significant roles in aquaticanimal health management and disease control.As described above, some diagnostic techniquesare used to screen healthy animals toensure that they are not carrying infection atsub-clinical levels by specific pathogens. Thisis most commonly conducted on stocks orpopulations of aquatic animals destined for livetransfer from one area or country to another.Such screening provides protection on twofronts: (a) it reduces the risk that animals arecarrying few, if any, opportunistic agents whichmight proliferate during shipping, handling orchange of environment; and (b) it reduces therisk of resistant or tolerant animals transferringa significant pathogen to a population whichmay be susceptible to infection. The secondrole of diagnostics is to determine the causeof unfavourable health or other abnormality(such as spawning failure, growth or behaviour)in order to recommend mitigating measures applicableto the particular condition. This is themost immediate, and clearly recognised, roleof diagnostics in aquatic animal health.Accurate diagnosis of a disease is often incorrectlydescribed as complicated and costly. Thismay be the case for some of the more difficultto diagnose diseases or newly emerging diseases.Disease diagnosis is not solely a laboratorytest. A laboratory test may confirm thepresence of a specific disease agent, or it mayexclude its presence with a certain level of certainty.Incorrect diagnosis can lead to ineffectiveor inappropriate control measures (whichmay be even more costly). For example, a “new”disease agent may get introduced to a majoraquaculture producing area, or the animals mayall die in shipment/during handling. Disease diagnosticsshould be made as a continuum ofobservations starting on the farm and, in fact,commencing prior to the disease event. Thedifferent levels of disease diagnostics which canbe undertaken when investigating a diseasesituation are discussed in the section below.I.6 Levels of DiagnosticsThe Asia Diagnostic Guide is built on a frameworkof “three levels” of diagnostics, agreedupon during the Second Regional Workshop ofthe TCP held in Bangkok in February 1999 (seeFAO/NACA 2000). Table I.6.1 below outlines thediagnostic activities at each level, who is responsible,and the equipment and training required.It should be noted that none of the levelsfunction in isolation, but build on each other,each contributing valuable data and informationfor optimum diagnoses. Level 1 providesthe foundation and is the basis of Levels II andIII since findings using higher level(s) can onlybe meaningfully interpreted only in conjunctionwith observations and results obtained fromlower levels.43

INTRODUCTIONLevel I (farm/production site observations,record-keeping and health management) isstrongly emphasized throughout the Asia DiagnosticGuide as this forms the basis for triggeringthe other diagnostic levels (II and III).Level II includes the specialisations of parasitology,histopathology, bacteriology and mycology,which require moderate capital and traininginvestment, and which, generally-speaking,cannot be conducted at the farm or culture site.Level III comprises the types of advanced diagnosticspecialisation which requires significantcapital and training investment. As thereader will note, immunology and biomoleculartechniques are included in Level III, althoughfield kits are now being developed for farm orpond-side use (Level I) as well as use in microbiologyor histology laboratories (Level II). Theseefforts are good indication that technologytransfer is now enhancing diagnostics and, withsolid quality control and field validation, it iscertain that more Level III technology will becomefield accessible in the near future (Walkerand Subasinghe 2000).One of the most important aspects ofmaximising the effectiveness of the three diagnosticlevels is ensuring that Level I diagnosticianshave access to, and know how to contactLevels II and III support (and at what cost),and vice versa. Level III diagnostic support isusually based on referrals, so has little contactwith field growing conditions. They, therefore,need feedback to ensure any diagnosis (andactions recommended) are relevant to aquaticanimal production situation being investigated.Thus, the baseline aim for initiating diagnosticcapability is Level I. Confirmatory diagnostics,or second opinion, where required, can be obtainedby referral until such capabilities are developedlocally. The period required to developLevel II and/or Level III diagnostic infrastructures,usually depends on the disease situationsbeing faced and tackled by Level I diagnosticiansin the area/country and the resourcesavailable. Where there are few problems, thereis little incentive to build diagnostic capability.This is a vulnerable position, and strong linkswith Level II and/or III diagnostics are good precautionarymeasures and strongly promotedunder the regional program – especially for introductionsof live aquatic animals into a relativelydisease-free area.44

IINTRODUCTIONTechnical requirements to support activitiesField keys.Farm record keeping formats.Eqiopment listsModel clinical observation sheets.Pond/Site record sheets.Preservation/transportation guidelines for Levels II/IIIdiagnoses.Model job descriptions/skill requirements.Asia Diagnostic Guide for Aquatic Animal DiseasesModel laboratory record-keeping systemProtocols for preservation/ transport of samples to Level IIIModel laboratory requirements/ equipment/ consumables listsModel job descriptions/ skill listsAccess to Level II and Level III specialist expertiseAsia Diagnostic Guide for Aquatic Animal DiseasesOIE Diagnostic Manual for Aquatic Animal DiseasesRegional General Diagnostics ManualsModel laboratory requirements/ equipment/ consumables listsModel job descriptions/ skill requirementsContact information for reference laboratoriesProtocols for preservation of samples for consultation/ validationAsia Diagnostic Guide for Aquatic Animal DiseasesOIE Diagnostic Manual for Aquatic Animal DiseasesGeneral molecular and microbiology diagnostic referencesTable I.6.1 Diagnostic Levels, Associated Requirements and Responsibilities.LevelActivityObservation of animaland environmentGross clinicalexaminationWork requirementsKnowledge of normal (feeding,behaviour, growth) of stock.Frequent / regular observation of stock.Regular, consistent record-keeping andassistance (Levels II, III).maintenance of records – includingfundamental environmental information.ResponsibilityFarm worker/manager.Fishery extensionofficers.On-site veterinarysupport.Local fishery biologists.Knowledge contacts for health diagnosisAbility to submit and/or preserverepresentative specimens for optimaldiagnosis (Levels II, III).IIParasitologyBacteriologyLaboratories with basic equipment andpersonnel trained/experienced in aquaticanimal pathology.Fish biologists/ technicians.Aquatic Veterinarians.MycologyHistopathologyKeep and maintain accurate diagnostic andlaboratory case records.Ability to preserve and storage specimens foroptimal Level III diagnoses.Parasitologists/ technicians.Mycologists/ techniciansBacteriologists/ technicians.Knowledge of/ contact with different areas ofspecialisation within Level II.Histopathologists/ technicians.Knowledge of who to contact for Level IIIdiagnostic assistance.IIIVirologyElectron microscopyMolecular biologyImmunologyHighly equipped laboratory with highlyspecialised and trained personnel.Keep and maintain accurate diagnosticand laboratory case records.Preserve and store specimens.Virologist/ technician.Ultrastructural histopathologist/technicians.Molecular biology scientists/technicians.Maintenance of contact with peopleresponsible for sample submission.45

INTRODUCTIONI.7 ReferencesAlday de Graindorge, V. and T.W. Flegel. 1999.Diagnosis of shrimp diseases with emphasison blacktiger prawn, Penaeus monodon.Food and Agriculture Organization of theUnited Nations (FAO),Multimedia Asia Co.,Ltd, BIOTEC, Network of Aquaculture Centresin Asia Pacific (NACA)and SoutheastAsian Chapter of the World Aquaculture Society(WAS). Bangkok, Thailand. (Inter-activeCD-ROM format).Chanratchakool, P., J.F. Turnbull, S.J. Funge-Smith, I.H. MacRae and C. Limsuan. 1998.Health management in shrimp ponds. ThirdEdition. Aquatic Animal Health Research Institute(AAHRI), Bangkok, Thailand. 152p.FAO/NACA. 2000. Asia Regional TechnicalGuidelines on Health Management for theResponsible Movement of Live Aquatic Animalsand the Beijing Consensus and ImplementationStrategy. FAO Fisheries TechnicalPaper. No. 402. Rome, FAO. 2000. 53p.FAO/NACA. 2001. Manual of Procedures for theImplementation of the Asia Regional TechnicalGuidelines on Health Management for theResponsible Movement of Live Aquatic Animals.FAO Fisheries Technical Paper. No.402,Suppl. 1. Rome, FAO. 2001. 106p.FAO/NACA/OIE. 1998. Report of the First TrainingWorkshop of the Regional Programme forthe Development of Technical Guidelines onQuarantine and Health Certification and Establishmentof Informations Systems, for theResponsible Movement of Live Aquatic Animalsin Asia. Bangkok, Thailand, 16-20 January1998. TCP/RAS/6714 Field DocumentNo. 1. Food and Agriculture Organization ofthe United Nations (FAO), Network of AquacultureCentres in Asia Pacific (NACA) and theOffice International des Epizooties (OIE).Bangkok, Thailand. 142p.Herfort, A. and G.T. Rawlin. 1999. AustralianAquatic Animal Disease Identification FieldGuide. Agriculture, Fisheries and Forestry –Australia (AFFA), Canberra. 90p.Koesharyani, I., D. Roza, K. Mahardika, F.Johnny, Zafran and K. Yuasa. 2001. Manualfor Fish Disease Diagnosis – II. Marfine Fishand Crustacean Diseases in Indonesia.Gondol Research Institute for Mariculture andJapan International Cooperation Agency.Bali, Indonesia. 49p.Lavilla-Pitogo, C.R. G.D. Lio-Po, E.R Cruz-Lacierda, E.V. Alapide-Tendencia and L.D. dela Pena. 2000. Diseases of penaeid shrimpsin the Philippines. Aquaculture ExtensionManual No. 16. Second edition. AquacultureDepartment of the Southeast Asian FisheriesDevelopment Center (SEAFDEC-AQD),Tigbauan, Iloilo, Philippines. 83p.Lilley, J.H., R.B. Callinan, S. Chinabut, S.Kanchanakhan, I.H. MacRae and M.J.Phillips. 1998. Epizootic Ulcerative Syndrome(EUS) Technical Handbook. AquaticAnimal Health Research Institute (AAHRI),Bangkok, Thailand. 88p.NACA/FAO. 1999. Quarterly Aquatic AnimalDisease Report (Asia and Pacific Region),July-September 1998. FAO Project TCP/RAS/6714. Network of Aquaculture Centresin Asia Pacific (NACA), Bangkok, Thailand.37p.OIE. 1997. OIE International Aquatic AnimalHealth Code, Second Edition. 1997. OfficeInternational des Epizooties (OIE), Paris,France. 192p.OIE. 1998. Quarterly Aquatic Animal DiseaseReport, October-December 1998 (Asian andPacific Region). 1998(2). The OIE Representationfor Asia and the Pacific, Tokyo, Japan.33p.OIE. 2000a. OIE International Aquatic AnimalHealth Code, Third Edition, 2000. Office Internationaldes Epizooties (OIE), Paris,France. 153p.OIE. 2000b. OIE Diagnostic Manual for AquaticAnimal Diseases, Third Edition, 2000. OfficeInternational des Epizooties (OIE), Paris,France. 237p.Snieszko, S.F. 1974. The effects of environmentalstress on outbreaks of infectious diseasesof fishes. J. Fish. Biol. 6:197-208.Subasinghe, R.P., M.G. Bondad-Reantaso andS.E. McGladdery. 2001. Aquaculture development,health and wealth. In: Subasinghe,R.P., P. Bueno, M.J. Phillips, C. Hough, S.E.McGladdery and J.R. Arthur (eds.). Aquaculturein the Third Millennium. Technical Proceedingsof the Conference on Aquaculturein the Third Millennium, Bangkok, Thailand,20-25 February 2000. (in press).46

INTRODUCTIONTonguthai, K. S. Chinabut, T. Somsiri, P.Chanratchakool and S. Kanchanakhan.1999. Diagnostic Procedures for Finfish Diseases.Aquatic Animal Health Research Institute,Bangkok, Thailand.Walker, P. and R.P. Subasinghe (eds.) 2000.DNA-based molecular diagnostic techniques:research needs for standardization and validationof the detection of aquatic animalpathogens and diseases. Report and proceedingsof the Expert Workshop on DNAbasedMolecular Diagnostic Techniques: ResearchNeeds for Standardization and Validationof the Detection of Aquatic AnimalPathogens and Diseases. Bangkok, Thailand,7-9 February 1999. FAO Fisheries TechnicalPaper. No. 395. Rome, FAO. 2000. 93p.47

Basic Anatomy of a Typical Bony Fishspinal cordswim bladderbrainheartliverpelvic findorsal finstomachkidney caudal peduncleBasic anatomy of a typical bony fish.intestineanal finspleencaudal fin48

SECTION 2 - FINFISH DISEASESBasic Anatomy of a Typical Bony FishSECTION 2 - FINFISH DISEASESF.1 GENERAL TECHNIQUESF.1.1F.1.1.1F.1.1.2F.1.1.2.1F.1.1.2.2F.1.1.2.3F.1.1.3F.1.1.3.1F.1.1.3.2F.1.2F.1.3F.1.3.1F.1.3.2F.1.3.3F.1.3.4F.1.3.5F.1.3.6F.1.3.7F.1.3.8F.1.4F.1.4.1F.1.4.2F.1.4.3F.1.5Gross ObservationsBehaviourSurface ObservationsSkin and FinsGillsBodyInternal ObservationsBody Cavity and MuscleOrgansEnvironmental ParametersGeneral ProceduresPre-Collection PreparationBackground InformationSample Collection for Health SurveillanceSample Collection for Disease DiagnosisLive Specimen Collection for ShippingDead or Tissue Specimen Collection for ShippingPreservation of Tissue SamplesShipping Preserved SamplesRecord-KeepingGross ObservationsEnvironmental ObservationsStocking RecordsReferencesVIRAL DISEASES OF FINFISHF.2 Epizootic Haematopoietic Necrosis (EHN)F.3 Infectious Haematopoietic Necrosis (IHN)F.4 Oncorhynchus masou Virus (OMV)F.5 Infectious Pancreatic Necrosis (IPN)F.6 Viral Encephalopathy and Retinopathy (VER)F.7 Spring Viraemia of Carp (SVC)F.8 Viral Haemorrhagic Septicaemia (VHS)F.9 LymphocystisBACTERIAL DISEASE OF FINFISHF.10 Bacterial Kidney Disease (BKD)FUNGUS ASSOCIATED DISEASE FINFISHF.11 Epizootic Ulcerative Syndrome (EUS)485050505050515252525253535354545455555656575757575759626568727679828690F.AIF.AIIF.AIIIANNEXESOIE Reference Laboratories for Finfish DiseasesList of Regional Resource Experts for FinfishDiseases in Asia-PacificList of Useful Diagnostic Manuals/Guides toFinfish Diseases in Asia-Pacific959810549

F.1 GENERAL TECHNIQUESGeneral fish health advice and other valuableinformation are available from the OIE ReferenceLaboratories, Regional Resource Expertsin the Asia-Pacific, FAO and NACA. Alist is provided in Annexes F.AI and AII, andup-to-date contact information may be obtainedfrom the NACA Secretariat in Bangkok(e-mail: naca@enaca.org). Other useful guidesto diagnostic procedures which provide valuablereferences for regional parasites, pestsand diseases are listed in Annex F.AIII.F.1.1 Gross ObservationsF.1.1.1 Behaviour (Level I)At a time when there are no problems on the farm,“normal behaviour” of the animals should be observedto establish and describe the “normal”situation. Any change from normal behaviourshould be a cause for concern and warrants investigation.Prior to the clinical expression of diseasesigns, individual finfish may exhibit increasedfeed consumption followed by cessationof feeding, or the fish may simply go off feedalone. Taking note of normal feed conversionratios, length/weight ratios or other body-shapesigns described below, is essential in order todetect impending disease.Abnormal behaviour includes fish swimming nearthe surface, sinking to the bottom, loss of balance,flashing, cork-screwing or air gulping (nonair-breathers) or any sign which deviates fromnormal behaviour. Bursts of abnormal activity areoften associated with a generalised lethargy.Behavioural changes often occur when a fish isunder stress. Oxygen deprivation leads to gulping,listlessness, belly-up or rolling motion. Thiscan be due to blood or gill impairment. Flashingcan indicate surface irritation, e.g., superficialsecondary infections of surface lesions. Corkscrewand other bizarre behaviour may also indicateneurological problems that may be diseaserelated (see F.6 - Viral Encephalopathy and Retinopathy).Patterns of mortalities should be closely monitored,as well as levels of mortality. If losses persistor increase, samples should be sent for laboratoryanalysis (Level II and/or III). Mortalities thatseem to have a uniform or random distributionshould be examined immediately and environmentalfactors during, pre- and post-mortalityrecorded. Mortalities that spread from one areato another may suggest the presence of aninfectious disease agent and should besampled immediately. Affected animals shouldbe kept (isolated) as far away as possible fromunaffected animals until the cause of the mortalitiescan be established.F.1.1.2 Surface Observations (Level I)Generally speaking, no surface observations canbe linked to a single disease problem, however,quick detection of any of the following clinicalsigns, plus follow-up action (e.g., removal or isolationfrom healthy fish, submission of samplesfor laboratory examination), can significantly reducepotential losses.F.1.1.2.1 Skin and Fins (Level I)Damage to the skin and fins can be the consequenceof an infectious disease (e.g., carperythrodermatitis). However, pre-existing lesionsdue to mechanical damage from contact withrough surfaces, such as concrete raceways, orpredator attack (e.g., birds, seals, etc., or chemicaltrauma) can also provide an opportunity forprimary pathogens or secondary pathogens (e.g.,motile aeromonads) to invade and establish. Thisfurther compromises the health of the fish.Common skin changes associated with disease,which should encourage further actioninclude red spots (Fig. F.1.1.2.1a), which maybe pin-point size (petechiae) or larger patches.These tend to occur around the fins, operculum,vent and caudal area of the tail, but maysometimes be distributed over the entire surface.Indications of deeper haemorrhaging orosmotic imbalance problem saredarkenedcolouration. Haemorrhagic lesions may precedeskin erosion, which seriously affect osmoregulationand defense against secondary infections.Erosion is commonly found on the dorsalsurfaces (head and back) and may be causedby disease, sunburn or mechanical damage. Insome species, surface irritation may be indicatedby a build up of mucous or scale loss.Surface parasites, such as copepods, ciliates orflatworms, should also be noted. As with the gills,these may not be a problem under most circumstances,however, if they proliferate to noticeablyhigher than normal numbers (Fig. F.1.1.2.1b),this may lead to secondary infections or indicatean underlying disease (or other stress)problem. The parasites may be attached superficiallyor be larval stages encysted in thefins, or skin. Such encysted larvae (e.g., flatwormdigenean metacercariae) may be detectedas white or black spots (Fig. F.1.1.2.1c)50

F.1 General Techniquesin the skin (or deeper muscle tissue).(K Ogawa)Abnormal growths are associated withtumourous diseases, which can be caused bydisease, such as Oncorhynchus masou virus(see F.4 - Oncorhynchus masou Virus Disease)and Lymphocystis (see F.9 - Lymphocystis), orother environmental problems.The eyes should also be observed closely fordisease indications. Shape, colour, cloudiness,gas bubbles and small haemorrhagic lesions (redspots) can all indicate emerging or actual diseaseproblems. For example, eye enlargementand distension, known as “Popeye”, is associatedwith several diseases (Fig. F.1.1.2.1d).Fig.F.1.1.2.1c. Ayu, Plecoglossus altivelis, infectedwith Posthodiplostomum cuticola (?)metacercariae appearing as black spots onskin.(R Chong)F.1.1.2.2 Gills (Level I)The most readily observable change to soft tissuesis paleness and erosion of the gills(Fig.F.1.1.2.2a). This is often associated withdisease and should be of major concern. Redspots may also be indicative of haemorrhagicproblems, which reduce the critical functioningability of the gills. Fouling, mucous build-up orparasites (ciliate protistans, monogeneans,copepods, fungi, etc.) may also reduce functionalsurface area and may be indicative ofother health problems (Fig.F.1.1.2.2b).Thesemay affect the fish directly or render it moresusceptible to secondary infections.(MG Bondad-Reantaso)Fig. F.1.1.2.1d. Typical ulcerative, popeye, finand tail rot caused by Vibrio spp.(SE McGladdery)Fig.F.1.1.2.2a.Example of gillerosion on Atlanticsalmon, Salmosalar, due to intenseinfestationby the copepodparasite Salmin -cola salmoneus.(MG Bondad-Reantaso)Fig.F.1.1.2.1a. Red spot disease of grass carp.(JR Arthur)Fig.F.1.1.2.2b. Fish gills infected with monogeneanparasites.>Fig.F.1.1.2.1b. Surface parasites, Lerneaecyprinacea infection of giant gouramy.51

F.1 General TechniquesF.1.1.2.3 Body (Level I)Any deviation from normal body shape in a fishis a sign of a health problem. Common changesinclude “pinhead” which usually affects young fishindicating developmental problems; lateral ordorso-vental bends in the spine (i.e., lordosis andscoliosis) can reveal nutritional or environmentalwater quality problems. Another common, andeasily detected, change in body shape is“dropsy”. Dropsy is a distention of the abdomen,giving the fish a “pot belly” appearance. This is astrong indicator of disease problems which mayinclude swelling of internal organs (liver, spleenor kidney), build up of body fluids (clear =oedema; bloody fluids = ascites), parasite problems,or other unknown cause. Dropsy is a commonelement in many of the serious diseaseslisted in the Asia Diagnostic Guide since it is commonlyassociated with systemic disruption of osmoregulationdue to blood-cell or kidney damage.F.1.1.3 Internal Observations (Level I)As a follow up to behavioural changes, samplesof sick fish should be examined and cut openalong the ventral surface (throat to anus). Thiswill allow gross observation of the internal organsand body cavity. A healthy-appearing fish shouldalso be opened up the same way, if the personhas little experience with the normal internal workingsof the fish they are examining. Organ arrangementand appearance can vary betweenspecies.Normal tissues should have no evidence of freefluid in the body cavity, firm musculature, creamwhitefat deposits (where present) around thepyloric caecae, intestine and stomach, a deepred kidney lying flat along the top of the bodycavity (between the spinal cord and swim-bladder),a red liver, a deep red spleen and pancreas.The stomach and intestine may contain food.Gonadal development will vary depending onseason. The heart (behind the gill chamber andwalled off from the body cavity) and associatedbulbous arteriosus should be distinct and shiny.F.1.1.3.1 Body Cavity and Muscle(Level I/II)Clues to disease in a body cavity most commonlyconsist of haemorrhaging and a build up of bloodyfluids. Blood spots in the muscle of the body cavitywall, may also be present. Body cavity wallswhich disintegrate during dissection may indicatea fish that has been dead for a while and whichis, therefore, of little use for accurate diagnosis,due to rapid invasion of secondary saprobionts(i.e., microbes that live on dead and decayingtissues).Necrotic musculature may also indicate amuscle infection, e.g., by myxosporean parasites.This can be rapidly investigated bysquashing a piece of the affected muscle betweentwo glass slides or between a Petri dishlid and base, and examining it under a compoundor dissection microscope. If spore-likeinclusions are present, a parasite problem canbe reasonably suspected. Some microsporidianand myxosporean parasites can form cysts inthe muscle (Fig.F.1.1.3.1a), peritoneal tissues(the membranous network which hold the organsin place in the body cavity), and organsthat easily visible to the naked eye as clumpsor masses of white spheres. These too, requireparasitology identification. Worms may also bepresent, coiled up in and around the organs andperitoneal tissues. None of these parasites(though unsightly) are usually a disease-problem,except where present in massive numberswhich compress or displace the organs(Fig.F.1.1.3.1b).F.1.1.3.2 Organs (Levels I-III)Any white-grey patches present in the liver, kidney,spleen or pancreas, suggest a disease problem,since these normally represent patches ofnecrosis or other tissue damage. In organs suchas kidney and spleen, this can indicate disruptionof blood cell production. Kidney lesions canalso directly affect osmoregulation and liver lesionscan affect toxin and microbial defensemechanisms. Swelling of any of these organs toabove normal size is equally indicative of a diseaseproblem which should be identified, as soonas possible.Swollen intestines (Fig.F.1.1.3.2a andFig.F.1.1.3.2b) should be checked to see if thisis due to food or a build up of mucous. The latteris indicative of feed and waste disposal disruption,as well as intestinal irritation, and is commonlyfound in association with several seriousdiseases. This may also occur due to opportunisticinvasion of bowels that have been irritatedby rapid changes in feed, e.g., by the flagellateprotistan Hexamita salmonis. Mucous filled intestinescan be spotted externally via the presenceof trailing, flocculent or mucous faeces (casts).52

F.1 General Techniques(H Yokoyama)(MG Bondad-Reantaso)(K Ogawa)Fig.F.1.1.3.1a. Myxobolus artus infection in theskeletal muscle of 0+ carp.(K Ogawa)Fig.F.1.3.1b. Ligula sp. (Cestoda) larvae infectionin the body cavity of Japanese yellow goby,Acanthogobius flavimanus.(H Yokoyama)Fig.F.1.3.2a. Distended abdomen of goldfish.F.1.2 Environmental Parameters(Level I)Water quality and fluctuating environmentalconditions, although not of contagious concern,can have a significant effect on finfish health,both directly (within the ranges of physiologicaltolerances) and indirectly (enhancing susceptibilityto infections). This is especiallyimportant for species grown in conditions thatbear little resemblance to the wild situation. Watertemperature, salinity, turbidity, fouling andFig.F.1.3.2b. Japanese Yamame salmon(Onchorynchus masou) fingerlings showingswollen belly due to yeast infection.plankton blooms are all important factors. Highstocking rates, common in intensive aquaculture,predispose individuals to stress as well asminor changes in environmental conditions thatcan precipitate disease. Accumulation of wastefeed indicates either overfeeding or a decreasein feeding activity. In either situation, the breakdownproducts can have a direct toxic effector act as a medium for microbial proliferationand secondary infections. Likewise, other pollutantscan also have a significant effect on fishhealth.F.1.3 General ProceduresF.1.3.1 Pre-Collection Preparation(Level I)Wherever possible, the number of specimensrequired for laboratory examination should beconfirmed before the samples are collected.Larger numbers are generally required forscreening purposes than for diagnosis of mortalities,or other abnormalities. The diagnosticlaboratory which will be receiving the sampleshould also be consulted to ascertain the bestmethod of transportation (e.g., on ice, preservedin fixative, whole or tissue samples). The laboratorywill also indicate if both clinically affected,as well as apparently healthy individuals, arerequired for comparative purposes.Inform the laboratory of exactly what is goingto be sent (i.e., numbers, size-classes or tissuesand intended date of collection and delivery)so the laboratory can be prepared prior tosample arrival. Such preparation can speed upprocessing of a sample (fixative preparation,labeling of slides, jars, cassettes, test-tubes,Petri-plates, data-sheets, etc.) by as much asa day.53

F.1 General TechniquesF.1.3.2 Background Information (Level I)All samples submitted for diagnosis should include as much supporting information as possibleincluding:• reason(s) for submitting the sample (i.e. health screening, certification)• gross observations,feed records,and envronmental parameters• history and origin of the fish population date of transfer and source location(s) if the stock doesnot originate from on-site.These information will help clarify whether handling stress, change of environment or infectiousagents are causes for concern. It will also help speed up diagnosis, risk assessment, and husbandrymanagement and treatment recommendations.F.1.3.3 Sample Collection for Health SurveillanceThe most important factors associated with collection of specimens for surveillance are:• sample numbers that are high enough (see Table F.1.3.3 below)• susceptible species are sampled• sampling includes age-groups and seasons that are most likely to manifest detectable infections.Such information is given under the specific disease sections.Prevalence (%)Population Size 0.5 1.0 2.0 3.0 4.0 5.0 10.050 46 46 46 37 37 29 20100 93 93 76 61 50 43 23250 192 156 110 75 62 49 25500 314 223 127 88 67 54 261000 448 256 136 92 69 55 272500 512 279 142 95 71 56 275000 562 288 145 96 71 57 2710000 579 292 146 96 72 29 27100000 594 296 147 97 72 57 271000000 596 297 147 97 72 57 27>1000000 600 300 150 100 75 60 30Table F.1.3.3 1 . Sample sizes needed to detect at least one infected host in a population of a givensize, at a given prevalence of infection. Assumptions of 2% and 5% prevalences are most commonlyused for surveillance of presumed exotic pathogens, with a 95% confidence limit.F.1.3.4 Sample Collection for Disease Diagnosis (Level I)All samples submitted for disease diagnosis should include as much supporting information aspossible including:• reason(s) for submitting the sample (mortalities, abnormal growth, etc.)• handling activities (net/cage de-fouling, size sorting/grading, site changes, predators, new species/stockintroduction, etc.)1Ossiander, F.J. and G. Wedermeyer. 1973. Journal Fisheries Research Board of Canada 30:1383-1384.54

F.1 General Techniques• environmental changes (rapid water qualitychanges, such as turbidity fluxes, saltwaterincursion into freshwater ponds, unusualweather events, etc.).These information will help clarify whether handlingstress, change of environment or infectiousagents may be a factor in the observedabnormalities/mortalities. Such information isnecessary for both rapid and accurate diagnosis,since it helps focus the investigative proceduresrequired.F.1.3.5 Live Specimen Collection for Shipping(Level I)Collection should take place as close to shippingtime as possible, to reduce mortalitiesduring transportation. This is especially importantfor moribund or diseased fish.The laboratory should be informed of the estimatedtime of arrival of the sample, in order toensure that the laboratory has the materials requiredfor processing prepared before the fisharrive. This shortens the time between removalof the fish from water and preparation of thespecimens for examination (see F.1.3.1).The fish should be packed in double plasticbags, filled with water to one third of theircapacity with the remaining 2/3 volume inflatedwith air/oxygen. The bags should be tightlysealed (rubber bands or tape) and packed insidea styrofoam box or cardboard box linedwith styrofoam. A plastic bag measuring 60 x180 cm is suitable for a maximum of four 200-300 g fish. The volume of water to fish volume/biomass is particularly important for live fishbeing shipped for ectoparasite examination, soadvance checking with the diagnostic laboratoryis recommended. The box should be sealedsecurely to prevent spillage and may be doublepacked inside a cardboard carton. The laboratoryshould be consulted about the packagingrequired.Containers should be clearly labeled as follows:“LIVE SPECIMENS, STORE AT ___ to ___°C,DO NOT FREEZE”(Insert temperature tolerance range of fish beingshipped)If being shipped by air also indicate“HOLD AT AIRPORT AND CALL FOR PICK-UP”(Clearly indicate the name and telephone numberof the person responsible for picking up thepackage, or receiving it at the laboratory).Where possible, ship early in the week to avoiddelivery during the weekend which may lead toimproper storage and loss of samples.Inform the contact person(s) as soon as the shipmenthas been sent and provide the name of thecarrier, flight number, waybill number and estimatedtime of arrival, as appropriate.F.1.3.6 Dead or Tissue Specimen Collectionfor Shipping (Level I)In some cases, samples may be unable to bedelivered live to a diagnostic laboratory due todistance or slow transport connections. In suchcases, diagnostic requirements should be discussedwith laboratory personnel prior to samplecollection. Shipping of non-preserved tissuesor dead specimens may require precautions toprevent contamination or decay. In addition,precautions should be taken to protect ectoparasites,if these are of probable significance.For bacteriology, mycology or virology:• Small fish may be bagged, sealed and transportedwhole on ice/frozen gel-packs.• For larger fish, the viscera can be asepticallyremoved, placed in sterile containers andshipped on ice/frozen gel-packs.• For bacteriology or mycology examinations– ship fish individually bagged and sealed,on ice/frozen gel-packs.• For virology examination - bag fish with fivevolumes of Hanks’ basal salt solution containingeither gentamycin (1,000 mg/ml) or penicillin(800 IU/ml) + dihydrostreptamycin (800mg/ml). Anti-fungal agents such as Mycostatinor Fungizone may also be incorporated at alevel of 400 IU/ml.Note: Intact or live specimens are ideally bestsince dissected tissues rapidly start autolysiseven under ice, making them useless for steriletechnique and bacteriology, particularly for tropicalclimates. Fish destined for bacteriologicalexamination can be kept on ice for a limited period.The icing should be done to ensure that theorgans/tissues destined for examination usingsterile technique are kept at temperatures belowambient water (down to 4°C is a standard low)but not freezing. Individual bagging is also recommendedin order to prevent contamination by2Further details are available in “Recommendations for euthanasia of experimental animals” Laboratory Animals 31:1-32 (1997).55

F.1 General Techniquesone individual within a sample.F.1.3.7 Preservation (Fixation) of TissueSamples (Level I)Fish should be killed prior to fixation. With smallfish, this can be done by decapitation, however,this causes mechanical damage to the tissuesand is unsuitable for larger fish. Alternatively,euthanasia with an overdose of anaestheticis a better (unless examination is for ectoparasites,which may be lost) option. Benzocaineor Etomidate, administered at triple therecommended dose is usually effective foranaesthesizing fish. Injection of anaestheticshould be avoided, wherever possible, due tohandling induced tissue trauma 2 . Putting fishin iced water is also recommended prior to killingof fish.Very small fish, such as fry or alevins, shouldbe immersed directly in a minimum of 10:1(fixative:tissue) volume ratio.For large fish (>6 cm), the full length of the bodycavity should be slit open (normally along themid-ventral line) and the viscera and swim bladdergently displaced to permit incision of eachmajor organ, at least once, to allow maximumpenetration of the fixative. Ideally, the organ, orany lesions under investigation, should be removed,cut into blocks (

F.1 General Techniquesestimated time of arrival, as appropriate.F.1.4 Record-Keeping (Level I)It is critical to establish, and record, normalbehaviour and appearance to compare with observationsmade during disease events. Recordkeepingis, therefore, an essential component ofeffective disease management. For fish, manyof the factors that should be recorded on a regularbasis are outlined in sections F.1.4.1, F.1.4.2and F.1.4.3.F.1.4.1 Gross Observations (Level I)These can be included in routine records of fishgrowth that, ideally would be monitored on aregular basis, either by sub-sampling from tanksor ponds, or by estimates made from surfaceobservations.For hatcheries, critical information that should berecorded include:• feeding activity• growth• mortalitiesThese observations should be recorded daily, forall stages, including date, time, tank #, broodstock(where there are more than one) and food source.Dates and times of tank and water changes, pipeflushing/back-flushing and/or disinfection,should also be recorded. Ideally, these recordsshould be checked (signed off) regularly by theperson responsible for maintaining the facility.For pond or net/cage sites, observations whichneed to be recorded include:• growth• fouling• mortalitiesThese should be recorded with date, site locationand any relevant activities (e.g., sample collectionfor laboratory examination). As elsewhere,these records should be checked regularly bythe person responsible for the facility.F.1.4.2 Environmental Observations(Level I)Environmental observations are most applicableto open water, ponds, cage and net culture systems.Information that should be recorded include:• weather• water temperature• oxygen• salinity• turbidity (qualitative evaluation or Secchi disc)• algal blooms• human activity (handling, neighbouring landuse/water activities)• pHThe frequency of these observations will vary withsite and fish species. Where salinity or turbidityrarely vary, records may only be required duringrainy seasons or exceptional weather conditions.Temperate climates may require more frequentwater temperature monitoring than tropicclimates. Human activity(ies) should also be recordedon an “as it happens” basis, since theremay be time-lag effects. In all cases, date andtime should be recorded, as parameters suchas temperature and pH can vary markedly duringthe day, particularly in open ponds and inter-tidalsites.It may not always be possible to monitor oxygenlevels in the pond. However, the farmer shouldbe aware that in open non-aerated ponds, oxygenlevels are lowest in the early morning whenplants (including algae) have used oxygen overnight.Photosynthesis and associated oxygenproduction will only commence after sunrise.F.1.4.3 Stocking Records (Level I)All movements of fish into and out of a hatcheryor site should be recorded, including:• the source of the broodstock/eggs/larvae/juvenilesand their health certification• the volume or number of fish• condition on arrival• date and time of delivery and name of personresponsible for receiving the fish• date, time and destination of stock shippedoutfrom a hatchery or site.Such records are also applicable (but less critical)to movements between tanks, ponds, cageswithin a site. Where possible, animals from differentsources should not be mixed. If mixing isunavoidable, keep strict records of which sourcesare mixed and dates of new introductions intothe holding site or system.F.1.5 ReferencesChinabut, S. and R.J. Roberts. 1999. Pathologyand histopathology of epizootic ulcerative syndrome(EUS). Aquatic Animal Health ResearchInstitute. Department of Fisheries, Royal ThaiGovernment. Bangkok, Thailand. 33p.57

F.1 General TechniquesClose, B., K. Banister, V. Baumans, E. Bernoth,N. Bromage, J. Bunyan, W. Erhardt, P.Flecknell, N. Gregory, H. Hackbarth, D.Morton and C. Warwick. 1997. Recommendationsfor euthanasia of experimental animals:Part 2. Lab. Anim.31:1-32.Ossiander, F.J. and G. Wedermeyer. 1973.Computer program for sample size requiredto determine disease incidence in fishpopulations. J. Fish. Res. Bd. Can. 30: 1383-1384.Tonguthai, K., S. Chinabut, T. Somsiri, P.Chanratchakool, and S. Kanchanakhan.1999. Diagnostic Procedures for Finfish Diseases.Aquatic Animal Health Research Institute,Department of Fisheries, Bangkok,Thailand.58

VIRAL DISEASES OF FINFISHESF.2 EPIZOOTIC HAEMATOPOIETICNECROSIS (EHN)F.2.1 Background InformationF.2.1.1 Causative AgentEpizootic Haematopoietic Necrosis (EHN) iscaused by a double-stranded DNA, non-envelopedIridovirus known as EpizooticHaematopoeitic Necrosis Virus (EHNV). Thisvirus shares at least one antigen with iridovirusesinfecting sheatfish (Silurus glanis) and the catfish(Ictalurus melas) in Europe and with amphibianiridoviruses from North America (frog virus3) and Australia (Bohle iridovirus). Recently, theOIE included the two agents, European catfishvirus and European sheatfish virus, as causativeagents of EHN (OIE 2000a; http://www.oie.int).Current classification in the genus Ranavirus isunder review (see http://www.ncbi.nlm.nih.gov/ICTV). More detailed information about the diseasecan be found in the OIE Diagnostic Manualfor Aquatic Animal Diseases (OIE 2000a).F.2.1.2 Host RangeEHNV infects redfin perch (Perca fluviatilis) andrainbow trout (Oncorhynchus mykiss). Other fishspecies found to be susceptible to EHNV afterbath exposure are Macquarie perch (Macquariaaustralasica), mosquito fish (Gambussia affinis),silver perch (Bidyanus bidyanus) and mountaingalaxias (Galaxias olidus).F.2.1.3 Geographic DistributionHistorically, the geographic range of EHNVinfections has been restricted to mainlandAustralia. However, a recent OIE decision toinclude sheatfish and catfish iridoviruses ascauses of EHN, increased the geographicdistribution to include Europe. A related virusrecently isolated from pike-perch in Finland, wasfound to be immunologically cross-reactive butnon-pathogenic to rainbow trout.F.2.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System(1999-2000)Australia reported the occurrence of EHN inVictoria (last year 1996), New South Wales (lastyear 1996) and South Australia (1992). It wasalso known to have occurred in New South Walesduring first quarter of 2000, with annualoccurrence in the Australian Capital Territory(without laboratory confirmation) (OIE 1999,2000b).India reported EHN during last quarter of 1999affecting murrels and catfishes (OIE 1999).F.2.2 Clinical AspectsThere are no specific clinical signs associatedwith EHN. Mortalities are characterised bynecrosis of liver (with or without white spots),spleen, haematopoietic tissue of the kidney andother tissues. Disruption of blood function leadsto osmotic imbalance, haemorrhagic lesions,build up of body fluids in the body cavity. Thebody cavity fluids (ascites) plus enlarged spleenand kidney may cause abdominal distension(dropsy).Clinical disease appears to be associated withpoor water quality, as well as water temperature.In rainbow trout, disease occurs at temperaturesfrom 11 to 17˚C (in nature) and 8 - 21˚C(experimental conditions). No disease is foundin redfin perch at temperatures below 12˚Cunder natural conditions. Both juvenile and adultredfin perch can be affected, but juvenilesappear more susceptible (Fig.F.2.2a). EHNV hasbeen detected in rainbow trout ranging from fryto market size, although mortality occurs mostfrequently in 0+ - 125 mm fork-length fish.F.2.3 Screening MethodsMore detailed information on methods forscreening EHN can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000a), at http://www.oie.int, or selectedreferences.As with other disease agents, screening for thepresence of an infectious agent in a sub-clinicalpopulation requires larger sample numbers thanfor a disease diagnosis. Numbers will varyaccording to the confidence level required (seeF.1.3.3).F.2.3.1 PresumptiveF.2.3.1.1 Gross Observations (Level 1)and Histopathology (Level II)It is not possible to detect infections in sub-clinicalfish, using gross observations (Level I) orhistopathology (Level II).F.2.3.1.2 Virology (Level III)EHNV can be isolated on Bluegill Fin 2 (BF-2) orFathead Minnow (FHM) cell lines. This requiressurveillance of large numbers (see Table F.1.3.3)59

F.2 Epizootic HaematopoieticNecrosis (EHN)of sub-clinical fish to detect low percentagecarriers.F.2.3.2 ConfirmatoryF.2.3.2.1 Immunoassays (Level III)Suspect cytopathic effects (CPE) in BF-2 or FHMcell-lines require confirmation of EHNV as thecause through immunoassay (indirect fluorescentantibody test (IFAT) or enzyme linkedimmunosorbent assay (ELISA) or PolymeraseChain Reaction (PCR) (Level III).F.2.4 Diagnostic MethodsMore detailed information on methods fordiagnosis of EHN can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000a), at http://www.oie.int, or selectedreferences.EHNV is a highly resistant virus that canwithstand freezing for prolonged periods, thus,fish may be stored and or/transported frozenwithout affecting the diagnosis.F.2.4.1 PresumptiveF.2.4.1.1 Gross Observation (Level I)As described under F.2.2, mass mortalities ofsmall redfin perch under cool water conditions(< 11 °C), which include cessation of feeding,abdominal distension, focal gill and finhaemorrhage, as well as overall skin darkening,should be considered suspect for EHNV infection.Similar observations in rainbow trout fingerlings(11-17 °C) may also be considered suspect,but the conditions are not specific to EHNin either host.Necropsy may reveal liver and spleenenlargement or focal pale spots on the liver, butthese, again, are non-specific.F.2.4.1.2 Histopathology (Level II)Histopathology in haematopoietic kidney, liver,spleen and heart tissues are similar in bothinfected redfin perch and rainbow trout, althoughperch livers tend to have larger focal or locallyextensive areas of necrosis. Gills of infectedperch show focal blood clots, haemorrhage andfibrinous exudate. Focal necrosis occurs in thepancreas and intestinal wall. In the former tissuesite necrosis can become extensive.F.2.4.1.3 Virology (Level III)Whole alevin or juvenile perch (

F.2 Epizootic HaematopoieticNecrosis (EHN)F.2.6Control Measures(AAHL)Prevention of movement of infected fishbetween watersheds, and minimising contactbetween trout farms and surrounding perchpopulations is recommended. In addition,reducing bird activity at farm sites may beeffective in reducing the chances of exposureand spread. Precautionary advice andinformation for recreational fishermen usinginfected and uninfected areas may also reduceinadvertent spread of EHN.F.2.7Selected ReferencesGould, A.R., A.D. Hyatt, S.H. Hengstberger, R.J.Whittington, and B.E.H. Couper. 1995. A polymerasechain reaction (PCR) to detect epizooticnecrosis virus and Bohle iridovirus. Dis.Aquat. Org. 22: 211-215.Hyatt, A.D., B.T. Eaton, S. Hengstberger,G.Russel. 1991. Epizootic haematopoieticnecrosis virus: detection by ELISA, immunohistochemistryand electron microscopy. J.Fish Dis.14: 605-618.Fig.F.2.2a. Mass mortality of single species ofredfin perch. Note the small size of fish affectedand swollen stomach of the individual to thecentre of the photograph. Note thecharacteristic haemorrhagic gills in the fish onthe left in the inset.(EAFP)OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Whittington, R.J. and K.A. Steiner. 1993. Epizootichaematopoietic necrosis virus (EHNV):improved ELISA for detection in fish tissuesand cell cultures and an efficient method forrelease of antigen from tissues. J. Vir.Meth.43: 205-220.Whittington, R.J., L.A. Reddacliff, I. Marsh, C.Kearns, Z. Zupanovic, Z. and R.B.Callinan.1999. Further observations on theepidemiology and spread of epizootichaematopoietic necrosis virus (EHNV) infarmed rainbow trout Oncorhynchus mykiss insoutheastern Australia and a recommendedsampling strategy for surveillance. Dis.Aquat. Org. 35: 125-130.F.3Fig.F.3.2a. IHN infected fry showing yolk sachaemorrhages.(EAFP)Fig.F.3.2b. Clinical signs of IHN infected fishinclude darkening of skin, haemorrhages on theabdomen and in the eye around the pupil.61

F.3 INFECTIOUS HAEMATOPOIETICNECROSIS (IHN)F.3.1 Background InformationF.3.1.1 Causative AgentInfectious Haematopoietic Necrosis (IHN) iscaused by an enveloped single stranded RNA(ssRNA) Rhabdovirus, known as InfectiousHaematopoietic Necrosis Virus (IHNV). It is currentlyunassigned to genus, but the InternationalCommittee on Taxonomy of Viruses (ICTV) iscurrently reviewing a new genus – Novirhabdovirus– which is proposed to include VHSV andIHNV (see http://www.ncbi.nlm.nih.gov/ICTV).More detailed information about the disease canbe found in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a).F.3.1.2 Host RangeIHNV infects rainbow or steelhead trout(Oncorhynchus mykiss), sockeye salmon(O. nerka), chinook (O. tshawytscha), chum(O.keta), yamame (O. masou), amago(O. rhodurus), coho (O. kisutch), and Atlanticsalmon (Salmo salar). Pike fry (Esox lucius),seabream and turbot can also be infected underexperimental conditions.F.3.1.3 Geographic DistributionHistorically, the geographic range of IHN waslimited to the Pacific Rim of North America but,more recently, the disease has spread tocontinental Europe and Asia.F.3.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System(1999- 2000)India reported occurrence of IHN during lastquarter of 1999 affecting murrels and catfishes;Korea RO reported IHN among rainbow troutduring 3 rd and 4 th (September) quarters of 2000while Japan reported occurrence of IHN everymonth during 1999 and 2000 (OIE 1999, 2000b).F.3.2 Clinical AspectsAmong individuals of each fish species, there isa high degree of variation in susceptibility toIHNV. Yolk-sac fry (Fig.F.3.2a) are particularlysusceptible and can suffer 90-100% mortality. Inrainbow trout, such mortalities are correlated withwater temperatures

F.3 Infectious HaematopoieticNecrosis (IHN)F.3.4.1 PresumptiveF.3.4.1.1 Gross Observations (Level I)Behavioural changes are not specific to IHN butmay include lethargy, aggregation in still areasof the pond with periodic bursts of erraticswimming (see F.3.2) and loss of equilibrium.Changes in appearance include darkdiscolouration of the body (especially the dorsalsurface and tail fin regions), especially in yolksacfry stages (90-100% mortality). Theabdomen can be distended due toaccumulation of fluids in the body cavity(dropsy) and haemorrhaging may be visible atthe base of the fins, on the operculum andaround the eyes. The eyes may also show signsof water imbalance in the tissues by bulging(“pop-eye”). There may be vent protrusion andtrailing white/mucoid casts.F.3.4.1.2 Histopathology (Level II)Tissue sections show varying degrees ofnecrosis of the kidney and spleen(haematopoietic) tissues, as well as in the brainand digestive tract.F.3.4.1.3 Virology (Level III)Whole alevins (body length ≤4 cm), visceraincluding kidney (fish 4 – 6 cm in length) or kidney,spleen and brain tissues from larger fish, arerequired for isolating the virus on EPC or BF-2cell lines. Confirmation of IHNV being the causeof any resultant CPE requires immunoassayinvestigation, as described below.F.3.4.2 ConfirmatoryF.3.4.2.1 Immunoassays (IFAT or ELISA)(Level III)Diagnosis of IHNV is achieved via immunoassayof isolates from cell culture using IFAT or ELISA,or immunological demonstration of IHNV antigenin infected fish tissues.F.3.4.2.2 Transmission Electron Microscopy(TEM) (Level III)TEM of cells infected in cell-culture reveals enveloped,slightly pleomorphic, bullet shaped virions,45-100 nm in diameter and 100-430 nm long.Distinct spikes are evenly dispersed over mostof the surface of the envelope (although thesemay be less evident under some cell-cultureconditions). The nucleocapsids are coiled andshow cross-banding (4.5 – 5.0 nm apart) innegative stain and TEM. Viral replication takesplace in the cytoplasm with particle maturationat the cell membrane or the Golgi cisternae.F.3.5 Modes of TransmissionIHNV is usually spread by survivors of infections,which carry sub-clinical infections. When suchfish mature, they may shed the virus duringspawning. Clinically infected fish can also spreadthe disease by shedding IHNV with faeces, urine,spawning fluids and mucus secretions. Othersources of infection include contaminatedequipment, eggs from infected fish, and bloodsucking parasites (e.g., leeches, Argulus spp.).Fish-eating birds are believed to be anothermechanism of spread from one site to another.The most prominent environmental factoraffecting IHN is water temperature. Clinicaldisease occurs between 8˚C and 15˚C undernatural conditions. Outbreaks rarely occur above15°C.F.3.6 Control MeasuresControl methods currently rely on avoidancethrough thorough disinfection of fertilised eggs.Eggs, alevins and fry should be reared on virusfreewater supplies in premises completelyseparated from possible IHNV-positive carriers.Broodstock from sources with a history of IHNoutbreaks should also be avoided whereverpossible. At present, vaccination is only at anexperimental stage.As with viral haemorrhagic septicaemia virus(VHSV, see F.8), good over-all fish healthcondition seems to decrease the susceptibilityto overt IHN, while handling and other types ofstress frequently cause sub-clinical infection tobecome overt.F.3.7 Selected ReferencesEnzmann, P.J., D. Fichtner, H. Schuetze, andG. Walliser. 1998. Development of vaccinesagainst VHS and IHN: Oral application,molecular marker and discrimination ofvaccinated fish from infected populations. J.Appl. Ichth.14: 179-183.Gastric, J., J. Jeffrey. 1991. Experimentallyinduced diseases in marine fish with IHHNVand a rhabdovirus of eel. CNEVA –Laboratoirede Pathologie des Animaux Aquatiques B.P.63

F.3 Infectious HaematopoieticNecrosis (IHN)70 – 29289 Plouzane, France. EAS Spec.Publ. No. 14.Hattenberger-Baudouy, A.M., M. Dabton, G.Merle, and P. de Kinkelin. 1995. Epidemiologyof infectious haematopoietic necrosis (IHN) ofsalmonid fish in France: Study of the courseof natural infection by combined use of viralexamination and seroneutralisation test anderadication attempts. Vet. Res. 26: 256-275 (inFrench).OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Park, M.S., S.G. Sohn, S.D. Lee, S.K. Chun,J.W. Park, J.L Fryer, and Y.C. Hah. 1993.Infectious haematopoietic necrosis virus insalmonids cultured in Korea. J. Fish Dis. 16:471-478.Schlotfeldt, H.-J. and D.J. Alderman. 1995.What Should I Do? A Practical Guide for theFreshwater Fish Farmer. Suppl. Bull. Eur.Assoc. Fish Pathol. 15(4). 60p.64

F.4 ONCORHYNCHUS MASOUVIRUS (OMV)F.4.1 Background InformationF.4.1.1 Causative AgentOncorhynchus masou virus disease (OMVD) iscaused by Oncorhynchus masou virus (OMV)is believed to belong to the FamilyHerpesviridae, based on an icosahedraldiameter of 120-200 nm, and enveloped,dsDNA properties. OMV is also known asYamame tumour virus (YTV), Nerka virusTowada Lake, Akita and Amori prefecture(NeVTA), coho salmon tumour virus (CSTV),Oncorhynchus kisutch virus (OKV), coho salmonherpesvirus (CSHV), rainbow trout kidney virus(RKV), or rainbow trout herpesvirus (RHV). OMVdiffers from the herpesvirus of Salmonidae type1, present in the western USA. Currently thissalmonid herpesvirus has not beentaxonomically assigned (see http://www.ncbi.nlm.nih.gov/ICTV). More detailedinformation about the disease can be found inthe OIE Diagnostic Manual for Aquatic AnimalDiseases (OIE 2000a).F.4.1.2 Host RangeKokanee (non-anadromous sockeye) salmon(Oncorhynchus nerka) is most susceptible,followed, in decreasing order of susceptibility, bymasou salmon (O. masou), chum salmon(O. keta), coho salmon (O. kisutch) and rainbowtrout (O. mykiss).F.4.1.3 Geographic DistributionOMVD is found in Japan and, probably (as yetundocumented) the coastal rivers of eastern Asiathat harbour Pacific salmon.F.4.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System (1999-2000)Japan reported OMVD during all months of 1999and 2000; and suspected by Korea RO for 1999,and during first two quarters of 2000 (OIE 1999,2000b).F.4.2 Clinical AspectsOMV infects and multiplies in endothelial cells ofblood capillaries, spleen and liver, causingsystemic oedema and haemorrhaging. Onemonth-oldalevins are the most susceptibledevelopment stage. Kidney, spleen, liver andtumours are the sites where OMV is mostabundant during the course of overt infection.Four months after the appearance of clinicalsigns, some surviving fish may developepitheliomas (grossly visible tumours) around themouth (upper and lower jaw) and, to a lesserextent, on the caudal fin operculum and bodysurface. These may persist for up to 1 year. In 1yr-old coho salmon, chronic infections manifestthemselves as skin ulcers, white spots on theliver and papillomas on the mouth and bodysurface. In rainbow trout, however, there are few(if any) external symptoms, but intestinalhaemorrhage and white spots on the liver areobserved.Survivors of OMVD develop neutralisingantibodies which prevent re-infection, however,they can remain carriers of viable virus.F.4.3 Screening MethodsMore detailed information on screening methodsfor OMV can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE 2000a),at http://www.oie.int or selected references.F.4.3.1 PresumptiveF.4.3.1.1 Gross Observations (Level I)Persistent superficial tumours are rare, butindicative of a potential carrier of viable OMV.Species, such as rainbow trout show no suchlesions. Sub-clinical carriage cannot normally bedetected using histology.F.4.3.1.2 Virology (Level III)OMV can be isolated from reproductive fluids,kidney, brain and spleen tissue samples onChinook salmon embryo-214 (CHSE-214) orrainbow trout gonad-2 (RTG-2) cell lines. Anyresultant CPE requires further immunological andPCR analyses to confirm the identity of the virusresponsible (see F.4.3.2.1).F.4.3.2 ConfirmatoryF.4.3.2.1 Immunoassays and Nucleic AcidAssays (Level III)Cytopathic effect (CPE) from cell cultures, as wellas analyses of reproductive fluids, kidney, brainand spleen tissue samples from suspect fish canbe screened using specific neutralisation antibodytests, indirect immunofluorescent antibodytests (IFAT) with immunoperoxidase staining,ELISA or Southern Blot DNA probe assays.65

F.4 Oncorhynchus Masou Virus (OMV)F.4.4 Diagnostic Methods(M Yoshimizu)More detailed information on diagnosticmethods for OMV can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000b), at http://www.oie.int or selectedreferences.F.4.4.1 PresumptiveF.4.4.1.1 Gross Observation (Level I)Behavioural changes include lethargy andaggregation around the water inflow by youngsalmonids of susceptible species. Pin-pointhaemorrhaging or ulcers may be visible on theskin, along with darkened colouration. Popeyemay also be present. Internally, white spots maybe present on the liver (Fig.F.4.4.1.1a). Afterapproximately 4 months, surviving fish may showsigns of skin growths around the mouth(Fig.F.4.4.1.1b) or, less commonly, on theoperculum, body surface or caudal fin area.Fig.F.4.4.1.1a. OMV-infected chum salmonshowing white spots on the liver.(M Yoshimizu)F.4.4.1.2 Histopathology (Level II)Tissue sections from suspect fish may showlesions with enlarged nuclei in the epithelialtissues of the jaw, inner operculum and kidney.F.4.4.1.3 Virology (Level III)Whole alevin (body length ≤ 4 cm), viscera includingkidney (4 – 6 cm length) or, for larger fish,skin ulcerative lesions, neoplastic (tumourous tissues),kidney, spleen and brain are required fortissue culture using CHSE-214 or RTG-2 celllines.The cause of resultant CPE should be confirmedas viral using the procedures outlined inF.4.3.2.1.Fig.F.4.4.1.1b. OMV-induced tumourdeveloping around the mouth of chum salmonfingerling.(M Yoshimizu)F.4.4.1.4 Transmission Electron Microscopy(TEM) (Level III)Detection of virions in the nuclei of affected tissuesand tumours by TEM. The dsDNA virionsare enveloped and icosahedral, measuring 120-200 nm in diameter (Fig.F.4.4.1.3).F.4.4.2ConfirmatoryF.4.4.2.1 Gross Observations (Level I)Gross behaviour and clinical signs at the onsetof OMVD are not disease specific. Thus, confirmatorydiagnosis requires additional diagnosticexamination or occurrence with a docu-Fig.F.4.4.1.3. OMV particles isolated frommasou salmon, size of nucleocapsid is 100 to110 nm.66

F.4 Oncorhynchus Masou Virus (OMV)mented history of OMVD on-site or mortalitiesseveral months preceding the appearance ofepithelial lesions and tumours.F.4.4.2.2 Virology (Level III)As described for F.4.3.1.2F.4.4.2.3 Immunoassays and Nucleic AcidAssays (Level III)As described for F.4.3.2.1.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Yoshimizu, M., T. Nomura, Y. Ezura, Y. and T.Kimura. 1993. Surveillance and control of infectioushaematopoieticnecrosis virus (IHNV)and Oncorhynchus masou virus (OMV) of wildsalmonid fish returning to the northern part ofJapan 1976-1991. Fish. Res.17: 163-173.F.4.5 Modes of TransmissionVirus is shed with faeces, urine, external andinternal tumours, and, possibly, with skinmucus. Reservoirs of OMV are clinically infectedfish as well as wild or cultured sub-clinicalcarriers. Maturation of survivors of early lifehistoryinfections may shed virus with theirreproductive fluids (“egg associated”, ratherthan true vertical transmission). Egg-associatedtransmission, although less frequent than othermechanisms of virus release, is the most likelysource of infection in alevins.F.4.6Control MeasuresThorough disinfection of fertilised eggs, in additionto rearing of fry and alevins, in water free ofcontact with contaminated materials or fish, hasproven effective in reducing outbreaks ofOMVD.Water temperatures

F.5 INFECTIOUS PANCREATICNECROSIS (IPN)F.5.1 Background InformationF.5.1.1 Causative AgentInfectious pancreatic necrosis (IPN) is causedby a highly contagious virus, Infectious pancreaticnecrosis virus (IPNV) belonging to theBirnaviridae. It is a bi-segmented dsRNA viruswhich occurs primarily in freshwater, but appearsto be saltwater tolerant. More detailed informationabout the disease can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000a).F.5.1.2 Host RangeIPN most commonly affects rainbow trout(Oncorhynchus mykiss), brook trout (Salvelinusfontinalis), brown trout (Salmo trutta), Atlanticsalmon (Salmo salar) and several Pacific salmonspecies (Oncorhynchus spp.). Serologicallyrelated are reported from Japanese yellowtailflounder (Seriola quinqueradiata), turbot(Scophthalmus maximus), and halibut(Hippoglossus hippoglossus). Sub-clinicalinfections have also been detected in a widerange of estuarine and freshwater fish speciesin the families Anguillidae, Atherinidae, Bothidae,Carangidae, Cotostomidae, Cichlidae, Clupeidae,Cobitidae, Coregonidae, Cyprinidae, Esocidae,Moronidae, Paralichthydae, Percidae, Poecilidae,Sciaenidae, Soleidae and Thymallidae.F.5.1.3 Geographic DistributionThe disease has a wide geographical distribution,occurring in most, if not all, major salmonidfarming countries of North and South America,Europe and Asia.F.5.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System(1999-2000)IPN was reported by Japan and suspected inKorea RO for 1999; in 2000, reported by Japanfor the whole year except for the month ofFebruary, and by Korea RO in April (OIE 1999,2000b).F.5.2 Clinical AspectsThe first sign of IPN in salmonid fry is the suddenonset of mortality. This shows a progressiveincrease in severity, especially following introductionof feed to post-yolk-sac fry. IPN also affectsAmerican salmon smolt shortly after transferto sea-cages. Clinical signs include darkeningof the lower third of the body and smallswellings on the head (Fig.F.5.2.a) and a pronounceddistended abdomen (Fig.F.5.2b andFig.F.5.2c) and a corkscrewing/spiral swimmingmotion. Some fish may also show ‘pop-eye’ deformities.Cumulative mortalities may vary fromless than 10% to more than 90% dependingon the combination of several factors such asvirus strain, host and environment. Survivorsof the disease, at early or late juvenile stages,are believed to be carriers of viable IPNV forlife. Mortality is higher when water temperaturesare warm, but there is no distinct seasonalcycle.The pancreas, oesophagus and stomach becomeulcerated and haemorrhagic. The intestinesempty or become filled with clear mucous (thismay lead to white fecal casts).F.5.3 Screening MethodsMore detailed information on screening methodsfor IPN can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE 2000b),at http://www.oie.int or selected references.As with other disease agents, screening for thepresence of an infectious agent in a sub-clinicalpopulation requires larger sample numbers thanfor a disease diagnosis. Numbers will varyaccording to the confidence level required (seeF.1.3.3).F.5.3.1 PresumptiveF.5.3.1.1 Gross Observations (Level I)and Histopathology (Level II)Carriers of sub-clinical infections show noexternal or internal evidence of infection at thelight microscope level.F.5.3.1.2 Virology (Level III)Screening procedures use viral isolation onChinook Salmon Embryo (CHSE-214) or BluegillFin (BF-2) cell lines. The cause of any CPE,however, has to be verified using confirmatorytechniques (F.5.3.2.2). Fish material suitable forvirological examination include whole alevin(body length ≤ 4 cm), viscera including kidney(fish 4 – 6 cm in length) or, liver, kidney and spleenfrom larger fish.68

F.5 Infectious Pancreatic Necrosis(IPN)F.5.3.2 Confirmatory(J Yulin)F.5.3.2.1 Immunoassays and MolecularProbe Assays (Level III)The viral cause of any CPE on CHSE-214 orBF-2 cell lines has to be confirmed by either animmunoassay (Neutralisation test or ELISA) or(EAFP)Fig.F.5.4.1.3. CPE of IHNV.(J Yulin)Fig.F.5.2a. IPN infected fish showing darkcolouration of the lower third of the body andsmall swellings on the head.(J Yulin)Fig.F.5.4.1.4. IPN Virus isolated from rainbowtrout fimported from Japan in1987. Virusparticles are 55 nm in diameter.Fig.F.5.2b. Rainbow trout fry showing distendedabdomen characteristic of IPN infection.Eyed-eggs of this species were importedfrom Japan into China in 1987.(EAFP)PCR techniques, including reversetranscriptasePCR (RT-PCR) and in situhybridization (ISH).F.5.4 Diagnostic MethodsMore detailed information on diagnosticmethods for IPN can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000b), at http://www.oie.int or selectedreferences.F.5.4.1 PresumptiveF.5.4.1.1 Gross Observations (Level I)Fig.F.5.2c. Top: normal rainbow trout fry, below:diseased fry.Clinical signs in salmonid fry and parr includelying on the bottom of tanks/ponds, or showingcork-screw swimming behaviour. High mortalitiesmay occur when fry are first fed or in smolt shortlyafter transfer to seawater. Chronic low mortalitiesmay persist at other times. Dark discolouration(especially of the dorsal and tail surfaces) maybe accompanied by swollen abdomens, pop-eyeand/or pale faecal casts.69

F.5 Infectious Pancreatic Necrosis(IPN)F.5.4.1.2 Histopathology (Level II)Tissue pathology is characterised by necroticlesions and ulcers in the pancreas, oesophagusand stomach. The intestines may be empty orfilled with clear mucus (NB difference fromparasite infection by Hexamita inflata(Hexamitiasis), where there is a yellowish mucusplug).F.5.4.1.3 Virology (Level III)As described for screening (F.5.3.1.2), fish materialsuitable for virological examination includewhole alevin (body length ≤ 4 cm), viscera includingkidney (fish 4 – 6 cm in length) or, liver,kidney and spleen for larger fish. The virus(Fig.F.5.4.1.3) can be isolated on CHSE-214 orBF-2 cell lines, but the cause of resultant CPEhas to be verified using confirmatory techniques(F.5.3.2).F.5.4.1.4 Transmission Electron Microscopy(TEM) (Level III)The ultrastructural characteristics of IPNV areshared by most aquatic birnavididae, thus,immunoassay or nucleic acid assays are requiredfor confirmation of identity. Birnaviruses are nonenveloped,icosahedral viruses, measuringapproximately 60 nm in diameter (Fig.F.5.4.1.4).The nucleic acid component is bi-segmented,dsRNA, which can be distinguished usingstandard histochemistry.F.5.4.2 ConfirmatoryF.5.4.2.1 Virology and Immunoassay(Level III)As described for screening (F.5.3.2.1), the viralcause of any CPE on CHSE-214 or BF-2 celllines has to be confirmed by either animmunoassay (Neutralisation test or ELISA) orPCR techniques, including RT-PCR and ISH.F.5.5Modes of TransmissionThe disease is transmitted both horizontallythrough the water route and vertically via the egg.Horizontal transmission is achieved by viral uptakeacross the gills and by ingestion. The virusshows strong survival in open water conditionsand can survive a wide range of environmentalparameters. This, in addition to its lack of hostspecificity, provides gives IPNV the ability to persistand spread very easily in the open-waterenvironment.F.5.6 Control MeasuresPrevention methods include avoidance offertilised eggs from IPNV carrier broodstock anduse of a spring or borehole water supply (freeof potential reservoir fish). Surface disinfectionof eggs has not been entirely effective inpreventing vertical transmission.Control of losses during outbreaks involves reducingstocking densities and dropping watertemperatures (in situations where temperaturecan be controlled).Vaccines are now available for IPN and theseshould be considered for fish being grown in IPNVendemic areas.F.5.7 Selected ReferencesFrost, P. and A. Ness. 1997. Vaccination ofAtlantic salmon with recombinant VP2 ofinfectious pancreatic necrosis virus (IPNV),added to a multivalent vaccine, suppressesviral replication following IPNV challenge. FishShellf. Immunol. 7: 609-619.Granzow, H., F. Weiland, D. Fichtner, and P.J.Enzmann. 1997. Studies on the ultrastructureand morphogenesis of fish pathogenic virusesgrown in cell culture. J. Fish Dis. 20: 1-10.Lee, K.K., T.I. Yang, P.C. Liu, J.L. Wu, and Y.L.Hsu. 1999. Dual challenges of infectiouspancreatic necrosis virus and Vibrio carchariaein the grouper Epinephelus sp.. Vir. Res. 63:131-134.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Seo, J-J., G. J. Heo, and C.H. Lee. 1998.Characterisation of aquatic Birnavirusesisolated from Rockfish (Sebastes schlegeli)cultured in Korea. Bull. Eur. Assoc. Fish Pathol.18: 87-92.70

F.5 Infectious Pancreatic Necrosis(IPN)Schlotfeldt, H.-J. and D.J. Alderman. 1995.What Should I Do? A Practical Guide for theFreshwater Fish Farmer. Suppl. Bull. Eur.Assoc. Fish Pathol. 15(4). 60p.Wang, W.S., Y.L. Wi, and J.S. Lee. 1997. Singletube, non interrupted reverse transcriptasePCR for detection of infectious pancreatic necrosisvirus. Dis. Aquat. Org. 28: 229-233.Yoshinaka, T., M. Yoshimizu, and Y. Ezura. 1998.Simultaneous detection of infectioushaematopoietic necrosis virus (IHNV) and infectiouspancreatic necrosis virus(IPNV) by reverse transcriptase (RT) polymerasechain reaction (PCR). Fish. Sci. 64:650-651.71

F.6 VIRAL ENCEPHALOPATHY ANDRETINOPATHY (VER)F.6.1F.6.1.1Background InformationCausative AgentsViral Encephalopathy and Retinopathy (VER) iscaused by icosahedral, non-envelopednodaviruses, 25-30 nm in diameter. Theseagents are also known as Striped Jack NervousNecrosis Virus (SJNNV), Viral Nervous Necrosis(VNN) and Fish Encephalitis Virus (FEV). Allshare serological similarities with the exceptionof those affecting turbot (F.6.1.2). More detailedinformation about the disease can be found inthe OIE Diagnostic Manual for Aquatic AnimalDiseases (OIE 2000a).F.6.1.2Host RangeThe pathology of VER occurs in larval and,sometimes, juvenile barramundi (sea bass, Latescalcarifer), European sea bass (Dicentrarchuslabrax), turbot (Scophthalmus maximus), halibut(Hippoglossus hippoglossus), Japaneseparrotfish (Oplegnathus fasciatus), red-spottedgrouper (Epinepheles akaara), and striped jack(Pseudocaranx dentex). Disease outbreaks withsimilar/identical clinical signs have been reportedin tiger puffer (Takifugu rubripes), Japaneseflounder (Paralichthys olivaceus), kelp grouper(Epinephelus moara), brown spotted grouper(Epinephelus malabaricus), rock porgy(Oplegnathus punctatus), as well as othercultured marine fish species.F.6.1.3 Geographic DistributionVER occurs in Asia, the Mediterranean and thePacific.F.6.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System(1999-2000)Australia reported VER occurrence during 8 of12 months in 1999, and 7 of 12 months in 2000.Japan reported VER in 6 of 12 months in 2000,and 3 of 12 months in 1999. Last major outbreakreported by Singapore was in 1997 and recentlyin April 1999 and November 2000 amongseabass. Korea RO suspected VER occurrencefor whole year of 1999 and half year of 2000(OIE 1999, OIE 2000b).F.6.2 Clinical AspectsVER affects the nervous system. All affectedspecies show abnormal swimming behaviour(cork-screwing, whirling, darting and belly-upmotion) accompanied by variable swim bladderhyperinflation, cessation of feeding, changes incolouration, and mortality (Fig.F.6.2). Differencesbetween species are most apparent with relationto age of onset and clinical severity. Earlierclinical onset is associated with greater mortality,thus onset at one day post-hatch in striped jackresults in more severe losses than suffered byturbot, where onset is up to three weeks posthatch.Mortalities range from 10-100%.Two forms of VER have been induced withexperimental challenges (Peducasse et al.1999):i) acute – induced by intramuscular inoculation,andii) sub-acute – by intraperitoneal inoculation, bath,cohabitation and oral routes.F.6.3 Screening MethodsMore detailed information on screening methodsfor VER can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE 2000b),at http://www.oie.int or selected references.F.6.3.1 PresumptiveThere are no obvious diagnostic lesions that canbe detected in sub-clinical carriers.F.6.3.2 ConfirmatoryF.6.3.2.1 Virology (Level III)The nodavirus from barramundi has beencultured on a striped snakehead (Channastriatus) cell line (SSN-1) (Frerichs et al. 1996).The applicability of this cell line to othernodaviruses in this group is unknown.F.6.3.2.2 Nucleic Acid Assays (Level III)A newly developed polymerase chain reaction(PCR) method has shown potential for screeningpotential carrier striped jack and other fishspecies (O. fasciatus, E. akaara, T. rubripes, P.olivaceus, E. moara, O. punctatus and D. labrax).F.6.4 Diagnostic MethodsMore detailed information on diagnosticmethods for VER can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000b), at http://www.oie.int or selectedreferences.72

F.6 Viral Encephalopathy andRetinopathy (VER)(J Yulin)(S Chi Chi)Fig.F.6.2. Fish mortalities caused by VER.F.6.4.1PresumptiveF.6.4.1.1 Gross ObservationsAbnormal swimming behaviour and swim-bladderinflation in post-hatch larvae and juvenilestages of the host. Species described above,along with associated mortalities are indicativeof VER. Different species show different grossclinical signs (Table F.6.4.1.1). Non-feeding,wasting and colour changes in association withbehavioural abnormalities, should also be consideredsuspect.F.6.4.1.2 Histopathology (Level II)Normal histological methods may reveal varyingdegrees of vacuolisation in the brain or retinaltissues (Fig.F.6.4.1.2a and Fig.F.6.4.1.2b). Smalllarvae can be embedded whole in paraffinblocks and serially sectioned to providesections of brain and eyeballs. Larger fish(juvenile) usually require removal and fixationof eyes and brain.All the diseases described/named under F.6.1.1demonstrate vacuolisation of the brain, althoughsome species (e.g., shi drum, Umbrina cirrosa)may show fewer, obvious, vacuolar lesions. Inaddition, vacuolisation of the nuclear layers ofthe retina may not be present in Japaneseparrotfish or turbot. Intracytoplasmic inclusions(≤ 5 µm diameter) have been described insections of European sea bass and Australianbarramundi, Japanese parrotfish and brownspottedgrouper nerve tissue. Neuronal necrosishas been described in most species.Vacuolisation of the gut is not caused by VERnodaviruses, but is typical.Figs.F.6.4.1.2a, b. Vacuolation in brain (Br) andretina (Re) of GNNV-infected grouper in ChineseTaipei (bar = 100 mm).F.6.4.2 ConfirmatoryF.6.4.2.1 Virology (Level III)As described under F.6.3.2.1.F.6.4.2.2 Immunoassays (Level III)Immunohistochemistry protocols for tissue sectionsfixed in Bouin’s or 10% buffered formalinand direct fluorescent antibody test (DFAT) techniquesuse antibodies sufficiently broad in specificityto be able to detect at least four other virusesin this group. An ELISA test is only applicableto SJNNV from diseased larvae of stripedjack.F.6.4.2.3 Transmission Electron Microscopy(TEM) (Level III)Virus particles are found in affected brain andretina by both TEM and negative staining.Positive stain TEM reveals non-enveloped,icosahedral, virus particles associated withvacuolated cells and inclusion bodies. Theparticles vary from 22-25 nm (European seabass) to 34 nm (Japanese parrotfish) and formintracytoplasmic crystalline arrays, aggregatesor single particles (both intra- and extracellular).In negative stain preparations, non-enveloped,73

F.6 Viral Encephalopathy andRetinopathy (VER)SpeciesBehaviourChangesAppearanceChangesOnset of Clinical SignsBarramundiUncoordinateddarting andPale colouration,anorexia andEarliest onset at 9 days post-hatch.Usual onset at15-18 days post-corkscrewswimming; off feedwastinghatchEuropean Sea BassWhirling swimming;off feedSwim-bladderhyperinflationEarliest onset at 10 days posthatch.Usual onset 25-40 dayspost-hatchJapanese ParrotfishSpiral swimmingDarkened colourFirst onset anywhere between 6-25 mm total lengthRed-spotted GrouperWhirling swimming-First onset at 14 days post-hatch(7-8 mm total length). Usual onsetat 9-10 mm total lengthBrown-spotted GrouperStriped Jack-Abnormal-Swim-bladder20-50 mm total length1-4 days post-hatchTurbotswimmingSpiral and/orhyperinflationDarkened colour< 21 days post-hatchlooping swimpattern, belly-up atrestTable F.6.4.1.1 – adapted from OIE (1997)round to icosahedral particles, 25-30 nm, aredetectable. These are consistent with VERnodaviruses.F.6.4.2.4 Nucleic Acid Assay (Level III)Reverse transcriptase PCR assays have beendeveloped for VER nodavirus detection andidentification.F.6.5 Modes of TransmissionVertical transmission of VER virus occurs instriped jack, and ovarian infection has beenreported in European sea bass. Other modes oftransmission have not been clearlydemonstrated, but horizontal passage fromjuvenile fish held at the same site, andcontamination of equipment cannot yet be ruledout. Experimental infections have been achievedin larval stripe jack and red-spotted grouper usingimmersion in water containing VER virus.Juvenile European sea bass have also beeninfected by inoculation with brain homogenatesfrom infected individuals.F.6.6 Control MeasuresControl of VNN in striped jack and other affectedspecies is complicated by the verticaltransmission of the virus(es). Strict hygiene inhatcheries may assist in controlling VNNinfection. Culling of detected carrier broodstockis one control option used for striped bass,however, there is some evidence that reducedhandling at spawning can reduce ovarianinfections and vertical transmission in somecarrier fish. Control of clinical disease in stripedbass using the following techniques has alsoshown some success:• no recycling of culture water• chemical disinfection of influent water and larvaltanks between batches, and• reduction of larval density from 15-30 larvae/litre to

F.6 Viral Encephalopathy andRetinopathy (VER)F.6.7 Selected ReferencesAnderson, I., C. Barlow, S. Fielder, D. Hallam,M. Heasman and M. Rimmer. 1993. Occurrenceof the picorna-like virus infecting barramundi.Austasia Aquacult. 7:42-44.Arimoto, M., J. Sato, K. Maruyama, G. Mimuraand I. Furusawa. 1996. Effect of chemical andphysical treatments on the inactivation ofstriped jack nervous necrosis virus (SJNNV).Aquac. 143:15-22.Boonyaratpalin, S., K. Supamattaya, J.Kasornchandra, and R.W. Hoffman.1996.Picorna-like virus associated with mortalityand a spongious encephalopathy in grouper,Epinephelus malabaricus. Dis. Aquat. Org. 26:75-80.Bovo, G., T. Nishizawa, C. Maltese, F.Borghesan, F. Mutinelli, F. Montesi, and S.De Mas. 1999. Viral encephalopathy andretinopathy of farmed marine fish species inItaly. Vir. Res. 63: 143-146.Chi, S.C., W.W. Hu, and B.L. Lo. 1999.Establishment and characterization of acontinuous cell line (GF-1) derived fromgrouper, Epinephelus coioides (Hamilton): acell line susceptible to grouper nervousnecrosis virus (GNNV). J. Fish Dis. 22: 172-182.Comps, M., M. Trindade, and C. Delsert. 1996.Investigation of fish encephalitis virus (FEV)expression in marine fishes using DIG-labelledprobes. Aquac. 143:113-121.jack Pseudocaranx dentex selected asspawners. J. Aquat. Anim. Health 8: 332-334.OIE. 1997. OIE Diagnostic Manual for AquaticAnimal Diseases. Second Edition.OfficeInternational des Epizooties, Paris, France.252p.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Peducasse, S., J. Castric, R. Thiery, J.Jeffroy,A. Le Ven, and F. Baudin-Laurencin,. 1999.Comparative study of viral encephalopathy andretinopathy in juvenile sea bass Dicentrarchuslabrax infected in different ways. Dis. Aquat.Org. 36: 11-20.Thiery, R., R.C. Raymond, and J. Castric. 1999.Natural outbreak of viral encephalopathy andretinopathy in juvenile sea bass,Dicentrarchus labrax: study by nested reversetranscriptase-polymerase chainreaction. Vir. Res. 63: 11-17.Frerichs, G.N., H.D. Rodger, and Z. Peric.1996.Cell culture isolation of piscine neuropathynodavirus from juvenile sea bass,Dicentrarchus labrax. J. Gen. Vir. 77: 2067-2071.Munday, B.L. and T. Nakai. 1997. Special topicreview: Nodaviruses as pathogens in larvaland juvenile marine finfish. World J. Microbiol.Biotechnol. 13:375-381.Nguyen, H.D., K. Mushiake, T. Nakai, and K.Muroga. 1997. Tissue distribution of stripedjack nervous necrosis virus (SJNNV) in adultstriped jack. Dis. Aquat. Org. 28: 87-91.Nishizawa, T., K. Muroga, K. and M.Arimoto.1996. Failure of polymerase chainReaction (PCR) method to detect striped jacknervous necrosis virus (SJNNV) in Striped75

F.7 SPRING VIRAEMIA OF CARP(SVC)F.7.1 Background InformationF.7.1.1 Causative AgentsSpring viraemia of carp (SVC) is caused byssRNA Vesiculovirus (Rhabdoviridae), known asSpring viraemia of carp Virus (SVCV) or Rhabdoviruscarpio (RVC) (Fijan 1999). More detailedinformation about the disease can be found inthe OIE Diagnostic Manual for Aquatic AnimalDiseases (OIE 2000a).F.7.1.2 Host RangeSVCV infects several carp and cyprinid species,including common carp (Cyprinus carpio), grasscarp (Ctenopharyngodon idellus), silver carp(Hypophthalmichthys molitrix), bighead carp(Aristichthys nobilis), crucian carp (Carassiuscarassius), goldfish (C. auratus), tench (Tincatinca) and sheatfish (Silurus glanis).F.7.1.3 Geographic DistributionSVC is currently limited to the parts of continentalEurope that experience low water temperaturesover winter.F.7.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System(1999 – 2000)No reported case in any country during reportingperiod for 1999 and 2000 (OIE 1999, 2000b).F.7.2 Clinical AspectsYoung carp and other susceptible cyprinids(F.7.1.2), up to 1 year old, are most severelyaffected. Overt infections are manifest in springwhen water temperatures reach 11-17 °C. Poorphysical condition of overwintering fish appearsto be a significant contributing factor. Mortalitiesrange from 30-70%.Viral multiplication in the endothelial cells of bloodcapillaries, haematopoietic tissue and nephroncells, results in oedema and haemorrhage andimpairs tissue osmoregulation. Kidney, spleen,gill and brain are the organs in which SVCV ismost abundant during overt infection. Survivorsdemonstrate a strong protective immunity,associated with circulating antibodies, however,this results in a covert carrier state.F.7.3 Screening MethodsMore detailed information on methods forscreening SVC can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000a), at http://www.oie.int or selected references.F.7.3.1 PresumptiveThere are no methods for detection of sub-clinicalinfections using gross observations or routinehistology.F.7.3.1.1 Virology (Level III)Screening for sub-clinical carriers uses tissuehomogenates from the brain of any size fish orthe ovarian fluids from suspect broodstock fish.Cell lines susceptible to SVCV are EPC andFHM. Any resultant CPE requires molecularbasedassays as described under F.7.3.2.F.7.3.2 ConfirmatoryF.7.3.2.1 Immunoassays (Level III)CPE products can be checked for SVCV using avirus neutralisation (VN) test, indirect fluorescentantibody tests (IFAT), and ELISA. IFAT can alsobe used on direct tissue preparations.F.7.4 Diagnostic MethodsMore detailed information on methods fordiagnosis of SVC can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000a), at http://www.oie.intor selected references.F.7.4.1 PresumptiveF.7.4.1.1 Gross Observations (Level I)Sudden mortalities may occur with no otherclinical signs. Behavioural clues are non-specificto SVC and include lethargy, separation from theshoal, gathering at water inlets or the sides ofponds and apparent loss of equilibrium.External signs of infection are also non-specific,with fish showing varying degrees of abdominaldistension (dropsy), protruding vents and trailingmucoid faecal casts. Haemorrhaging at the basesof the fins and vent, bulging eye(s) (pop-eye orexophthalmia), overall darkening and pale gillsmay also be present (Figs.F.7.4.1.1a, b, c andd).Internal macroscopic signs of infection includean accumulation of body cavity fluids (ascites)76

F.7 Spring Viraemia of Carp (SVC)which may lead to the dropsy visible asabdominal distension, bloody and mucousfilledintestines, swim-bladder haemorrhageand gill degeneration.F.7.4.1.2 Transmission Electron Microscopy(TEM) (Level III)Detection of enveloped, bullet-shaped, viralparticles measuring 90-180 nm in length and witha regular array of spicules on the surface inspleen, kidney and brain tissues, or in isolatesfrom CPE in the cell-lines described underF.7.4.1.3, should be considered indicative of SVCin susceptible carp species showing other clinicalsigns of the disease. Viral replication takes placein the cytoplasm with maturation in associationwith the plasma membrane and Golgi vesicles.F.7.4.1.3 Virology (Level III)Whole fish (body length ≤4 cm), or visceraincluding kidney (fish 4 - 6 cm in length) or kidney,spleen and brain of larger fish, can be preparedfor tissue culture using Epithelioma papulosumcyprinae (EPC) or FTM cell lines. Resultant CPEshould be examined using the diagnostictechniques outlined below and under F.7.3.2.1to confirm SVCV as the cause.F.7.4.2ConfirmatoryF.7.4.2.1 Immunoassays (Level III)As described under F.7.4.1.3, SVCV can beconfirmed in CPE products using a virusneutralisation (VN) test, indirect fluorescentantibody tests (IFAT), and ELISA. IFAT can alsobe used on direct tissue preparations.F.7.4.2.2 Nucleic Acid Assay (Level III)RT-PCR techniques are under development.F.7.5 Modes of TransmissionHorizontal transmission can be direct (contactwith virus shed into the water by faeces, urine,reproductive fluids and, probably, skin mucous)or indirectly via vectors (fish-eating birds, the carplouse Argulus foliaceus or the leech Piscicolageometra). Vertical transmission is also possiblevia SVCV in the ovarian fluids (however, the rarityof SVC in fry and fingerling carp indicatesthat this may be a minor transmission pathway).SVCV is hardy and can retain infectivity afterexposure to mud at 4°C for 42 days, streamwater at 10°C for 14 days, and after drying at4-21°C for 21 days. This means that avenuesfor establishing and maintaining reservoirs ofinfection are relatively unrestricted. This, plusthe broad direct and indirect mechanisms fortransmission, makes this disease highly contagiousand difficult to control.F.7.6 Control MeasuresNo treatments are currently available althoughsome vaccines have been developed. Most effortis applied to optimising the overwinteringcondition of the fish by reducing stocking density,reduced handling and strict maintenance ofhygiene. New stocks are quarantined for at leasttwo weeks before release into ponds for growout.Control of spread means rapid removal anddestruction of infected and contaminated fishimmediately on detection of SVC. Repeatoutbreaks may allow action based onpresumptive diagnosis. First time outbreaksshould undertake complete isolation of affectedfish until SVC can be confirmed.F.7.7 Selected ReferencesDixon, P.F., A.M. Hattenberger-Baudouy, andK. Way. 1994. Detection of carp antibodies tospring viraemia of carp virus by competitiveimmunoassay. Dis. Aquat. Org. 19: 181-186.Fijan, N. 1999. Spring viraemia of carp and otherviral diseases and agents of warm-water fish,pp 177-244. In: Woo, P.T.K and Bruno, D.W.(eds).Fish Diseases and Disorders. Vol 3.Viral, Bacterial and Fungal Infections. CABIPublishing, Oxon, UK.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Oreshkova, S.F., I.S. Shchelkunov, N.V.Tikunova, T.I. Shchelkunova, A.T. Puzyrev,and A.A. Ilyichev. 1999. Detection of spring77

F.7 Spring Viraemia of Carp (SVC)viraemia of carp virus isolates byhybridisation with non-radioactive probesand amplification by polymerase chainreaction. Vir. Res. 63: 3-10.(EAFP)Rodak, L., Z. Pospisil, J. Tomanek, T. Vesley, T.Obr, and L. Valicek. 1993. Enzyme-linkedimmunosorbent assay (ELISA) for thedetection of spring viraemia of carp virus(SVCV) in tissue homogenates of the carpCyprinus carpio L. J. Fish Dis. 16: 101-111.Schlotfeldt, H.-J. and D.J. Alderman. 1995. WhatShould I Do? A Practical Guide for the FreshwaterFish Farmer. Suppl.Bull. Eur. Assoc. FishPathol. 15(4). 60p.Fig.F.8.4.1.1. Non-specific internal sign(petechial haemorrhage on muscle) of VHSinfected fish.(EAFP)abcdFigs.F.7.4.1.1a, b, c, d. Non-specific clinical signs of SVC infected fish, which may include swollenabdomen, haemorrhages on the skin, abdominal fat tissue, swim bladder and other.78

F.8 VIRAL HAEMORRHAGICSEPTICAEMIA (VHS)F.8.1 Background InformationF.8.1.1 Causative AgentViral haemorrhagic septicaemia (VHS) is causedby ssRNA enveloped rhabdovirus, known as viralhaemorrhagic septicaemia virus (VHSV).VHSV is synonymous with Egtved virus. Althoughpreviously considered to fall within theLyssavirus genus (Rabies virus), the ICTV haveremoved it to “unassigned” status, pending evaluationof a proposed new genus – Novirhabdovirus– to include VHSV and IHNV (see http://www.ncbi.nlm.nih.gov/ICTV). Several strains ofVHSV are recognised. More detailed informationabout the disease can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000a).F.8.1.2 Host RangeVHS has been reported from rainbow trout(Oncorhynchus mykiss), brown trout (Salmotrutta), grayling (Thymallus thymallus), white fish(Coregonus spp.), pike (Esox lucius) and turbot(Scophthalmus maximus). Genetically distinctstrains of VHSV have also been associated withdisease in Pacific salmon (Oncorhynchus spp.),Pacific cod (Gadus macrocephalus) and Pacificherring (Clupea pallasi). These strains show littlevirulence in rainbow trout challenges (OIE2000a). VHSV has also been isolated from Atlanticcod (Gadus morhua), European sea bass(Dicentrarchus labrax), haddock (Melanogrammusaeglefinus), rockling (Rhinonemuscimbrius), sprat (Sprattus sprattus), herring(Clupea harengus), Norway pout (Trisopterusesmarkii), blue whiting (Micromesistiuspoutassou), whiting (Merlangius merlangius) andlesser argentine (Argentina sphyraena)(Mortensen 1999), as well as turbot(Scophthalmus maximus) (Stone et al. 1997).Among each species, there is a high degree ofvariability in susceptibility with younger fish showingmore overt pathology.F.8.1.3 Geographic DistributionVHSV is found in continental Europe, the AtlanticOcean and Baltic Sea. Although VHSV-likeinfections are emerging in wild marine fish inNorth America, VHS continues to be considereda European-based disease, until the phylogeneticidentities of the VHSV-like viruseswhich do not cause pathology in rainbow troutcan be clearly established.F.8.1.4 Asia-Pacific Quarterly Aquatic Animal Disease Reporting System(1999-2000)Japan reported the disease during second quarterof 2000, no other reports from other countries(OIE 1999, 2000b).F.8.2 Clinical AspectsThe virus infects blood cells (leucocytes), theendothelial cells of the blood capillaries,haematopoietic cells of the spleen, heart, nephroncells of the kidney, parenchyma of the brainand the pillar cells of the gills. Spread of the viruscauses haemorrhage and impairment of osmoregulation.This is particularly severe in juvenilefish, especially during periods when water temperaturesranging between 4 – 14°C.F.8.3 Screening MethodsMore detailed information on methods for screeningVHS can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE 2000a),at http://www.oie.int or selected references.F.8.3.1 PresumptiveF.8.3.1.1 Gross Observations (Level I)and Histopathology (Level II)There are no gross visible clues (Level I) or histopathologyclues (Level II) to allow presumptivediagnosis of sub-clinical VHS infections. Subclinicalcarriers should be suspected, however,in populations or stocks which originate fromsurvivors of clinical infections or from confirmedcarrier broodstock.F.8.3.1.2 Virology (Level III)VHSV can be isolated from sub-clinical fish onBluegill Fry (BF-2), Epithelioma papulosumcyprinae (EPC) or rainbow trout gonad (RTG-2).Any resultant CPE requires further immunoassayor nucleic acid assay to confirm VHSV asthe cause (F.8.3.2).F.8.3.1.3 Immunoassay (Level III)Immunohistochemistry can be used to highlightVHSV in histological tissue samples (which ontheir own cannot be used to screen sub-clinicalinfections). Due to the wide range of hosts andserotypes, however, any cross-reactions needto be confirmed via tissue culture and subsequentviral isolation as described under F.8.3.1.2.79

F.8 Viral Haemorrhagic Septicaemia(VHS)F.8.3.2 ConfirmatoryF.8.3.2.1 Immunoassay (Level III)Identification of VHSV from cell-line culture canbe achieved using a virus neutralisation test,indirect fluorescent antibody test (IFAT) or ELISA.F.8.3.2.2 Nucleic Acid Assay (Level III)RT-PCR techniques have been developed.F.8.4 Diagnostic ProceduresMore detailed information on methods for diagnosisof VHS can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE 2000a),at http://www.oie.int or selected references.F.8.4.1 PresumptiveF.8.4.1.1 Gross Observation (Level I)There are no VHS-specific gross clinical signs.General signs are shared with bacterialsepticaemias, IHN, osmotic stress, handlingtrauma, etc., and include increased mortality,lethargy, separation from the shoal, gatheringaround the sides of ponds, nets or water inlets.The skin may become darkened andhaemorrhagic patches may be visible at the caseof the fins, the vent and over the body surface.Gill may also be pale. Internal organ changes mayor may not be present depending on the speedof onset of mortalities (stressed fish die quicker).Where present these include an accumulation ofbloody body cavity fluids (ascites), mucous-filledintestines and pale rectal tissues. Pin-pointhaemorrhages may also be present throughoutthe muscle (Fig.F.8.4.1.1), fat (adipose) tissueand swim-bladder.F.8.4.1.2 Virology (Level III)VHSV can be isolated from whole alevin (bodylength ≤ 4 cm), viscera including kidney (fish 4– 6 cm in length) or kidney, spleen and braintissue samples from larger fish, using BF-2, EPCor RTG-2 (as described under F.8.3.1.2). Anyresultant CPE requires further immunoassay ornucleic acid assay to confirm VHSV as the cause(F.8.3.2.1/2).F.8.4.1.3 Immunoassay (Level III)Immunohistochemistry can be used to highlightVHSV in histopathological lesions (however, histologyis not a normal method of diagnosingVHS). Due to the wide range of hosts and serotypes,however, any cross-reactions need to beconfirmed via tissue culture and subsequent viralisolation as described under F.8.3.1.2.F.8.4.2 ConfirmatoryAs described under F.8.3.2.F.8.5 Modes of TransmissionVHSV is shed in the faeces, urine and sexualfluids of clinically infected and sub-clinical carrierfish (wild and cultured). Once established at asite or in a water catchment system, the diseasebecomes enzootic because of the virus carrierfish. Water-borne VHSV can be carried 10-26 kmdownstream and remain infective. Mechanicaltransfer by fish-eating birds, transport equipmentand non-disinfected eggs from infectedbroodstock, have all been demonstrated as viableroutes of transmission (Olesen 1998).F.8.6 Control MeasuresNo treatments are currently available, althoughDNA-based vaccines have shown some successunder experimental conditions. Most controlmethods aim towards breaking the transmissioncycle and exposure to carriers, as well as reducingstress. Pathogenic proliferation occurs at temperatures

F.8 Viral Haemorrhagic Septicaemia(VHS)Heppell, J., N. Lorenzen, N.K. Armstrong, T. Wu,E. Lorenzen, K. Einer-Jensen, J. Schorr, andH.L. Davis. 1998. Development of DNA vaccinesfor fish: Vector design, intramuscularinjection and antigen expression using viralhaemorrhagic septicaemia virus genes as amodel. Fish and Shellf. Immunol. 8: 271-286.Lorenzen, N., E. Lorenzen, K. Einer-Jensen, J.Heppell, T. Wu, and H. Davis. 1998. Protectiveimmunity to VHS in rainbow trout(Oncorhynchus mykiss, Walbaum) followingDNA vaccination. Fish and Shellf. Immunol. 8:261-270.Mortensen, H.F. 1999. Isolation of viralhaemorrhagic septicaemia virus (VHSV) fromwild marine fish species in the Baltic Sea,Kattegat, Skagerrak and the North Sea. Vir.Res. 63: 95-106.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Olesen, N.J. 1998. Sanitation of viralhaemorrhagic septicaemia (VHS). J. Appl.Ichth. 14: 173-177.Schlotfeldt, H.-J. and D.J. Alderman. 1995. WhatShould I Do? A Practical Guide for the FreshwaterFish Farmer. Suppl. Bull. Eur. Assoc.Fish Pathol. 15(4). 60p.Stone, D.M., K. Way, and P.F. Dixon. 1997.Nucleotide sequence of the glycoprotein geneof viral haemorrhagic septicaemia (VHS) virusesfrom different geographical areas: A linkbetween VHS in farmed fish species and virusesisolated from North Sea cod (Gadusmorhua L.). J. Gen. Vir. 78: 1319-1326.81

F.9 LYMPHOCYSTISF.9.1 Background InformationF.9.1.1 Causative AgentLymphocystis is caused by dsDNA, non-enveloped,iridoviruses, with particles measuring200±50 nm, making them among the largest ofthe Iridoviridae. The iridovirus associated with giltheadsea bream (Sparus aurata) is known asLymphocystis Disease Virus (LDV).F.9.1.2 Host RangeLymphocystis occurs in many marine and somefreshwater fish families, including, herring(Clupeidae), smelt (Osmeridae), sea bass(Serranidae), flounder (Paralichthidae), snappers(Lutjanidae), perch (Percidae), drum(Sciaenidae), butterfly fishes (Chaetodontidae),cichlids (Cichlidae), gobies (Gobiidae) and sole(Soleidae).F.9.1.3 Geographic DistributionThe geographic range of lymphocystis is probablyglobal. The disease has been reported fromEurope, North and Central America, Australia,Africa, Hawaii, the South Pacific and Asia.F.9.2 Clinical AspectsLymphocystis is a common chronic and benigninfection by an iridovirus that results in uniquelyhypertrophied cells, typically in the skin and finsof fishes. The main clinical signs are white(occasionally pale red), paraffin-like nodulescovering the skin and fins of sick fish (Fig.F.9.2a). Some particulate inclusions maybe observed in the lymphoma lesion.At maturity the lesions are irregularly elevatedmasses of pebbled texture. The colour is lightcream to grayish, but covering epithelial tissuemay be normally pigmented. Vascularitysometimes gives large clusters of cells a reddishhue. Considerable variation occurs in size,location, and distribution of the masses. Infectedcells may also occur singly.Although the infection is rarely associated withovert disease, mortalities can occur under cultureconditions, possibly due to impaired gill,swimming or feeding capability with mechanicallyintrusive lesions. The primary effect, however, iseconomical, as fish with such grossly visible lesionsare difficult to market.F.9.3 Screening MethodsCurrently there are no detection techniques thatare sensitive enough to detect or isolate thisgroup of iridoviruses from sub-clinically infectedfish. To date, cell-culture techniques have beenlimited to isolation of virus from evident lymphomalesions.F.9.4Diagnostic MethodsF.9.4.1 PresumptiveF.9.4.1.1 Gross Observations (Level I)The main external signs associated withlymphocystis are white (or occasionally palepink), paraffin wax-like nodules or growths overthe skin and fins. Such growths may containsmall granular-like particles, and some mayshow signs of vascularisation (extension ofblood capillaries into the tissue growth) (Fig.F.9.4.1.1a and Fig. F.9.4.1.1b). The presence ofthe granular inclusions is an important for distinguishinglymphocystis from Carp Pox disease(caused by a Herpesvirus) (Fig.F.9.4.1.1c).The wax-like appearance is also an importantfeature which distinguishes Lymphocystis fromfungal (mycotic) skin growths (Fig.F.9.4.1.1d).F.9.4.2 ConfirmatoryF.9.4.2.1 Histopathology (Level II)Light microscopy of tissue sections of lymphomareveal that the particulate inclusions arevirus-induced giant cells of connective tissueorigin, enveloped with a thick capsule. The diameterof each giant cell is about 500 Îm, amagnification of normal cell volume of 50,000to 100,000-fold (Fig.F.9.4.2.1a). The distinctivecapsule (Fig.F.9.4.2.1b), enlarged and centrallylocated nucleus and nucleolus, and cytoplasmicinclusions, are unique. No such cell alterationsoccur with Carp Pox Herpesvirus infections.In addition, there may be some eosinophilic,reticulate (branching) inclusion bodies incytoplasm of the giant cell. These correspondto the viral replicating bodies which have a lightrefractive density which renders them “cytoplasmic-like”and demonstrate one of the fewinstances when a viral aetiology can be diagnosedat the light microscope level with a highdegree of confidence. Identification of the exactvirus(es) involved requires further investigation,however, this level of diagnosis is sufficientto allow control advice to be made (F.9.6).82

F.9 Lymphocystis(MG Bondad-Reantaso)(J Yulin)Fig.F.9.2a. Wild snakehead infected withlymphocystis showing irregularly elevatedmasses of pebbled structure.(J Yulin)Fig.F.9.4.1.1d. Goldfish with fungal (mycotic)skin lesions(J Yulin)Fig.F.9.4.1.1a. Flounder with severelymphocystis.(J Yulin)Fig.F.9.4.2.1a. Giant (hypertrophied lymphomacells with reticulate or branching inclusion bodiesaround the nuclei.(J Yulin)Fig.F.9.4.1.1b. Lymphocystis lesions showinggranular particle inclusions.(J Yulin)Fig.F.9.4.2.1b. Impression smear oflymphocystis showing some giant cells, andhgaline capsules (membrane).Fig.F.9.4.1.1c. Carp Pox Disease caused byHerpesvirus.83

F.9 LymphocystisF.9.4.2.2 Transmission Electron Microscopy(TEM) (Level III)TEM of ultra-thin section of lymphocystis tissueis the principal method of further confirming grossand light microscope observations. The viral particlesshow up as large icosahedral (roughly hexagonal-spherical)particles measuring 150-300nm within the encapsulated cell cytoplasm. Ultrastructuralfeatures of lymphocystis iridovirusparticles include a dense core within two unitmembranes making up the capsid(Fig.F.9.4.2.2aand Fig.F.9.4.2.2b). Note the difference fromHerpesvirus in Carp Pox, which are envelopedand smaller virions (Fig.F.9.4.2.2c).F.9.4.2.3 Virology (Level III)Due to the relative ease of finding and identifyingthe viruses associated with lymphocystis lesions(compared with the viral agents of other fish diseases),there has been little emphasis on cellculture as a means of confirming diagnosis ofthe disease. However, the growing impact of thisdisease in aquaculture situations around theworld has increased interest in differentiatingbetween the iridoviral agents involved and enhancingapparent acquired immunity to infection.A new cell-line from gilt-head sea bream is currentlyunder investigation and has shown promisefor isolating lymphocystis iridoviruses.F.9.5 Modes of TransmissionHorizontal contact and water-borne transmissionappear to be the principal mechanism forlymphocystis virus spread. This is reinforced byproliferation of the problem under intensive cultureconditions. High population density and externaltrauma enhance transmission. External surfacesincluding the gills appear to be the chief portal ofepidermal entry. The oral route seems not to beinvolved, and there is no evidence of verticaltransmission.(J Yulin)Fig.F.9.4.2.2a. Electron micrograph showingnumerous viral particles in cytoplasm.(J Yulin)Fig.F.9.4.2.2b. Enlarged viral particles showingtypical morphology of iridovirus (100 mm =bar).(J Yulin)F.9.6 Control MethodsAt present, there is no known method of therapyor of immunization. There is some evidence ofantibodies in at least one flatfish species, however,this remains to be investigated further.Avoidance of stocking with clinically infected fish,early detection through monitoring and sterile(land-fill or chemical) disposal, along withminimising stocking densities and handling skintrauma,have proven to be effective controls.Fig.F.9.4.2.2c. Herpervirus in Carp Pox showingenveloped and smaller virions compared tolymphocystis virus.84

F.9 LymphocystisF.9.7 Selected ReferencesBowden, R.A., D.J. Oestmann, D. H. Lewis, andM.S. Frey. 1995. Lymphocystis in red drum.J. Aquat. Anim. Health 7: 231-235.Bowser, P.R., G.A. Wooster, and R.G. Getchell.1999. Transmission of walleye dermal sarcomaand lymphocystis via waterborne exposure.J. Aquat. Anim. Health 11: 158-161.Yulin, J., Z. Chen, H. Liu, J. Pen, and Y. HuangY. 1999. Histopathological and electron microscopicobservation of Lymphocystic diseasevirus in flounder (Paralichthys), PP30. In: Bookof Abstracts. Fourth Symposium on Diseasesin Asian Aquaculture. Fish Health Section ofthe Asian Fisheries Society, Philippines.Chao, T.M. 1984. Studies on the transmissibility of lymphocystis disease occurring inseabass (Lates calcarifer Bloch). Sing. J.Prim. Ind. 12: 11-16.Dixon, P., D. Vethaak, D. Bucke, and M.Nicholson, M. 1996. Preliminary study of thedetection of antibodies to lymphocystis diseasevirus in flounder, Platichthys flesus L., exposedto contaminated harbour sludge. Fishand Shellf. Immunol. 6: 123-133.Garcia-Rosado, E., D. Castro, S. Rodriguez, S.I.Perez-Prieto, and J.J. Borrego. 1999. Isolationand characterization of lymphocystis virus(FLDV) from gilt-edged sea bream (Sparusaurata L.) using a new homologous cell line.Bul. Europ. Assoc. Fish Pathol. 19: 53-56.Limsuan, C., S. Chinabut and Y. Danayadol.1983. Lymphocystis disease in seabass (Latescalcarifer). National Inland Fisheries Institute,Fisheries Division, Department of Fisheries.Tech. Pap. No. 21. 6p. (In Thai, with Englishabstract).Perez-Prieto, S.I., S. Rodrigues-Saint-Jean, E.Garcia-Rosado, D. Castro, D., M..C. Alvarez,and J.J. Borrego. 1999. Virus susceptibility ofthe fish cell line SAF-1 derived from gilt-headseabream. Dis. Aquat. Org. 35:149-153.Williams, T. 1996. The iridoviruses. Adv. Vir.Res. 46: 345-412.Wolf, K. 1988. Fish viruses and fish viral diseases.Cornell University Press, Ithaca, NY.Xue, L., G. Wang, X. Xu, and M. Li. 1998. Preliminarystudy on lymphocystis disease of marinecage cultured Lateolabrax japonicus. MarineSciences/Haiyang Kexue. Qingdao No. 2:54-57.Yulin, J., Y. Li, and Z. Li. 1991. Electron microscopicobservation of pathogen of carp-poxdisease. Acta Hydrobiologica Sinica/Shuisheng Shengwu Xuebao.15: 193-195.85

BACTERIAL DISEASE OF FINFISHF.10 BACTERIAL KIDNEY DISEASE(BKD)F.10.1 Background InformationF.10.1.1 Causative AgentBacterial kidney disease (BKD) is caused byRenibacterium salmoninarum, a coryneform, rodshaped,Gram-positive bacterium that is the solespecies belonging to the genus Renibacterium.More detailed information about the diseases canbe found in the OIE Manual for Aquatic AnimalDiseases (OIE 2000a).F.10.1.2 Host RangeFish of the Salmonidae family are clinicallysusceptible, in particular the Oncorhynchusspecies (Pacific salmon and rainbow trout).F.10.1.3 Geographic DistributionBKD occurs in North America, Japan, WesternEurope and Chile.F.10.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System(1999-2000)Japan reported BKD occurrence for whole yearexcept for the month of December for both 1999and 2000 reporting period. Pakistan suspectedthe disease from July to December of 1999 (OIE1999, 2000b).F.10.2 Clinical AspectsRenibacterium salmoninarum infections can buildup over a long period of time, with clinical diseaseonly appearing in advanced infections, usuallywhen the fish have completed their first year oflife. Virulence of R. salmoninarum varies with:• the strain of bacterium• the salmon species infected• environmental and holding conditions.The bacteria can evade lysosomal breakdownby the blood cells that engulf them, thus avoiddestruction by the fishes’ primary defencemechanism. Nutrition and seawater transfers canalso affect the pathogenicity of R. salmoninaruminfections and broodstock infection levels arebelieved to have a direct correlation tosusceptibility in their offspring. Progeny of parentstock with low levels or no infection with R.salmoninarum show better survival than offspringfrom BKD compromised fish. This may reflectgreater transmission titres by the latter (F.10.5).F.10.3 Screening MethodsDetailed information on methods for screeningBKD can be found at the OIE Diagnostic Manualfor Aquatic Animal Diseases (OIE 2000a), at http://www.oie.int, or selected references.F.10.3.1 PresumptiveF.10.3.1.1 Gross Observations (Level I)and Histopathology (Level II)There are no gross signs or histological lesionsthat can be detected in sub-clinical carriers ofRenibacterium salmoninarum.F.10.3.1.2 Bacteriology (Level II)When no lesions are present, the kidney shouldbe selected for culture. In mature females,coelomic fluids may also be used. Specialisedgrowth media, such as, kidney disease mediumenriched with serum (KDM2) or charcoal(KDMC), or selective kidney disease medium(SKDM) are required due to the fastidious natureof Renibacterium salmoninarum.Growth requires 2-3 weeks, but may take up to12 weeks. Colonies are pinpoint to 2 mm indiameter, white-creamy, shiny, smooth, raisedand entire (Fig.F.10.3.1.2a). The rods(Fig.F.10.3.1.2b) are 0.3-1.5 x 0.1-1.0 mm,Gram-positive, PAS-positive, non-motile, nonacid-fast, frequently arranged in pairs or chainsor in pleiomorphic forms (“Chinese letters”). Oldcultures may achieve a granular or crystallineappearance. Transverse sections through suchcolonies will reveal the presence of Gram-positiverods in a crystalline matrix. Although few otherbacteria have these growth characteristics,identification of the bacteria should be confirmedby immunoassay (F.10.3.2.1) or nucleic acidassay (F.10.3.2.2).F.10.3.2 ConfirmatoryF.10.3.2.1Immunoassays (Level II/III)Agglutination tests, direct and indirectfluorescent antibody tests (DFAT, IFAT) andELISA kits are now available that can be usedto detect R. salmoninarum antigen in fishtissues, as well as from bacterial cultures. TheELISA tests are believed to be the mostsensitive to low-titre infections, hence they arerecommended for screening for sub-clinicalcarriers (such as ovarian fluids from broodstocksalmonids). Commercially produced kits are86

F.10 Bacterial Kidney Disease (BKD)(M Yoshimizu)(EAFP)Fig.F.10.3.1.2a. Pinpoint colonies up to 2 mmin diameter of Renibacteriium salmonimarum,white-creamy, shiny, smooth, raised and entire;three weeks after incubation at 15°C on KDM-2 medium.(M Yoshimizu)Fig.F.10.4.1.1b. Enlargement of spleen is alsoobserved from BKD infected fish.especially for sub-clinical cases, or first timeisolations (Griffiths et al. 1996).F.10.3.2.2 Nucleic Acid Assays (Level III)Renibacterium salmoninarum primers have beendeveloped for PCR-probes. These can detect R.salmoninarum DNA in tissue homogenates. Theprimers have been published and some kits arenow commercially available.F.10.4 Diagnostic MethodsDetailed information on methods for diagnosis ofBKD can be found at the OIE Diagnostic Manualfor Aquatic Animal Diseases (OIE 2000a), at http://www.oie.int, or selected references.Fig.F.10.3.1.2b. Renibacterium salmoninarumrods, isolated from masou salmon.(M Yoshimizu)Fig.F.10.4.1.1a. Kidney of masou salmonshowing swelling with irregular grayish.also available, which contain specificinstructions. Positive ELISA results, using eitherpolyclonal or monoclonal antibodies, should becorroborated with other diagnostic tests,F.10.4.1 PresumptiveF.10.4.1.1 Gross Observations (Level I)Gross clinical signs are not usually evident untilinfections have become well-advanced (usuallyafter at least 1 year). These include exophthalmia(pop-eye), varying degrees of abdominaldistension (dropsy) due to disruption of thekidney excretory function, skin lesions andhaemorrhaging.Internally, there is evidence of grey/white lesions(granulomas) in all the organs, but especially thekidney (Fig.F.10.4.1.1a); enlargement of spleen(Fig.F.10.4.1.1b) is also observed. The greyishspots may show signs of multiplication andcoalescence until the whole kidney appearsswollen and bloated with irregular greyishpatches. BKD can be distinguished fromproliferative kidney disease (PKD) insalmonids, where the kidney becomes enlargedbut there is no associated grey discolouration.Another salmonid kidney disease –nephrocalcinosis – only affects the urinary87

F.10 Bacterial Kidney Disease (BKD)ducts, which develop a white porcelain textureand colour.F.10.4.1.2 Smears (Level I)Smears from tissue lesions of susceptible hostsstained with Gram’s stain or other metachromaticstain may reveal large numbers of small Grampositive,rod-shaped, bacteria. Care should betaken not to confuse these with the melaningranules commonly present in kidney tissues.Other Gram-positive bacteria, such as Lacticspecies, may also be present, so furtherbacteriological identification methods arerequired.F.10.4.1.3 Bacteriology (Level II)Whenever possible, culturing should be used forconfirmation despite the difficulties imposed bythe slow, fastidious growth of Renibacteriumsalmoninarum. Presumptive diagnosis is alsopossible from bacterial culture due its slow growth(2-3 weeks) at 15°C. Kidney and other organswith suspicious lesions should be sampled. Protocolsfor culture are as described underF.10.3.1.2. Although few other bacteria havethese growth characteristics, identification of thebacteria should be confirmed by immunoassay(F.10.3.2.1) or nucleic acid assay (F.10.3.2.2)F.10.4.2 ConfirmatoryF.10.4.2.1 Immunoassay (Level II/III)Slide agglutination tests can be used for rapididentification of culture colonies. Bacterialagglutination is determined by comparison withduplicate suspension containing rabbit serum, asa control. Co-agglutination with Staphylococcusaureus (Cowan I strain) sensitised with specificimmunoglobulins is also effective at enhancingthe agglutination process (Kimura and Yoshimizu1981).For immunofluorescence (direct and indirect) andELISA tests, MAbs against specific determinantsare recommended to avoid cross-reactions withother bacteria. As noted under F.10.3.2.1, positiveresults, using either polyclonal or monoclonalantibodies, should be corroborated with otherdiagnostic tests, especially for first time isolations(Griffiths et al. 1996).F.10.4.2.2 Nucleic Acid Assays (Level III)As described under F.10.3.2.2, Renibacteriumsalmoninarum PCR-probes are now available.Cross-checking positive samples with other diagnosticmethods (bacteriology, immunoassay),however, is highly recommended, especially forfirst time isolations (Hiney and Smith 1999).F.10.5 Modes of TransmissionRenibacterium salmoninarum is widely distributedin both freshwater and marine environments. Itcan be transmitted horizontally by water-bornerelease and faecal contamination, as well as viareservoir hosts which span all salinity ranges.Indirect vertical transmission via reproductivefluids and spawning products is also possiblefor sub-clinical carriers of the bacteria.F.10.6 Control MeasuresDue to its intracellular location in the host fish,BKD is difficult to treat with antibiotics. Injectionof female broodstock with erythromycin at regularintervals prior to spawning appears to have somesuccess in preventing vertical transmission toeggs. Vaccination and medicated feeds have alsoshown some success in reducing the occurrenceof BKD, however, results have varied with strainof R. salmoninarum and host species.Most emphasis is placed on breaking vertical andhorizontal transmission routes (F.10.5). Cullingof high BKD-titre broodstock, reducing stockingdensity, avoiding contact with sub-clinicalcarriers/reservoirs, reducing handling stress andavoiding unacclimatised transfer from fresh tosaltwater, have all proven effective in reducingBKD pathogenicity.F.10.7 Selected ReferencesAustin, B., T.M. Embley, and M. Goodfellow.1983. Selective isolation of Renibacteriumsalmoninarum. FEMS Micro. Let. 17: 111-114.Brown, L.L., G.K. Iwama, T.P.T. Evelyn, W.S.Nelson, and R.P. Levine. 1994. Use of thepolymerase chain reaction (PCR) to detectDNA from Renibacterium salmoninarumwithin individual salmon eggs. Dis. Aquat.Org. 18: 165-171.Daly, J.G. and R.M.W. Stephenson. 1985.Charcoal agar, a new growth medium for thefish disease bacterium Renibacteriumsalmoninarum. Appl. Environ. Micro. 50: 868-871.Evelyn, T.P.T. 1977. An improved growthmedium for the kidney disease bacterium and88

F.10 Bacterial Kidney Disease (BKD)some notes on using the medium. Bull. ofthe OIE. 87: 511-513.Griffiths, S.G., K. Liska, and W.H. Lynch. 1996.Comparison of kidney tissueand ovarian fluidfrom broodstock Atlantic salmon for detectionof Renibacterium salmoninarum, and use ofSKDM broth culture with western blotting toincrease detection in ovarian fluid. Dis. Aquat.Org. 24: 3-9.Hiney, M.P. and P.R. Smith. 1999. Validation ofpolymerase chain reaction-based techniquesfor proxy detection of bacterial fish pathogens:Framework, problems and possible solutionsfor environmental applications. Aquac. 162:41-68.Kimura, T. and M. Yoshimizu. 1981. Acoagglutination test with antibody-sensitisedstaphylococci for rapid and simple diagnosisof bacterial kidney disease (BKD). Dev. Biol.Standard. 49: 135-148.Leon, G., M.A. Martinez, J.P. Etchegaray, M.I.Vera, Figueroa and M. Krauskopf. 1994.Specific DNA probes for the identification ofthe fish pathogen Renibacterium salmoninarum.Wor. J. Micro.Biotech. 10: 149-153.Meyers, T.J., S. Short, C. Farrington, K. Lipson,H.J. Geiger, and R. Gates. 1993. Establishmentof a positive-negative threshold opticaldensity value for the enzyme-linked immunosorbentassay (ELISA) to detect solubleantigen of Renibacterium salmoninarum inAlaskan Pacific salmon. Dis. Aquat. Org.16:191-197.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Schlotfeldt, H.-J. and D.J. Alderman. 1995.What Should I Do? A Practical Guide for theFreshwater Fish Farmer. Suppl. Bull. Eur.Assoc. Fish Pathol. 15(4). 60p.89

FUNGUS ASSOCIATED DISEASEF.11 EPIZOOTIC ULCERATIVESYNDROME (EUS)F.11.1Background InformationF.11.2 Clinical AspectsF.11.1.1 Causative FactorsThe mycotic granulomas in EUS-affected tissuesare caused by the Oomycete fungusAphanomyces invadans (also known as A.invaderis, A. piscicida, Mycotic Granuloma-fungus(MG) and ERA [EUS-relatedAphanomyces]). It is also known as Red spotdisease (RSD). More detailed information aboutthe disease can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE 2000a).F.11.1.2 Host RangeEUS affects freshwater and estuarine warm waterfish and was first reported in farmed ayu(Plecoglossus altivelis) in Japan (Fig.F.11.1.2a).Severe outbreaks occurred in Eastern Australiaaffecting estuarine fish, particularly grey mullet(Mugil cephalus). Region-wide, over 50 species(Fig.F.11.1.2b) have been confirmed affected byhistopathological diagnosis (Lilley et al., 1998),but some important culture species includingtilapia, milkfish and Chinese carps have beenshown to be resistant.F.11.1.3 Geographic DistributionEUS was first reported in Japan and subsequentlyin Australia. Outbreaks have shown awestward pattern of spread through Southeastand South Asia. EUS has also spread westwardwith major outbreaks reported in Papua NewGuinea, Malaysia, Indonesia, Thailand, Philippines,Sri Lanka, Bangladesh and India. EUS hasmost recently been confirmed in Pakistan. Thepathology demonstrated by ulcerative mycosis(UM)-affected estuarine fish along the Atlanticcoast of USA is indistinguishable from EUS, butfurther work is required to compare the causalagents involved in each case.F.11.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System(1999-2000)Australia, Bangladesh, India, Japan, Lao PDR,Nepal, Philippines, Sri Lanka, and Thailandreported the disease on various months for thereporting year 1999; for the year 2000, Australia,Bangladesh, India, Japan, Lao PDR, Nepal,Pakistan, Philippines and Thailand reportedpositive occurrence of EUS (OIE 1999, OIE2000b).Affected fish typically show necrotic dermal ulcers,characterised histologically by the presenceof distinctive mycotic granulomas in underlyingtissues. The mycotic granulomas inEUS-affected tissues are caused by theOomycete fungus Aphanomyces invadans. Initiallesions may appear as red spots(Fig.F.11.2a), which become deeper as the infectionprogresses and penetrate underlyingmusculature (Fig.F.11.2b). Some advanced lesionsmay have a raised whitish border. Highmortalities are usually associated with EUS outbreaksbut, in certain cases, where fish do notsuccumb to secondary invasion of these gapingwounds, ulcers may be resolved.F.11.3 Screening MethodsThere are no screening methods for sub-clinicalanimals available.F.11.4 Diagnostic MethodsMore detailed information on methods for diagnosisof EUS can be found in the OIE Manualfor Aquatic Animal Diseases (OIE 2000a), athttp://www.oie.int, or selected references..F.11.4.1 PresumptiveF.11.4.1.1Gross Observations (Level I)The gross appearance of lesions varies betweenspecies, habitat and stage of lesion development(Fig.F.11.4.1.1a). The most distinctive EUSlesion is the open dermal ulcer. However, otherdiseases may also result in similar clinical lesions(Fig.11.4.1.1b) and it is, therefore, importantto confirm the presence of A. invadans toensure accurate diagnosis.F.11.4.1.2Rapid Squash Muscle Preparation(Level I)Presumptive diagnosis of EUS in susceptiblefish showing dermal lesions can be made bydemonstrating aseptate hyphae (12-30 ?m indiameter) in squash preparations of the muscleunderlying the visible lesion (Fig.11.4.1.2). Thiscan be achieved using a thin piece of musclesquashed between two glass plates or microscopeslides, examined using a light or dissectingmicroscope under field conditions.90

F.11 Epizootic Ulcerative Syndrome(EUS)(K Hatai)(MG Bondad-Reantaso)Fig.F.11.1.2a. Ayu, Plecoglatus altivelis, infectedwith mycotic granulomatosis.(RB Callinan)Fig.F.11.2b. Snakehead in Philippines (1985)showing typical EUS lesions.(MG Bondad-Reantaso)Fig.F.11.1.2b. EUS affected farmed silver perchBidyanus bidyanus from Eastern Australia.(MG Bondad-Reantaso)Fig.F.11.4.1.1b. Red spot disease of grass carpin Vietnam showing ulcerative lesions.F.11.4.2 ConfirmatoryFig.F.11.2a. Catfish showing initial EUS redspots.(MG Bondad-Reantaso)Fig.F.11.4.1.1a. Wild mullet in Philippines (1989)with EUS.F.11.4.2.1 Histopathology (Level II)Confirmatory diagnosis requires histologicaldemonstration of typical granulomas and invasivehyphae using haematoxylin and eosin(Fig.F.11.4.2.1a) or a general fungus stain (e.g.Grocott’s) (Fig.F.11.4.2.1b). Early EUS lesionsshow shallow haemorrhagic dermatitis with noobvious fungal involvement. Later lesions demonstrateA. invadans hyphae penetrating the skeletalmuscle tissues and increasing inflammation.The fungus elicits a strong inflammatory responseand granulomas are formed around the penetratinghyphae, a typical characteristic of EUS. Thelesion progresses from a mild chronic dermatitis,to a severe, locally pervasive, necrotisingdermatitis, with severe degeneration of themuscle. The most typical lesions are large, open,haemorrhagic ulcers about 1-4 cm in diameter.These commonly show secondary infections withbacteria, and pathogenic strains of Aeromonashydrophila have been isolated from lesions.F.11.4.2.2 Mycology (Level II)Moderate, pale, raised, dermal lesions are mostsuitable for fungal isolation attempts. Remove91

F.11 Epizootic Ulcerative Syndrome(EUS)the scales around the periphery of the lesionand sear the underlying skin with a red-hotspatula to sterilise the surface. Using a sterilescalpel blade and sterile, fine pointed, forceps,cut through skin underlying the seared area andcut horizontally to lift the superficial tissues andexpose the underlying muscle. Ensure the instrumentsdo not contact the external surfaceand contaminate the underlying muscle. Asepticallyexcise 2 mm 3 pieces of muscle, approximately2 mm 3 , and place on a Petri dish containingCzapek Dox agar with penicillin G (100units/ml) and oxolinic acid (100 mg/ml). Sealplates and incubate at room temperature examiningdaily. Transfer emerging hyphal tipsonto fresh plates of Czapek Dox agar until culturesare free of contamination.The fungus can be identified to genus by inducingsporogenesis (Fig.F.11.4.2.2a) and demonstratingthe asexual characteristics ofAphanomyces as described in Lilley et al. (1998).A. invadans is characteristically slow growing inculture (Fig.F.11.4.2.2b) and fails to grow at 37 o Con GPY agar (GP broth with 0.5 g/l yeast extractand 12 g/l technical agar). Detailed temperaturegrowthprofiles are given in Lilley and Roberts(1997). Confirmation that the isolate is A.invadans can be made by injecting a 0.1 ml suspensionof 100+ motile zoospores intramuscularlyin EUS-susceptible fish (preferablyChanna striata) at 20 o C, and demonstrating histologicallygrowth of aseptate hyphae 12-30 µmin diameter in muscle of fish sampled after 7days, and typical mycotic granulomas in muscleof fish sampled after 14 days.F.11.5 Modes of TransmissionThe spread of EUS is thought to be due to floodingand movement of affected and/or carrierfish. Aphanomyces invadans is considered tobe the “necessary cause” of EUS, and ispresent in all cases, however, an initial skin lesionis required for the fungus to attach andinvade underlying tissues. This lesion may beinduced by biotic or abiotic factors. In Australiaand Philippines, outbreaks have been associatedwith acidified water (due to acid sulfatesoil runoff), along with low temperatures, presenceof susceptible fish and A. invadanspropagules. In other areas, where acid waterdoes not occur, it is possible that other biological(e.g., rhabdovirus infection) or environmentalfactors (e.g., temperature) may initiate lesions.F.11.6 Control MeasuresControl in wild populations is impossible in mostcases. Selection of resistant species for culturepurposes currently appears to be the mosteffective means of farm-level control. Wherechanging culture species is not an option, measuresshould be taken to eradicate or excludethe fungus through:• drying and liming of ponds prior to stocking• exclusion of wild fish• use of prophylactically-treated, hatcheryrearedfry• use of well-water• salt bath treatments• disinfection of contaminated nets and equipment.F.11.7 Selected ReferencesBlazer, V.S., W.K. Vogelbein, C.L. Densmore,E.B. May, J.H. Lilley and D.E. Zwerner. 1999.Aphanomyces as a cause of ulcerative skinlesions of menhaden from Chesapeake Baytributaries. J. Aquat. Anim. Health 11:340-349.Bondad-Reantaso, M,G., S.C. Lumanlan, J.M.Natividad and M.J. Phillips. 1992. Environmentalmonitoring of the epizootic ulcerativesyndrome (EUS) in fish from Munoz, NuevaEcija in the Philippines, pp. 475-490. In: Diseasesin Asian Aquaculture 1. M. Shariff, R.P.Subasinghe and J.R. Arthur (eds). Fish HealthSection, Asian Fisheries Society, Manila, Philippines.Callinan, R.B., J.O. Paclibare, M.G. Bondad-Reantaso, J.C. Chin and R.P. Gogolewsky.1995. Aphanomyces species associated withepizootic ulcerative syndrome (EUS) in thePhilippines and red spot disease (RSD) inAustralia: preliminary comparative studies. Dis.Aquat. Org. 21:233-238.Callinan, R.B., J.O. Paclibare, M.B. Reantaso,S.C. Lumanlan-Mayo, G.C. Fraser andJ.Sammut. 1995. EUS outbreaks in estuarinefish in Australia and the Philippines: associationswith acid sulphate soils, rainfall andAphanomyces, pp. 291-298. In:Diseases inAsian Aquaculture 1. M. Shariff, J.R. Arthurand R.P. Subasinghe (eds). Fish Health Section,Asian Fisheries Society, Manila, Philippines.92

F.11 Epizootic Ulcerative Syndrome(EUS)(MG Bondad-Reantaso)(K Hatai)Fig.F.11.4.1.2. Granuloma from squash preparationof muscle of EUS fish.Fig.F.11.4.2.2a.Typical characteristicof Aphanomycessporangium.(MG Bondad-Reantaso)(MG Bondad-Reantaso)Fig.F.11.4.2.1a. Typical severe mycotic granulomasfrom muscle section of EUS fish (H & E).(MG Bondad-Reantaso)Fig.F.11.4.2.2b. Growth of Aphanomycesinvadans on GP agar.Fig.F.11.4.2.1b. Mycotic granulomas showingfungal hyphae (stained black) using Grocottsstain.Chinabut, S. and R.J. Roberts. 1999. Pathologyand Histopathology ofEpizooticUlcerative Syndrome (EUS). AquaticAnimal Health Research Institute,Departmentof Fisheries, Royal Thai Government,Bangkok, Thailand, 33p. ISBN 974-7604-55-8.Fraser, G.C., R.B. Callinan, and L.M. Calder.1992. Aphanomyces species associatedwithred spot disease: an ulcerative disease of estuarinefish from eastern Australia. J. Fish Dis.15:173-181.Hatai, K. and S. Egusa. 1978. Studies on thepathogenic fungus of mycotic granulomatosis-II.Some of the note on the MG-fungus.Fish Pathol. 13(2):85-89 in Japanese, withEnglish abstract).Hatai, K., S. Egusa, S. Takahashi and K. Ooe.1977. Study on the pathogenic fungus ofmycotic granulomatosis-I. Isolation andpathogenicity of the fungus from cultured ayuinfected with the disease. Fish Pathol. 11(2):129-133.93

F.11 Epizootic Ulcerative Syndrome(EUS)Lilley, J.H., Callinan, R.B., Chinabut, S.,Kanchanakhan, S., MacRae, I.H., andPhillips,M.J. 1998. Epizootic Ulcerative Syndrome(EUS) Technical Handbook.TheAquatic Animal Health Research Institute,Bangkok. 88p.Lilley, J.H. and Roberts, R.J. 1997. Pathogenicityand culture studies comparingAphanomyces involved in epizootic ulcerativesyndrome (EUS) with other similar fungi. J.Fish Dis. 20: 135-144.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Roberts, R.J., B. Campbell and I.H. MacRae(eds). 1994. Proceedings of the RegionalSeminar on Epizootic Ulcerative Syndrome,25-27 January 1994. The Aquatic AnimalHealth Research Institute. Bangkok, Thailand.Tonguthai, K. 1985. A preliminary account of ulcerativefish diseases in the Indo-Pacific region(a comprehensive study based on Thaiexperiences). National Inland Fisheries Institute,Bangkok, Thailand. 39p.94

ANNEX F.AI. OIE REFERENCE LABORATORIESFOR FINFISH DISEASESDiseaseEpizootic haematopoietic necrosisvirus (EHNV)Infectious haematopoietic necrosisvirus (Rhabdoviruses)Onchorhynchus masou virusSpring viremia of carp virusViral haemorrhagic septicaemia virusExpert/LaboratoryDr. A. HyattAustralian Animal Health LaboratoryGeelong, Victoria 3213, AUSTRALIATel: 61-3-52275000Fax: 61-3-52275555E-mail: alex.hyatt@dah.csiro.auDr. R. WhittingtonElizabeth MacArthur Agricultural InstitutePMB 8, CamdenNSW 2570, AUSTRALIATel: 61-2-46293333Fax: 61-2-46293343E-mail: Richard.Whittington@smtpgwy.agric.nsw.gov.auDr. J. A. LeongOregon State UniversityDepartment of MicrobiologyNash Hall 220, Corvallis, Oregon 93331-3804UNITED STATES of AMERICATel: 1-541-7371834Fax: 1-541-7370496E-mail: leongj@orst.eduDr. J. WintonWestern Fisheries Research Center6505 N.E. 65th StreetSeattle, Washington 98115UNITED STATES of AMERICAE-mail: jim_winton@nbs.govDr. M. YoshimizuLaboratory of MicrobiologyFaculty of FisheriesHokkaido University3-1-1, Minato-cho, HakodateHokkaido 041-0821JAPANTel./Fax: 81-138-408810E-mail: yosimizu@pop.fish.hokudai.ac.jpDr. B.J. HillThe Centre for Environment, Fisheries and AquacultureSciences (CEFAS)Barack Road, the Nothe, Weymouth, DorsetDT4 8UB UNITED KINGDOMTel: 44-1305-206626Fax:44-1305-206627E-mail: b.j.hill@cefas.co.ukDr. N.J. OllesenDanish Veterinary LaboratoryHangovej 2, DK-8200 Aarhus NDENMARKTel: 45-89372431Fax:45-89372470E-mail: njo@svs.dk95

Annex F.AI. OIE Reference Laboratoriesfor Finfish DiseasesChannel catfish virusViral encephalopathy and retinopathyInfectious pancreatic necrosisInfectious salmon anaemiaEpizootic ulcerative syndromeBacterial kidney diseaseDr. L.A. HansonFish Diagnostic LaboratoryCollege of Veterinary MedicineMississippi State UniversityBox 9825, Spring StreetMississippi 39762UNITED STATES of AMERICATel: 1-662-3251202Fax: 1-662-3251031E-mail: hanson@cvm.msstate.eduDr. G. BovoInstituto Zooprofilaticco Sperimentale delle VenezieDipartimento di Ittiopatologia, Via Romea 14/A35020 Legnaro PD ITALYTel: 39-049-8830380Fax: 39-049-8830046E-mail: bovo.izs@interbusiness.itDr. T. NakaiFish Pathology LaboratoryFaculty of Applied Biological SciencesHiroshima UniversityHigashihiroshima 739-8528JAPANTel: 81-824-247947Fax: 81-824-227059E-mail: nakaitt@ipc.hiroshima-u.ac.jpDr. B.J. HillThe Centre for Environment, Fisheries and AquacultureSciences (CEFAS)Barack Road, the Nothe, Weymouth, DorsetDT4 8UB UNITED KINGDOMTel: 44-1305-206626Fax:44-1305-206627E-mail: b.j.hill@cefas.co.ukDr. B. DannevigNational Veterinary InstituteUllevalsveien 68P.O. Box 8156Dep., 0033 OsloNORWAYTel: 47-22-964663Fax: 47-22-600981E-mail: birgit.dannevig@vetinst.noDr. Kamonporn TonguthaiAquatic Animal Health Research InstituteDepartment of FisheriesKasetsart University CampusJatujak, Ladyao, Bangkok 10900THAILANDTel: 662-5794122Fax: 662-5613993E-mail: kamonpot@fisheries.go.thDr. R.J. PaschoWestern Fisheries Research CenterU.S. Geological SurveyBiological Resources Division6505 N.E. 65th StreetSeattle, Washington 9811596

Annex F.AI. OIE Reference Laboratoriesfor Finfish DiseasesEnteric septicaemia of catfishPiscirikettsiosisUNITED STATES of AMERICATel: 1-206-5266282Fax: 1-206-5266654E-mail: ron_pascho@usgs.govDr. L.A. HansonFish Diagnostic LaboratoryCollege of Veterinary MedicineMississippi State UniversityBox 9825, Spring StreetMississippi 39762UNITED STATES of AMERICATel: 1-662-3251202Fax: 1-662-3251031E-mail: hanson@cvm.msstate.eduDr. J.L. FryerDistinguished Professor EmeritusDepartment of Biology220 Nash HallOregon State UniversityCorvallis, Oregon 97331-3804Tel: 1-541-7374753Fax:1-541-7372166E-mail: fryerj@bcc.orst.eduGyrodactylosis (Gyrodactylus salaris) Dr. T. Atle MoNational Veterinary InstituteUllevalsvein 68P.O. Box 8156Dep., 0033 OsloNORWAYTel: 47-22-964722Fax: 47-22-463877E-mail: tor-atle.mo@vetinst.noRed sea bream iridoviral diseaseDr. K. NakajimaVirology Section, Fish Pathology DivisionNational Research Institute of AquacultureFisheries Agency422-1 Nakatsuhama, Nansei-choWatarai-gun Mie 516-0913JAPANTel: 81-599661830Fax: 81-599661962E-mail: kazuhiro@nria.affrc.go.jp97

ANNEX F.AII. LIST OF REGIONAL RESOURCEEXPERTS FOR FINFISH DISEASES INASIA PACIFIC 1DiseaseEpizootic ulcerativesyndrome (EUS)ExpertDr. Richard CallinanNSW Fisheries, Regional Veterinary LaboratoryWollongbar NSW 2477AUSTRALIATel (61) 2 6626 1294Mob 0427492027Fax (61) 2 6626 1276E-mail: richard.callinan@agric.nsw.gov.auDr. C.V. MohanDepartment of AquacultureCollege of Fisheries, UASMangalore-575002INDIATel: 91 824 439256 (College); 434356 (Dept), 439412(Res)Fax: 91 824 438366E-mail: cv_mohan@yahoo.comProf. Kishio HataiDivison of Fish DiseasesNippon Veterinary and Animal Science University1-7-1 Kyonan-cho, Musashino, Tokyo 180JAPANTel: 81-0422-31-4151Fax: 81-0422-33-2094E-mail: hatai@scan-net.ne.jpMs. Susan Lumanlan-MayoFish Health SectionBureau of Fisheries and Aquatic ResourcesArcadia Building, 860 Quezon AvenueQuezon City, Metro ManilaPHILIPPINESTel/Fax: 632-372-5055E-mail: slmayo99@yahoo.comMr. Jose O. PaclibareFish Health SectionBureau of Fisheries and Aquatic ResourcesArcadia Building, 860 Quezon AvenueQuezon City, Metro ManilaPHILIPPINESTel/Fax: 632-372-5055E-mail: jopac@edsamail.com.phDr. Erlinda LacierdaFish Health SectionAquaculture DepartmentSoutheast Asian Fisheries Development CenterTigbauan, Iloilo 5021PHILIPPINESTel: 63 33 335 1009Fax: 63 33 335 1008E-mail: eclacier@aqd.seafdec.org.phDr. Somkiat KanchanakhanAquatic Animal Health Research InstituteDepartment of FisheriesKasetsart University Campus1The experts included in this list have previously been consulted and agreed to provide valuable information and health adviseconcerning their particular expertise.98

Annex F.AII.List of Regional Resource Experts forFinfish Diseases in Asia PacificViral nervous necrosis (VNN)Viral encephalopathy andretinopathy (VER)Jatujak, Ladyao, Bangkok 10900THAILANDTel: 662-5794122Fax: 662-5613993E-mail: somkiatkc@fisheries.go.thDr. Supranee ChinabutAquatic Animal Health Research InstituteDepartment of FisheriesKasetsart University CampusJatujak, Ladyao, Bangkok 10900THAILANDTel: 662-5794122Fax: 662-5613993E-mail: supranee@fisheries.go.thDr. Melba B. ReantasoNetwork of Aquaculture Centres in Asia PacificDepartment of Fisheries CompoundKasetsart University CampusJatujak, Ladyao, Bangkok 10900THAILANDTel: 662- 561-1728 to 9 ext. 113Fax: 662-561-1727E-mail: Melba.Reantaso@enaca.orgDr. Kei YuasaFisheries and Aquaculture International Co., Ltd.No. 7 Khoji-machi Bldg., Room B1054-5 Khoji-machi, Chiyoda-kuTokyo 102-0083JAPANTel: 81-3-3234-8847Fax:81-3-3239-8695E-mail: fai@faiaqua.com; yuasakei@hotmail.comDr Myoung-Ae ParkPathology DivisionNational Fisheries Research and Development InstituteSouth Sea Regional Fisheries Research Institute347 Anpo-ri, Hwayang-myun, Yeosu-CityChullanam-do, 556-820KOREA ROTel: 82-662-690-8989Fax: 82-662-685-9073E-mail : mapark@haema.nfrda.re.krDr. Lin LiGuangdong Daya Wan Fisheries Development CenterAotou Town Huizhou City, Guangdong ProvincePEOPLE’s REPUBLIC OF CHINATel: 86-752-5574225Fax: 86-752-5578672E-mail: hzgdfdc@pub.huizhou.gd.cn; linli39@hotmail.comDr. Qiwei QinTropical Marine Science InstituteNational University of Singapore10 Kent Ridge Crescent 119260SINGAPORETel: +65-7749656Fax: +65-7749654Email: tmsqinqw@nus.edu.sg99

Annex F.AII.List of Regional Resource Experts forFinfish Diseases in Asia PacificLymphocystisDiseases of Grass CarpParasitic DiseasesDr Shau Chi ChiDepartment of ZoologyNational Taiwan UniversityTAIWAN PROVINCE of CHINAFax : 886 2 2367 3852E-mail : shauchi@ccms.ntu.edu.twDr Nguyen Huu DungCenter for Bio-Tech and Environment ResearchUniversity of Fisheries02 Nguyen Dinh Chieu St.Nha Trang CityVIETNAMTel: 84 58 83 2065Fax: 84 58 83 1147E-mail : huudung@dng.vnn.vnProf. Jiang YulinShenzhen Exit and Entry Inspection and Quarantine Bureau40 Heping Road, Shenzhen 518010PEOPLE’s REPUBLIC OF CHINATel: 86-755-5592980Fax:86-755-5588630E-mail: szapqbxi@public.szptt.net.cnProf. Jiang YulinShenzhen Exit and Entry Inspection and Quarantine Bureau40 Heping Road, Shenzhen 518010PEOPLE’s REPUBLIC OF CHINATel: 86-755-5592980Fax:86-755-5588630E-mail: szapqbxi@public.szptt.net.cnDr. Robert D. AdlardQueensland MuseumPO Box 3300, South Brisbane,Queensland 4101AUSTRALIATel: +61 7 38407723Fax: +61 7 38461226E-mail: RobertAd@qm.qld.gov.au;http://www.qmuseum.qld.gov.auProfessor R.J.G. LesterDepartment of Microbiology and ParasitologyThe University of Queensland, Brisbane 4072AUSTRALIATel: +61-7-3365-3305Fax:+61-7-3365-4620E-mail:R.Lester@mailbox.uq.edu.au; http://www.biosci.uq.edu.au/micro/academic/lester/lester.htmDr. Ian D. WhittingtonDepartment of Microbiology and ParasitologyThe University of QueenslandBrisbane, Queensland 4072AUSTRALIATel: 61-7-3365-3302Fax:61-7-3365-4620E-mail: i.Whittington@mailbox.uq.edu.auProf. Abu Tweb Abu AhmedDepartment of ZoologyDhaka – 100BANGLADESHTel: 880-2-9666120100

Annex F.AII.List of Regional Resource Experts forFinfish Diseases in Asia PacificFax:880-2-8615583E-mail: zooldu@citechno.netProf. Kazuo OgawaDepartment of Aquatic BioscienceGraduate School of Agricultural and Life SciencesThe University of TokyoYayoi, Bunkyo-ku Tokyo 113-8657JAPANTel: 81-3-5841-5282Fax:81-3-5841-5283E-mail: aogawak@mail.ec-u-tokyo.ac.jpDr. Bo Young JeePathology DivisionNational Fisheries Research and Development Institute408-1,Shirang-ri, Kitang-upKitang-gun, PusanKOREA ROTel: 82-51-720-2498Fax:82-51-720-2498E-mail: protjee@nfrdi.re.krDr. Kim Jeong-HoLaboratory of Aquatic Animal DiseasesCollege of Veterinary MedicineChungbuk National UniversityCheongju, Chungbuk 361-763KOREA ROTel: +82-43-261-3318Fax: +82-43-267-3150E-mail: kimjh89@trut.chungbuk.ac.kr, kimjh89@yahoo.comDr. Craig J. HaywardLaboratory of Aquatic Animal DiseasesCollege of Veterinary MedicineChungbuk National UniversityCheongju, Chungbuk 361-763,KOREA ROTel: +82-43-261-2617Fax: +82-43-267-3150E-mail: chayward@trut.chungbuk.ac.kr, dameunagi@hotmail.comDr. Susan Lim Lee-HongInstitute of Biological SciencesInstitute of Postgraduate Studies and ResearchUniversiti of Malaya50603 Kuala LumpurMALAYSIATel: 603-7594502Fax: 603-7568940E-mail: susan@umcsd.um.edu.myProf. Mohammed ShariffFaculty of Veterinary MedicineUniversiti Putra Malaysia43400 Serdang, SelangorMALAYSIATel: 603-9431064; 9488246Fax: 603-9488246; 9430626E-mail: shariff@vet.upm.edu.my101

Annex F.AII.List of Regional Resource Experts forFinfish Diseases in Asia PacificBacterial DiseasesDr. Leong Tak SengNo. 3 Cangkat Minden, Lorong 1311700 Glugor, Pulau PinangMALAYSIAE-mail: mhpg@pc.jaring.myDr. Tin TunDepartment of ZoologyUniversity of MandalayMYANMARE-mail: phys.mdy@mptmail.net.mmDr. Erlinda LacierdaFish Health SectionAquaculture DepartmentSoutheast Asian Fisheries Development CenterTigbauan, Iloilo 5021PHILIPPINESTel: 63 33 335 1009Fax: 63 33 335 1008E-mail: eclacier@aqd.seafdec.org.phDr. Supranee ChinabutAquatic Animal Health Research InstituteDepartment of FisheriesKasetsart University CampusJatujak, Ladyao, Bangkok 10900THAILANDTel: 662-5794122Fax: 662-5613993E-mail: supranee@fisheries.go.thDr. Melba B. ReantasoNetwork of Aquaculture Centres in Asia PacificDepartment of Fisheries CompoundKasetsart University CampusJatujak, Ladyao, Bangkok 10900THAILANDTel: 662- 561-1728 to 9 ext. 113Fax: 662-561-1727E-mail: Melba.Reantaso@enaca.orgMr. Bui Quang TeResearch Institute for Aquaculture No. 1Dinh Bang, Tu Son, Bac NinhVIETNAMDr. Indrani KarunasagarDepartment of Fishery MicrobiologyUniversity of Agricultural SciencesMangalore – 575 002INDIATel: 91-824 436384Fax: 91-824 436384E-mail: mircen@giasbg01.vsnl.net.inProf. Kiyokuni MurogaFish Pathology LaboratoryFaculty of Applied Biological ScienceHiroshima UniversityHigashi-hiroshima 739JAPANE-mail: fpath@hiroshima-u.ac.jpProf. Mohammed ShariffFaculty of Veterinary MedicineUniversiti Putra Malaysia102

Annex F.AII.List of Regional Resource Experts forFinfish Diseases in Asia PacificViral Diseases43400 Serdang, SelangorMALAYSIATel: 603-9431064; 9488246Fax: 603-9488246; 9430626E-mail: shariff@vet.upm.edu.myMr. Jose O. PaclibareFish Health SectionBureau of Fisheries and Aquatic ResourcesArcadia Building, 860 Quezon AvenueQuezon City, Metro Manila, PHILIPPINESTel/Fax: 632-372-5055E-mail: jopac@edsamail.com.phMrs. Celia Lavilla-TorresFish Health SectionAquaculture DepartmentSoutheast Asian Fisheries Development CenterTigbauan, Iloilo 5021PHILIPPINESTel: 63 33 335 1009Fax: 63 33 335 1008E-mail: celiap@aqd.seafdec.org.phDr. Temdoung SomsiriAquatic Animal Health Research InstituteDepartment of FisheriesKasetsart University CampusJatujak, Ladyao, Bangkok 10900THAILANDTel: 662-5794122Fax: 662-5613993E-mail: temdouns@fisheries.go.thMr. Jeong Wan DoPathology DivisionNational Fisheries Research and Development Institute408-1, Shirang-ri, Kitang-upKitang-gun, PusanKOREA ROTel: 82-51-720-2481Fax:82-51-720-2498E-mail: jwdo@nfrdi.re.krProf. Jiang YulinShenzhen Exit and Entry Inspection and Quarantine Bureau40 Heping Road, Shenzhen 518010PEOPLE’s REPUBLIC OF CHINATel: 86-755-5592980Fax:86-755-5588630E-mail: szapqbxi@public.szptt.net.cnDr. Gilda Lio-PoFish Health SectionAquaculture DepartmentSoutheast Asian Fisheries Development CenterTigbauan, Iloilo 5021PHILIPPINESTel: 63 33 335 1009Fax: 63 33 335 1008E-mail: liopo@aqd.seafdec.org.ph103

Annex F.AII.List of Regional Resource Experts forFinfish Diseases in Asia PacificFungal DiseasesFinfish DiseasesDr. Somkiat KanchanakhanAquatic Animal Health Research InstituteDepartment of FisheriesKasetsart University CampusJatujak, Ladyao, Bangkok 10900THAILANDTel: 662-5794122Fax: 662-5613993E-mail: somkiatkc@fisheries.go.thProf. Kishio HataiDivison of Fish DiseasesNippon Veterinary and Animal Science University1-7-1 Kyonan-cho, Musashino, Tokyo 180JAPANTel: 81-0422-31-4151Fax: 81-0422-33-2094E-mail: hatai@scan-net.ne.jpDr. Kei YuasaFisheries and Aquaculture International Co., Ltd.No. 7 Khoji-machi Bldg., Room B1054-5 Khoji-machi, Chiyoda-kuTokyo 102-0083JAPANTel: 81-3-3234-8847Fax:81-3-3239-8695E-mail: fai@faiaqua.com; yuasakei@hotmail.comDr. Mark CraneAAHL Fish Diseases LaboratoryAustralian Animal Health LaboratoryCSIRO Livestock IndustriesPrivate Bag 24, Geelong Vic 3220AUSTRALIATel : +61 3 52 275118Fax: +61 3 52 275555E-mail: mark.crane@li.csiro.auDr. Shuqin WuPearl River Fisheries Research InstituteChinese Academy of Fishery SciencesBaihedong, Guangzhou, Guangdong 510380PEOPLE’s REPUBLIC OF CHINATel: +86 (20) 81517825 ; +86 (20) 81501543Fax: +86 (20) 81504162E-mail: sqwxm@163.netDr. Jian-Guo HeSchool of Life SciencesZhongshan UniversityGuangzhou 510275PEOPLE’s REPUBLIC OF CHINATel: +86-20-84110976Fax: +86-20-84036215Email: lsbrc05@zsu.edu.cnDr. N. NilakarawasamDepartment of ZoologyThe Open UniversityNawala, NugegodaSRI LANKATel.: 094-1-853777 ext. 270E-mail: nnila@ou.ac.lk104

F.AIII. LIST OF USEFUL GUIDES/MANUAL OFFINISH DISEASES IN ASIA-PACIFIC• Atlas of Fish Diseases (1989) by Kishio Hatai, Kazuo Ogawa and Hitomi Hirose (eds.)Midori Shobo, Tokyo, 267 p. (in Japanese)Information: Prof. Kazuo OgawaDepartment of Aquatic BioscienceGraduate School of Agricultural and Life SciencesThe University of TokyoYayoi, Bunkyo, Tokyo 113-8657Tel: +81-3-5841-5282/5284Fax: +81-3-5841-5283E-mail: aogawak@mail.ecc.u-tokyo.ac.jp• Parasites and Diseases of Culture Marine Finfishes in Southeast Asia (1994) by LeongTak SengInformation: Dr. Leong Tak SengNo. 3 Cangkat Minden, Lorong 1311700 Glugor, Pulau PinangMalaysiaE-mail: mhpg@pc.jaring.my• Asian Fish Health Bibliography III Japan by Wakabayashi H (editor). Fish Health SpecialPublication No. 3. Japanese Society of Fish Pathology, Japan and Fish Health Section of AsianFisheries Society, Manila, PhilippinesInformation: Japanese Society of Fish Pathology• Checklist of the Parasites of Fishes of the Philippines by J. Richard Arthur and S. Lumanlan-Mayo. 1997. FAO Fisheries Technical Paper 369. 102p.Information: Dr. Rohana P. SubasingheFAO of the United NationsViale delle Terme di CaracallaRome 00100 ItalyE-mail: Rohana.Subasinghe@fao.org• Manual for Fish Diseases Diagnosis: Marine Fish and Crustacean Diseases in Indonesia(1998) by Zafran, Des Roza, Isti Koesharyani, Fris Johnny and Kei YuasaInformation: Gondol Research Station for Coastal FisheriesP.O. Box 140 Singaraja, Bali, IndonesiaTel: (62) 362 92278Fax: (62) 362 92272• Diagnostic Procedures for Finfish Diseases (1999) by Kamonporn Tonguthai, SupraneeChinabut, Temdoung Somsiri, Pornlerd Chanratchakool and Somkiat KanchanakhanInformation: Aquatic Animal Health Research InstituteDepartment of FisheriesKasetsart University CampusJatujak, Ladyao, Bangkok 10900THAILANDTel: (66.2) 579.41.22Fax: (66.2) 561.39.93E-mail: aahri@fisheries.go.th• Pathology and Histopathology of Epizootic Ulcerative Syndrome (EUS) by SupraneeChinabut and RJ RobertsInformation: Aquatic Animal Health Research InstituteDepartment of FisheriesKasetsart University CampusJatujak, Ladyao, Bangkok 10900THAILANDTel: (66.2) 579.41.22Fax: (66.2) 561.39.93105

F.AIII. List of Useful Guides/Manual of FinfishDiseases in Asia-PacificE-mail: aahri@fisheries.go.th• Fish Health for Fisfarmers (1999) by Tina ThorneInformation: Fisheries Western Australia3rd Floor, SGIO Atrium186 St. Georges Terrace, Perth WA 6000Tel: (08) 9482 7333Fax: (08) 9482 7389Web: http://www.gov.au.westfish• Australian Aquatic Animal Disease – Identification Field Guide (1999) by Alistair Herfortand Grant RawlinInformation: AFFA Shopfront – Agriculture, Fisheries and Forestry – AustraliaGPO Box 858, Canberra, ACT 2601Tel: (02) 6272 5550 or free call: 1800 020 157Fax: (02) 6272 5771E-mail: shopfront@affa.gov.au• Manual for Fish Disease Diagnosis - II: Marine Fish and Crustacean Diseases in Indonesia (2001) by Isti Koesharyani, Des Roza, Ketut Mahardika, Fris Johnny, Zafran and KeiYuasa, edited by K. Sugama, K. Hatai, and T NakaiInformation: Gondol Research Station for Coastal FisheriesP.O. Box 140 Singaraja, Bali, IndonesiaTel: (62) 362 92278Fax: (62) 362 92272106

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Basic Anatomy of an Oysterdigestive glanddenticlemouthstomachrectumpericardial cavitymantleanusgilladductor muscleBasic anatomy of an oyster.108

SECTION 3 - MOLLUSCAN DISEASESBasic Anatomy of an OysterSECTION 3 - MOLLUSCAN DISEASESM.1 GENERAL TECHNIQUESM.1.1M.1.1.1M.1.1.2M.1.1.3M.1.1.4M.1.2M.1.3M.1.3.1M.1.3.2M.1.3.3M.1.3.4M.1.3.5M.1.3.6M.1.3.7M.1.4M.1.4.1M.1.4.2M.1.4.3M.1.5Gross ObservationsBehaviourShell Surface ObservationsInner Shell ObservationsSoft-Tissue SurfacesEnvironmental ParametersGeneral ProceduresPre-Collection PreparationBackground InformationSample Collection for Health SurveillanceSample Collection for Disease DiagnosisLive Specimen Collection for ShippingPreservation of Tissue SamplesShipping Preserved SamplesRecord KeepingGross ObservationsEnvironmental ObservationsStocking RecordsReferencesDISEASES OF MOLLUSCSM.2 Bonamiosis (Bonamia sp., B. ostreae)M.3 Marteiliosis (Marteilia refringens, M. sydneyi)M.4 Mikrocytosis (Mikrocytos mackini, M. roughleyi)M.5 Perkinsosis (Perkinsus marinus, P. olseni)M.6 Haplosporidiosis (Haplosporidium costale,H. nelsoni)M.7 Marteilioidosis (Marteilioides chungmuensis, M.branchialis)M.8 Iridovirosis (Oyster Velar Virus Disease)108110111111111111114114116116116116116116117118118118119119119121125129133138144147M.AIM.AIIM.AIIIANNEXESOIE Reference Laboratories forMolluscan DiseasesList of Regional Resource Experts for MolluscanDiseases in Asia-PacificList of Useful Diagnostic Guides/Manuals toMolluscan Health149150152109

M.1 GENERAL TECHNIQUES(SE McGladdery)(MG Bondad- Reantaso)cFig.M.1.1.1. Gaping hard shell clam,Mercenaria mercenaria, despite air exposureand mechnical tapping.(MG Bondad-Reantaso)dFig.M.1.1.2c,d. Pteria penguin shell with densemulti-taxa fouling, Guian Pearl Farm, EasternPhilippines (1996).(SE McGladdery)Fig.M.1.1.2a. Mollusc encrustment (arrows) ofwinged oyster, Pteria penguin, Guian PearlFarm, Eastern Samar, Philippines (1996).(D Ladra)Fig.M.1.1.2e. Polydora sp. tunnels and shelldamage at hinge of American oyster,Crassostrea virginica, plus barnacle encrustingof other shell surfaces.(MG Bondad-Reantaso)Fig.M.1.1.2b. Pteria penguin cultured at GuianPearl Farm, Eastern Samar, Philippines with extensiveshell damage due to clionid (boring)sponge (1992).>Fig.M.1.1.2f. Winged oyster, Pteria penguin,shell with clionid sponge damage. Guian PearlFarm, Eastern, Philippines (1996).110

M.1 General TechniquesGeneral molluscan health advice and othervaluable information are available from the OIEReference Laboratories, Regional ResourceExperts in the Asia-Pacific, FAO and NACA.A list is provided in Annexes M.A1 and M.AII,and up-to-date contact information may beobtained from the NACA Secretariat inBangkok (e-mail: naca@enaca.org). Otheruseful guides to diagnostic procedures whichprovide valuable references for molluscan diseasesare listed in Annex M.AIII.M.1.1 Gross ObservationsM.1.1.1 Behaviour (Level I)It is difficult to observe behavioural changes inmolluscs in open-water, however, close attentioncan be made of behaviour of both broodstockand larvae in hatcheries. Since disease situationscan erupt very quickly under hatchery conditions,regular and close monitoring is worthLevel I efforts (see Iridovirus - M.8).Feeding behaviour of larval molluscs is also agood indicator of general health. Food accumulationin larval tanks should be noted and samplesof larvae examined, live, under a dissecting microscopefor saprobiotic fungi and protists (e.g.ciliates) and/or bacterial swarms. Pre-settlementstages may settle to the bottom prematurely orshow passive circulation with the water flow currentsin the holding tanks.Juvenile and adult molluscs may also cease feeding,and this should be cause for concern undernormal holding conditions. If feeding does notresume and molluscs show signs of weakening(days to weeks depending on water temperature)samples should be collected for laboratory examination.Signs of weakening include gaping (i.e.bivalve shells do not close when the mollusc istouched or removed from the water) (Fig.M.1.1.1), accumulation of sand and debris inthe mantle and on the gills, mantle retractionaway from the edge of the shell, and decreasedmovement in mobile species (e.g. scallop swimming,clam burrowing, abalone grazing, etc.).Open-water mortalities that assume levels ofconcern to the grower should be monitored todetermine if there are any patterns to the losses.Sporadic moralities following periods of intensehandling should be monitored with minimal additionalhandling if at all possible. If the mortalitiespersist, or increase, samples should be collectedfor laboratory analysis. Mortalities that appear tohave a uniform distribution should be examinedimmediately and environmental factors pre- andpost-mortality recorded. Mortalities that appearto spread from one area to another suggest thepresence of an infectious disease agent andshould be sampled immediately. Affected animalsshould be kept as far away as possible from unaffectedanimals until the cause of the mortalitiescan be determined.M.1.1.2 Shell Surface Observations(Level I)Fouling organisms (barnacles, limpets, sponges,polychaete worms, bivalve larvae, tunicates,bryozoans, etc.) are common colonists of molluscshell surfaces and do not normally presenta threat to the health of the mollusc (Fig.M.1.1.2a,b). Suspension and shallow water culture,however, can increase exposure to foulingand shells may become covered by other animalsand plants (Fig. M.1.1.2c,d). This can affecthealth directly by impeding shell opening andclosing (smothering) or indirectly through competitionfor food resources. Both circumstancescan weaken the mollusc so cleaning may be required.Such defouling should be undertaken asrapidly as possible, to minimise the period of removalfrom the water, during cooler periods ofthe day. Rapid cleaning is usually achieved usinghigh pressure water or mechanical scapers.Defouled molluscs should be returned to cleanwater. Fouling organisms should not be discardedin the same area as the molluscs, since this willaccelerate recolonisation. Signs of weakeningthat persist or increase after cleaning, should beinvestigated further by laboratory examination.Shell damage by boring organisms, such assponges and polychaete worms (Fig. M.1.1.2e,f) is normal in open-water growing conditions.Although usually benign, under certain conditions(especially in older molluscs) shells may be renderedbrittle or even become perforated. Suchdamage can weaken the mollusc and render itsusceptible to pathogen infections.Shell deformities (shape, holes in the surface),fragility, breakage or repair should be noted, butare not usually indicative of a disease condition(Fig.M.1.1.2g, h). Abnormal colouration andsmell, however, may indicate a possible soft-tissueinfection which may require laboratory examination.M.1.1.3 Inner Shell Observations (Level I)The presence of fouling organisms (barnacles,sponges, polychaete worms, etc.) on the innershell surface is a clear indication of a weak/111

M.1 General Techniques(MG Bondad-Reantaso)(B Jones)gFig.M.1.1.3a1. Abalone (Haliotis roei) from abatch killed by polydoriid worms.hFig.M.1.1.2g,h. Pinctada maxima, shell withclionid sponge damage due to excavation oftunnels exhalent-inhalent openings (holes) tothe surface (arrows). Other holes (small arrows)are also present that may have been causedby polychaetes, gastropod molluscs or otherfouling organisms. Guian Pearl Farm, EasternPhilippines (1996).(MG Bondad-Reantaso)(D Ladra)bFig.M.1.1.3a. Winged oyster, Pteria penguin,shell showing clionid sponge damage throughto the inner shell surfaces, Guian Pearl Farm,Eastern Philippines (1996).cFig.M.1.1.3b,c. b. Shells of Pinctada maximashowing a erosion of the nacreous inner surfaces(arrows), probably related to chronicmantle retraction; c. Inner surface of shellshowing complete penetration by boringsponges (thin arrows).112

M.1 General Techniques(MG Bondad-Reantaso)(MG Bondad-Reantaso)dFig.M.1.1.3g. Inner shell of winged pearl oystershowing: tunnels at edge of the shell (straightthick arrow); light sponge tunnel excavation(transparent arrow); and blisters (small thickarrow) at the adductor muscle attachment site.Guian Pearl Farm, Eastern Philippines (1996).(SE McGladdery)eFig.M.1.1.3.h. Extensive shell penetration bypolychaetes and sponges causing weakeningand retraction of soft-tissues away from theshell margin of an American oyster Crassostreavirginica.fFig.M.1.1.3d,e,f. Pinctada maxima (d), Pteriapenguin (e) and edible oyster (Crassostrea sp.)(f) shells showing Polydora-related tunnel damagethat has led to the formation of mud-filledblisters.113

M.1 General Techniquessick mollusc (Fig.M.1.1.3a and Fig.M.1.1.3a1).The inner surfaces are usually kept cleanthrough mantle and gill action. Perforation ofthe inner surface can be sealed off by depositionof additional conchiolin and nacre(Fig.M.1.1.3b,c). This may result in the formationof a mud- or water-filled “blister”(Fig.M.1.1.3d,e,f). Shell coverage can also occurover irritants attached to, or lying up againstthe inner shell, a process that may result in a“blister pearl” (Fig.M.1.1.3g).Where perforation or other irritants exceed repair,the health of the mollusc is jeopardisedand it becomes susceptible to opportunistic infections(Fig.M.1.1.3.h). The degree of shellperforation can be determined by holding theshell up to a strong light.Where abnormalities occurring within the matrixof the shell warrant further investigation,freshly collected specimens can be sent intactto the laboratory or fixed for subsequent decalcification,as required.M.1.1.4 Soft-Tissue Surfaces (Level I)The appearance of the soft-tissues is frequentlyindicative of the physiological condition of theanimal. Gross features which should be recordedinclude:• condition of the animal as listed below:fat - the soft-tissues fill the shell, are turgidand opaquemedium - the tissues are more flaccid,opaque and may not fill the shell cavitywatery - the soft-tissues are watery/transparentand may not fill the shell cavity (Fig.(Fig.M.1.1.4a, Fig.M.1.1.4b)• colour of the digestive gland – e.g., pale,mottled, dark olive• any abnormal enlargement of the heart orpericardial cavity – e.g., cardiac vibriosis,tumours• presence of focal lesions such as:abnormal coloration (eg., patches of green,pink, red, black, etc.)abscesses (Fig. M.1.1.4c)tumour-like lesions (Fig. M.1.1.4d)tissue (e.g. gill) erosion• presence of water blisters in the viscera, palp,or mantle (Fig.M.1.1.4e)• presence of pearls or other calcareous deposits(Fig.M.1.1.4f) within the soft tissues• presence of parasites or commensals suchas:pea crabs in mantle cavityparasitic copepods attached to gillspolychaetes, nematodes and turbellarians inmantle cavity or on surrounding surfaces(Fig. M.1.1.4g)redworm (Mytilicola spp.) usually exposedonly on dissection of the digestive tractciliates (sessile or free-swimming) and otherprotistans (for larvae only)bacteria (for larvae only)• any mechanical (e.g., knife) damage to thesoft-tissues during the opening of the shell.Abscess lesions, pustules, tissuediscolouration, pearls, oedema (water blisters),overall transparency or wateriness, gill deformities,etc., can be present in healthy molluscs,but, if associated with weak or dying animals,should be cause for concern. Record the levelsof tissue damage and collect samples ofboth affected and unaffected animals for laboratoryexamination. Moribund animals, or thosewith foul-smelling tissues may be of little usefor subsequent examination (especially fromwarm water conditions), however, numbers affectedshould be recorded.Worms or other organisms (e.g. pea crabs,copepods, turbellarians) on the soft-tissues arealso common and not generally associated withdisease. If present in high numbers on weakmolluscs, however, numbers should be notedand samples of intact specimens collected forlaboratory examination and identification. Fixationin 10% buffered formalin is usually adequatefor preserving the features necessaryfor subsequent identification.M.1.2 Environmental Parameters(Level I)Environmental conditions have a significant effecton molluscan health, both directly (withinthe ranges of physiological tolerances) and indirectly(enhancing susceptibility to infections).This is especially important for species grownunder conditions which differ significantly fromthe wild (e.g. oysters grown in suspension).Important environmental factors for molluscanhealth include water temperature, salinity, turbidity,fouling and plankton blooms. Extremesand/or rapid fluctuations in these can seriouslycompromise molluscan health. Anthropogenicfactors include a wide range of biologic andchemical pollutants. Since molluscs are, essentially,sessile species (especially under cultureconditions) this renders them particularly susceptibleto pollution. In addition, molluscs have114

M.1 General Techniques(SE McGladdery)(SE McGladdery)Fig.M.1.1.4a. Normal oyster (Crassostreavirginica) tissues.(SE McGladdery)Fig.M.1.1.4e. Water blister (oedema/edema) inthe soft-tissues of the mantle margin of anAmerican oyster (Crassostrea virginica).(SE McGladdery)Fig.M.1.1.4b. Watery oyster (Crassostreavirginica) tissues – compare with M.1.1.4a.(SE McGladdery)Fig.M.1.1.4f. Calcareous deposits (“pearls”) inthe mantle tissues of mussels in response toirritants such as mud or digenean flatwormcysts.(SE McGladdery and M Stephenson)Fig.M.1.1.4c. Abscess lesions (creamy-yellowspots, see arrows) in the mantle tissue of a Pacificoyster (Crassostrea gigas).(MS Park and DL Choi)Fig.M.1.1.4g. Polydoriid tunnels underlying thenacre at the inner edge of an American oyster(Crassostrea virginica) shell, plus another freelivingpolychaete, Nereis diversicolor on the innershell surface.>Fig.M.1.1.4d. Gross surface lesions in Pacificoyster (Crassostrea gigas) due to Marteiliodeschungmuensis.115

M.1 General Techniqueslow tolerance of some other water-uses/abuses(e.g., dynamite and cyanide fishing; dragging;creosote and other anti-fouling chemical compounds;agricultural run-off).Maintaining records of temperature, salinity (inestuarine or coastal areas), turbidity and manmadedisturbances provide valuable backgrounddata, essential for accurate interpretation of mortalityobservations and results from laboratoryanalyses.M.1.3 General ProceduresM.1.3.1 Pre-Collection PreparationWherever possible, check the number of specimensrequired for laboratory examination withlaboratory personnel before collecting thesample. Ensure that each specimen is intact,i.e., no empty or mud-filled shells. Largersample numbers are generally needed forscreening purposes compared with numbersrequired for disease diagnosis.M.1.3.2 Background Information (Level I)All samples being submitted for laboratory examinationshould include as much backgroundinformation as possible, such as:• reason(s) for submitting the sample (mortalities, abnormal growth/spawning, healthscreening, etc.);• gross observations and environmental paraeters (as described under M.1.1 and M.1.2);• where samples are submitted due to mortalties, approximate prevalences and patterns ofmortality (acute or chronic/sporadic cumulativelosses), and• whether or not the molluscs are from local populationsor from another site. If the stock is notlocal, the source and date of transfer shouldalso be noted.The above information will help identify if handlingstress, change of environment or infectiousagents may be a factor in mortalities. Itwill also help speed up accurate diagnosis of adisease problem or disease-risk analysis.M.1.3.3 Sample Collection for Health SurveillanceThe most important factors associated with collectionof specimens for surveillance are:• sample numbers that are high enough (seeTable M.1.3.3 below)• susceptible species are sampled• samples include age- or size-groups that aremost likely to manifest detetctable infections.Such information is given under specific diseasesections.The standard sample sizes for screening healthyaquatic animals, including molluscs, is given inTable M.1.3.3 below.M.1.3.4 Sample Collection for Disease DiagnosisAll samples submitted for disease diagnosisshould include as much supporting informationas possible including:• reason(s) for submitting the sample (mortalities,abnormal growth, etc.)• handling activities (de-fouling, size sorting/grading, site changes, new species/stock introduction,etc.)• history and origin(s) of the affectedpopulation(s);• environmental changesM.1.3.5 Live Specimen Collection for Shipping(Level I)Once the required number of specimens hasbeen determined, and the laboratory has provideda date or time for receipt of the sample, the molluscsshould be collected from the water. Thisshould take place as close to shipping as possibleto reduce air-storage changes in tissues andpossible mortalities during transportation. This isespecially important for moribund or diseasedmollusc samples.The laboratory should be informed of the estimatedtime of arrival to ensure they have thematerials required to process the sample preparedbefore the sample arrives. This helps reducethe time between removal from the waterand preservation of the specimens for examination.The molluscs should be wrapped in papersoaked with ambient seawater. For small seed(

M.1 General TechniquesPrevalence (%)Population Size 0.5 1.0 2.0 3.0 4.0 5.0 10.050 46 46 46 37 37 29 20100 93 93 76 61 50 43 23250 192 156 110 75 62 49 25500 314 223 127 88 67 54 261000 448 256 136 92 69 55 272500 512 279 142 95 71 56 275000 562 288 145 96 71 57 2710000 579 292 146 96 72 29 27100000 594 296 147 97 72 57 271000000 596 297 147 97 72 57 27>1000000 600 300 150 100 75 60 30Table M.1.3.3 1 . Sample sizes needed to detect at least one infected host in a population of agiven size, at a given prevalence of infection. Assumptions of 2% and 5% prevalences are mostcommonly used for surveillance of presumed exotic pathogens, with a 95% confidence limit.to freshwater ice (gel-paks or plastic bottlescontaining frozen water are recommended overloose ice to keep specimens cool) and to reduceloss of mantle fluids.Label containers clearly:“Live Specimens, Store at _____°C to ______°CDO NOT FREEZE”If being shipped by air also indicate:“HOLD AT AIRPORT AND CALL FOR PICK-UP”Clearly indicate the name and telephone numberof the contact person responsible for pickingup the package at the airport or receiving itat the laboratory.Ship early in the week to avoid arrival duringthe weekend with possible loss of samples dueto improper storage. Inform the contact personas soon as the shipment has been sent and,where appropriate, give them the name of thecarrier and waybill number.M.1.3.6 Preservation (Fixation) of TissueSamples (Level I - with basic training)For samples that cannot be delivered live to adiagnostic laboratory, due to distance or slowtransportation, specimens should be fixed (preserved)on site. This is suitable for subsequenthistology examination, but means that routinebacteriology, mycology or media culture (e.g.,Fluid Thioglycollate Medium culture of Perkinsusspp.) cannot be performed. Diagnostic needsshould, therefore, be discussed with laboratorypersonnel prior to collecting the sample.The following fixatives can be used for preservationof samples:i) 1G4F solution (1% Glutaraldehyde : 4% Formaldehyde)*Stock 1G4F solution - may be held at 4oC forup to 3 months:120 ml 37-40% buffered formalin solution**20 ml 50% glutaraldehyde360 ml tap water**Buffered formalin solution:1 litre 37-40% formaldehyde15 gm disodium phosphate (Na 2HPO 4)1Ossiander, F.J. and G. Wedermeyer. 1973. Journal of Fisheries Research Board of Canada 30:1383-1384.117

M.1 General Techniques0.06 gm sodium hydroxide (NaOH)0.03 gm phenol red (pH indicator)Working solution – should be preparedimmediately prior to use:500 ml filtered ambient seawater orInstant Ocean500 ml Stock 1G4F solution*The required tissue thickness is about 2-3 mm.Tissues can tolerate long storage in this fixativeat room temperature. (N.B. Thicker tissues, orwhole animals, may be fixed using the 10%buffered formalin solution as described below).ii) 10% Buffered formalin in filtered ambientseawater (This is the easiest solution to prepareand store).10 ml 37-40% buffered formalinsolution**90 ml filtered ambient seawaterN.B. Whole specimens less than 10 mm thickcan be fixed with this solution. If the specimensare larger, cut them into two or more piecesbefore fixing (ensure that pieces from differentspecimens do not get mixed up).iii) Davidson’s FixativeTissue up to 10 mm in thickness can be fixedin Davidson’s fixative. Prior to embedding, tissuesneed to be transferred to either 50% ethanolfor 2 hours (minimum) and then to 70%ethanol, or directly to 70% isopropanol. Bestresults are obtained if fixative is made up in thefollowing order of ingredients.Stock Solution:400 ml glycerin800 ml formalin(37-40% formaldhyde)1200 ml 95% ethanol (or 99% iso propanol)1200 ml filtered natural or artificialseawaterWorking Solution: dilute 9 parts stock solutionwith 1 part glacial acetic acidImportant Notes:• All fixatives should be kept away from openwater and used with caution against contactwith skin and eyes.• If the molluscs cannot be fixed intact, contactthe diagnostic laboratory to get guidancefor cracking shell hinges or removing the requiredtissues.M.1.3.7 Shipping Preserved Samples(Level I)Many transport companies (especially air carriers)have strict regulations regarding shippingany chemicals, including fixed samples for diagnosticexamination. Check with the carrierbefore collecting the sample to prevent loss oftime and/or specimens due to inappropriatepackaging, labeling, etc.. If the tissues havebeen adequately fixed (as described in M.1.3.4),most fixative or storage solution can be drainedfrom the sample for shipping purposes. As longas sufficient solution is left to keep the tissuesfrom drying out, this will minimise the quantityof chemical solution being shipped. Pack fixedsamples in a durable, leak-proof container.Label containers clearly with the information asdescribed for live specimens (M.1.3.3.). Clearlyindicate the name and telephone number of thecontact person responsible for picking up thepackage at the airport or receiving it at the laboratory.Ship early in the week to avoid arrivalduring the weekend with possible loss ofsamples due to improper storage. Inform thecontact person as soon as the shipment hasbeen sent and, where appropriate, give themthe name of the carrier and waybill number.If being shipped by air also indicate:“HOLD AT AIRPORT AND CALL FOR PICK-UP”M.1.4 Record-Keeping (Level I)Record keeping is essential for effective diseasemanagement. For molluscs, many of thefactors that should be recorded are outlined insections M.1.4.1, M.1.4.2, and M.1.4.3.M.1.4.1 Gross Observations (Level I)Gross observations can be included with routinemonitoring of mollusc growth, either bysub-sampling from suspension cages, lines orstakes, or by guess estimates from surfaceobservations.For hatchery operations, the minimum essentialinformation which should be recorded/logged are:• feeding activity• growth• mortalitiesThese observations should be recorded on adaily basis for larval and juvenile molluscs, includingdate, time, tank, broodstock (where118

M.1 General Techniquesthere are more than one) and food-source (algalculture batch or other food-source). Datesand times for tank and water changes shouldalso be noted, as well as dates and times forpipe flushing and/or disinfection. Ideally, theselogs should be checked regularly by the personresponsible for the site/animals.For open-water mollusc sites, the minimum essentialobservations which need to be recorded/logged include:• growth• fouling• mortalitiesThese should be recorded with date, site locationand any action if taken (e.g., defouling orsample collection for laboratory examination).Ideally, these logs should be checked regularlyby the person responsible for the site/animals.M.1.4.2 Environmental Observations(Level I)This is most applicable to open water sites, butshould also be included in land-based systemswith flow-through or well-based water sources.The minimum essential data which should berecorded are:• temperature• salinity• turbidity (qualitative evaluation or secchi disc)• algal blooms• human activityThe frequency of these observations will varywith site. Where salinity or turbidity rarely vary,records may only be required during rainy seasonsor exceptional weather conditions. Temperateclimates will require more frequent watertemperature monitoring than tropical climates.Human activity should be logged on an“as it happens” basis for reference if no infectionsor natural environmental changes can beattributed to a disease situation.M.1.4.3 Stocking Records (Level I)Information on movements of molluscs into andout of a hatchery should be recorded. Thisshould include:• exact source of the broodstock/seed• condition on arrival• date, time and person responsible for receivingdelivery of the stock• date, time and destination of stock shippedout of the hatcheryWhere possible, animals from different sourcesshould not be mixed.All movements of molluscs onto and off anopen-water site should also be recorded, including:• exact source of the molluscs• condition on arrival• date, time and person responsible for receivingdelivery of the stock• date, time and destination of stock shippedoff siteIn addition, all movements of stocks within ahatchery, nursery or grow out site should belogged with the date for tracking purposes if adisease situation arises.M.1.5 ReferencesElston, R.A. 1989. Bacteriological methods fordiseased shellfish, pp. 187-215. In: Austin,B. and Austin, D.A. (eds.) Methods for theMicrobiological Examination of Fish andShellfish. Ellis Horwood Ser. Aquac. Fish.Sup.. Wiley and Sons, Chichester, UK.Elston, R.A. 1990. Mollusc diseases: Guide forthe Shellfish Farmer. Washington Sea GrantProgram, University of Washington Press,Seattle. 73 pp.Elston, R.A., E.L. Elliot, and R.R. Colwell. 1982.Conchiolin infection and surface coatingVibrio: Shell fragility, growth depression andmortalities in cultured oysters and clams,Crassostrea virginica, Ostrea edulis andMercenaria mercenaria. J. Fish Dis. 5:265-284.Fisher, W.S. (ed.). 1988. Disease Processes inMarine Bivalve Molluscs. Amer. Fish. Soc.Spec. Public. 18. American Fisheries Society,Bethesda, Maryland, USA.Howard, D.W. and C.S. Smith. 1983. HistologicalTechniques for Bivalve Mollusks. NOAATech. Memo. NMFS-F/NEC-25.Woods Hole,Massachusetts. 97 pp.Lauckner, G. 1983. Diseases of Mollusca:Bivalvia, pp. 477-520. In: O. Kinne(ed.) Diseasesof Marine Animals Volume II. IntroductionBivalvia to Scaphopoda. BiologischeAnstalt Helgoland, Hamburg.119

M.1 General TechniquesLuna, L.G. 1968. Manual of Histologic StainingMethods of the Armed Forces Institute of Pathology.McGraw-Hill Book Company, NewYork. 258 pp.McGladdery, S.E., R.E. Drinnan, and M.F.Stephenson. 1993. A manual of the parasites,pests and diseases of Canadian Atlanticbivalves. Can. Tech. Rep. Fish. Aquat.Sci.1931.121 pp.Ossiander, F.J. and G. Wedermeyer. 1973.Computer program for sample size requiredto determine disease incidence in fish populations.J. Fish. Res. Bd. Can. 30: 1383-1384.Pass, D.A., R. Dybdahl, and M.M. Mannion.1987. Investigations into the causes of mortalityof the pearl oyster (Pinctada maxima)(Jamson), in Western Australia. Aquac.65:149-169.Perkins, F.O. 1993. Infectious diseases ofmoluscs, pp. 255-287. In: Couch, J.A. andFournie, J.W. (eds.). Pathobiology of Marineand Estuarine Organisms. CRC Press, BocaRaton, Florida.120

MOLLUSCAN DISEASESM.2 BONAMIOSIS(BONAMIA SP., B. OSTREAE)M.2.1 Background InformationM.2.1.1 Causative AgentsBonamiosis (a.k.a. Microcell Disease;haemocyte disease of flat or dredge oysters) iscaused by two Protistan (= Protozoan = singlecelled)species belonging to the Haplosporidia:Bonamia ostreae and Bonamia sp.. More informationabout the disease can be found in theOIE Diagnostic Manual for Aquatic Animal Diseases(OIE 2000a).M.2.1.2 Host RangeBonamia ostreae occurs naturally in Ostrea edulis(European oyster) and O. conchaphila (O. lurida)(Olympia oyster). Other ostreiid species can becomeinfected if transferred to enzootic areas,namely O. puelchana, O. angasi and Ostrealutaria (Tiostrea lutaria) (New Zealand oyster),Tiostrea chilensis (Ostrea chilensis) (SouthAmerican oyster), thus, all species of Ostrea,Tiostrea and some Crassostrea (C. ariakensis)should be considered susceptible. To date,Crassostrea gigas (Pacific oyster), Mytilus edulisand M. galloprovincialis (edible mussels) andRuditapes decussatus and R. philippinarum (Europeanand Manila clams) have been found tobe resistant to infection. These species have alsobeen shown to be incapable of acting as reservoirsor sub-clinical carriers of infection.M.2.1.3 Geographic DistributionBonamia ostreae: The Netherlands, France,Spain, Italy, Ireland, the United Kingdom (excludingScotland) and the United States of America(States of California, Maine and Washington).Denmark, although stocked with infected oystersin the early 1980’s, has shown no sign of persistenceof the infection and their European oystersare now considered to be free of B. ostreae.Bonamia sp.: Australia (Western Australia,Victoria and Tasmania) and New Zealand (SouthIsland and southern North Island).M.2.1.3 Asia-Pacific Quarterly Aquatic AnimalDisease Reporting System (1999 to2000)For the reporting year 1999, Bonamia sp. waspositively reported in Australia in April, July andOctober in Tasmania; July and October in WesternAustralia. For the year 2000, Bonamia sp.was reported in March and April in Western Australia.In New Zealand, Bonamia sp. wasreported every month for 1999 and 2000reporting periods (OIE 1999, OIE 2000b).M.2.2 Clinical AspectsMost infections show no clinical signs until theparasites have proliferated to a level that elicitsmassive blood cell (haemocyte) infiltration anddiapedesis (Fig.M.2.2a). The pathology of infectionvaries with the species of Bonamia, andwith host species. Bonamia ostreae infects thehaemocytes of European oysters (Fig.M.2.2b),where it divides until the haemocyte bursts, releasingthe parasites into the haemolymph. Infectionslikely occur through the digestive tract,but gill infections suggest this may also be anotherinfection route and macroscopic gill lesionsare sometimes visible. The pathology of Bonamiasp. in Australian Ostrea angasi and New Zealandpopulations of Tiostrea chilensis is very different.In Australia’s O. angasi, the first indicationof infection is high mortality. Surviving oystersrapidly start to gape on removal from the waterand may have “watery” tissues and a raggedappearance to the margin of the gill(unpublished data, B. Jones, Fisheries WesternAustralia). Bonamia sp. infects the walls of thegills, digestive ducts and tubules (Fig.M.2.2c),from which the parasites may be released intothe gut or surrounding water. Infectedhaemocytes may contain up to 6 Bonamia parasites(Fig.M.2.2d). Infections induce massiveabscess-like lesions (haemocytosis), even in thepresence of only a few parasites. In T. chilensis,Bonamia sp. appears to enter via the gut wall(Fig.M.2.2e), and then infects the haemocytes,where up to 18 Bonamia per haemocyte can befound (Fig.M.2.2f). The resultant haemocytosisis less severe than in O. angasi. When infectedhaemocytes enter the gonad of T. chilensis toreabsorb unspawned gametes, the parasites proliferateand may be released via the gonoduct.Alternative release is also possible via tissuenecrosis following the death of the host. Despitethe differences in pathology, gene sequencingstudies (unpublished data, R. Adlard, Universityof Queensland, Australia) have shown that theAustralian and New Zealand Bonamia sp. are thesame species.M.2.3 Screening MethodsMore detailed methods for screening can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.121

M.2 Bonamiosis(Bonamia sp., B. ostreae)(SE McGladdery)(PM Hine)Fig.M.2.2a. Haemocyte infiltration and diapedesisacross intestinal wall of a European oyster(Ostrea edulis) infected by Bonamia ostreae.(SE McGladdery)Fig.M.2.2d. Oil immersion of Bonamia sp. infectingblood cells and lying free (arrows) in thehaemolymph of an infected Australian flat oyster,Ostrea angasi. Scale bar 20µm (H&E).(PM Hine)Fig.M.2.2b. Oil immersion of Bonamia ostreaeinside European oyster (Ostrea edulis)haemocytes (arrows). Scale bar 20 µm.(PM Hine)Fig.M.2.2e. Focal infiltration of haemocytesaround gut wall (star) of Tiostrea lutaria (NewZealand flat oyster) typical of infection byBonamia sp. (H&E).(PM Hine)Fig.M.2.2c. Systemic blood cell infiltration inAustralian flat oyster (Ostrea angasi) infectedby Bonamia sp. Note vacuolised appearanceof base of intestinal loop and duct walls (H&E).Fig.M.2.2f. Oil immersion of haemocytespacked with Bonamia sp. (arrows) in an infectedTiostrea lutaria (H&E). >122

M.2 Bonamiosis(Bonamia sp., B. ostreae)M.2.3.1 PresumptiveM.2.3.1.1 Gross Observations (Level I).Slowed growth, presence of gill lesions (in somecases), gaping and mortalities of Ostrea edulisshould be considered suspect for Bonamiosis.Gross signs are not disease specific and requireLevel II examination.M.2.3.1.2 Cytological Examination(Level II)Spat or heart (preferably ventricle) smears or impressions(dabs) can be made onto a clean microscopeslide and air-dried. Once dry, the preparationis fixed in 70% methanol. Quick and effectivestaining can be achieved using commerciallyavailable blood-staining (cytological) kits, followingthe manufacturer’s instructions. The stainedslides are then rinsed (gently) in tapwater, allowedto dry and cover-slipped using a synthetic resinmounting medium. The parasite has basophilic(or colourless - Bonamia sp. in O. angasi) cytoplasmand an eosinophilic nucleus (dependingon the stain used). An oil immersion observationtime of 10 mins per oyster preparation is consideredsufficient for screening cytology, tissue imprintand histology preparations (OIE 2000a).M.2.3.2 ConfirmatoryM.2.3.2.1 Histopathology (Level II)It is recommended that at least two dorso-ventralsections through the cardiac cavity, gonadand gills of oysters over 18 months – 2 years (>30 mm shell height) be examined for screeningpurposes. These sections should be fixed immediatelyin a fast fixative such as 1G4F. Fixativessuch as Davidsons or 10% seawater bufferedformalin may be used for whole oysters (seeM.1.3.3.3), but these do not allow serial tissuesections to be collected for subsequent confirmatoryElectron Microscopy (EM) diagnosis, ifrequired. Davidson’s fixative is recommendedfor subsequent PCR-based confirmation techniques.Several standard stains (e.g., haematoxylineosin)enable detection of Bonamia spp.. Theparasites measure 2-5 µm and occur within thehaemocytes or epithelia (as described above) or,more rarely, loose within the haemolymph or gut/mantle lumens.M.2.4 Diagnostic MethodsMore detailed methods for diagnosis can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.M.2.4.1 PresumptiveM.2.4.1.1 Histopathology and Cytology(Level II)Histology and cytology (Level II), as describedunder M.2.3.2.1, may be used. For first-timediagnoses, a back up tissue specimen fixed forEM is recommended (M.2.4.2.1).M.2.4. 2 ConfirmatoryM.2.4.2.1 Transmission Electron Microscopy(TEM) (Level III)Tissue for TEM can be fixed in 1G4F (M.1.3.3.3),however, where it is likely that TEM will be requiredfor confirmatory diagnosis (M.2.4.1.1),small (< 1 mm cubed) sub-samples of infectedtissue should be fixed in 2-3% buffered glutaraldehydeprepared with ambient salinity filteredseawater. Fixation should not exceed 1 hr.Longer storage in gluteraldehyde fixative is possible,however membrane artifacts can be produced.Tissues should be rinsed in a suitablebuffer prior to post-fixing in 1-2% osmium tetroxide(= osmic acid - highly toxic). This post-fixativemust also be rinsed with buffered filtered(0.22 µm) seawater prior to dehydration andresin-embedding.Post-fixed tissues should be stored in a compatiblebuffer or embedded post-rinsing in a resinsuitable for ultramicrotome sectioning. Screeningof 1 micron sections melted onto glass microscopeslides and stained with Toluidine Blueis one method of selecting the tissue specimensfor optimum evidence of putative Bonamia spp.Ultrathin sections are then mounted on coppergrids (with or without formvar coating) for stainingwith lead citrate + uranyl acetate or equivalentEM stain.Ultrastructural differences between B. ostreaeand Bonamia sp. include diameter (B. ostreae =2.4 ± 0.5 µm; Bonamia sp. = 2.8 ± 0.4 µm in O.angasi, 3.0 ± 0.3 µm in T. chilensis); mean numberof mitochondrial profiles/section (B. ostreae= 2 ± 1; Bonamia sp. = 4 ± 1 in O. angasi, 3 ±1in T. chilensis), mean number ofhaplosporosomes/section (B. ostreae = 7 ± 5;123

M.2 Bonamiosis(Bonamia sp., B. ostreae)Bonamia sp. = 10 ± 4 in O. angasi, 14 ± 6 in T.chilensis); percentage of sections containinglarge lipid globules (B. ostreae = 7%; Bonamiasp. in O. angasi = 30%; in T. chilensis = 49%),large lipid globules/section (B. ostreae = 0.3 ±0.6; Bonamia sp. = 0.5 ± 0.8 in O. angasi, 0.8 ±0.9 in T. chilensis). Both species are distinguishedfrom Mikrocytos spp. by having a centrally-placednucleus.Plasmodial forms of Bonamia sp. in T. chilensisare distinguished from Bonamia ostreae by theirsize (4.0 -4.5 µm diameter), irregular cell andnucleus profile, amorphous cytoplasmic inclusions(multi-vesicular bodies) and arrays of Golgilikesmooth endoplasmic reticula. Other developmentalstages are more electron dense andsmaller in diameter (3.0 -3.5 µm).M.2.5 Modes of TransmissionPrevalence and intensity of infection tends to increaseduring the warm water season withpeaks in mortality in September/October in thenorthern hemisphere, and January to April, inthe southern hemisphere. The parasite is difficultto detect prior to the proliferation stage ofdevelopment or in survivors of an epizootic. Cohabitationand tissue homogenate/haemolymphinoculations can precipitate infections indicatingthat transmission is direct (no intermediatehosts are required). There is a pre-patent periodof 3-5 months between exposure and appearanceof clinical signs of B. ostreae infection.In New Zealand the pre-patent period forBonamia sp. infection may be as little as 2.5months and rarely exceeds 4 months.M.2.6 Control MeasuresNone known. Reduced stocking densities andlower water temperatures appear to suppressclinical manifestation of the disease, however,no successful eradication procedures haveworked to date. Prevention of introduction ortransfer of oysters from Bonamia spp. enzooticwaters into historically uninfected waters is recommended.M.2.7Selected ReferencesAdlard, R.D. and R.J.G. Lester. 1995. Developmentof a diagnostic test for Mikrocytosroughleyi, the aetiological agent of Australianwinter mortality in the commercial rock oyster,Saccostrea commercialis (Iredale &Roughley). J. Fish Dis. 18: 609-614.Balouet, G., M. Poder, and A. Cahour. 1983.Haemocytic parasitosis: Morphology and pathologyof lesions in the French flat oyster,Ostrea edulis L. Aquac. 43: 1-14.Banning, P. van 1982. The life cycle of the oysterpathogen Bonamia ostreae with a presumptivephase in the ovarian tissue of theEuropean flat oyster, Ostrea edulis. Aquac.84: 189-192.Carnegie, R.B., B.J. Barber, S.C. Culloty, A.J.Figueras, and D.L. Distel. 2000. Developmentof a PCR assay for detection of the oysterpathogen Bonamia ostreae and support forits inclusion in the Haplosporidia. Dis. Aquat.Org. 42(3): 199-206.Culloty, S.C., B. Novoa, M. Pernas, M.Longshaw, M. Mulcahy, S.W. Feist, and A.J.Figueras.1999. Susceptibility of a number ofbivalve species to the protozoan parasiteBonamia ostreae and their ability to act as vectorsfor this parasite. Dis. Aquat. Org. 37(1):73-80.Dinamani, P., P.M. Hine, and J.B. Jones. 1987.Occurrence and characteristics of thehemocyte parasite Bonamia sp. on the NewZealand dredge oyster Tiostrea lutaria. Dis.Aquat. Org. 3: 37-44.Farley, C.A., P.H. Wolf, and R.A. Elston. 1988.A long term study of “microcell” disease in oysterswith a description of a new genus,Mikrocytos (g.n.) and two new species,Mikrocytos mackini (sp.n.) and Mikrocytosroughleyi (sp.n.). Fish. Bull. 86: 581-593.Friedman, C.S. and F.O. Perkins. 1994. Rangeextensiion of Bonamia ostreae to Maine,U.S.A. J. Inverteb. Pathol. 64: 179-181.Hine, P.M. 1991. Ultrastructural observations onthe annual infection pattern of Bonamia sp. inflat oysters, Tiostrea chilensis. Aquac. 93: 241-245.McArdle, J.F., F. McKiernan, H. Foley, and D.H.Jones. 1991. The current status of Bonamiadisease in Ireland. Aquac. 93:273-278.Mialhe, E., E. Bachere, D. Chagot, and H. Grizel.1988. Isolation and purification of the protozoanBonamia ostreae (Pichot et al., 1980), aparasite affecting flat oyster Ostrea edulis L.Aquac. 71: 293-299.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35 p.124

M.3 MARTEILIOSIS(MARTEILIA REFRINGENS,M. SYDNEYI)M.3.1Background InformationM.3.1.1 Causative AgentsMarteiliosis is caused by two species of parasites,belonging to the Phylum Paramyxea.Marteilia refringens is responsible for Aber Disease(a.k.a Digestive Gland Disease) of Europeanoysters (Ostrea edulis) and Marteiliasydneyi is responsible for QX Disease ofSaccostrea glomerata (syn. Crassostreacommercialis, Saccostrea commercialis) and,possibly, Saccostrea echinata. More informationabout the disease can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000a).M.3.1.2 Host RangeOstrea edulis is infected by Marteilia refringens.Other host species include Tiostrea chilensis,Ostrea angasi, O. puelchana, Cerastoderma (=Cardium) edule, Mytilus edulis, M.galloprovincialis, Crassostrea gigas and C.virginica. Marteilia sydneyi infects Saccostreaglomerata and possibly S. echinata. Anothermarteiliad, Marteilia maurini, infects mussels(Mytilus edulis and M. galloprovincialis) fromFrance, Spain and Italy. This species is notreadily distinguished morphologically from M.refringens and distinct species status is underinvestigation. An unidentified marteiliad was responsiblefor mass mortalities of the Calico scallop(Argopecten gibbus) in Florida in the late1980’s, but has not re-appeared since. AnotherMarteilia-like species was reported from the giantclam, Tridacna maxima. Other species ofMarteilia that have been described include M.lengehi from Saccostrea (Crassostrea) cucullata(Persian Gulf and north Western Australia) andM. christenseni in Scrobicularia plana (France).These are differentiated from M. refringens andM. sydneyi by the cytoplasmic contents of thesporangia and spore morphology.M.3.1.3 Geographic DistributionMarteilia refringens is found in O. edulis in southernEngland, France, Italy, Portugal, Spain, Moroccoand Greece. Marteilia sydneyi is found inS. glomerata in Australia (New South Wales,Queensland and Western Australia).M.3.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System (1999-2000)No positive report of the disease in any countryfor the 2 year reporting periods. Most countrieshave no information about the occurrenceof the disease (OIE 1999,OIE 2000b).M.3.2 Clinical AspectsEarly stages of Marteilia refringens develop in thedigestive ducts, intestinal and stomach epitheliaand gills (Fig.M.3.2a). Later, spore-formingstages appear in the blind-ending digestive tubuleepithelia (Fig.M.3.2b). Proliferation of theparasite is associated with emaciation and exhaustionof glycogen reserves, grossdiscolouration of the digestive gland, cessationof feeding and weakening. Mortalities appear tobe associated with sporulation of the parasite anddisruption of the digestive tubule epithelia.M.3.3 Screening MethodsMore detailed methods for screening can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.M.3.3.1 PresumptiveM.3.3.1.1 Gross Observations (Level I)Slowed growth, gaping and mortalities of Ostreaedulis and other susceptible species should beconsidered suspect for Marteiliosis. Gross signsare not specific for Bonamiosis or Marteiliosis andrequire Level II examination.M.3.3.1.2 Tissue Imprints (Level II)Cut a cross-section through the digestive gland,blot away excess water with blotting paper anddab the cut section of the digestive gland onto aclean microscope slide. Fix tissue imprint for 2-3min in 70% methanol. Quick and effective stainingcan be achieved with a commercially availableblood-staining (cytological) kit, using themanufacturer’s instructions. The stained slidesare then rinsed (gently) under tap water, allowedto dry and cover-slipped using a synthetic resinmounting medium.The parasite morphology is as described for histology(M.3.3.2.1), although colouration may varywith the stain chosen. Initial screening with ahaematoxylin or trichrome stain, as used for his-125

M.3 Marteiliosis(Marteilia refringens, M. sydneyi)tology, may assist familiarisation with tissue imprintcharacteristics prior to using a dip-quickmethod. An observation time of 10 mins at 10-25x magnification is considered sufficient forscreening purposes.(SE McGladdery)M.3.3.2 ConfirmatoryM.3.3.2.1 Histopathology (Level II)Two dorso-ventral tissue section (2-3 mm thick)are recommended for screening purposes. Thesecan be removed from oysters over 18-24 monthsold (or >30 mm shell height) for immediate fixationin a fast fixative, such as 1G4F. Davidsonsor 10% buffered formalin may be used for largersamples or whole oysters (see M.1.3.3.3) butthese provide less satisfactory results if subsequentprocessing for Transmission Electron Microscopy(TEM) (M.3.4.2.1) is required (e.g., forspecies identification). Several standard stains(e.g., haematoxylin-eosin) enable detection ofMarteilia spp..The early stages of development occur in thestomach, intestine and digestive duct epithelia(usually in the apical portion of the cell) and appearas basophilic, granular, spherical inclusions(Fig.M.3.2a). Later stages occur in thedigestive tubules, where sporulation may inducehypertrophy of the infected cell. Marteiliaspp. spores contain eosinophilic, “refringent”,bodies which are easily detected at 10-25 xmagnification under light microscopy(Fig.M.3.2b).(SE McGladdery)Fig.M.3.2b. Digestive tubule of a Europeanoyster, Ostrea edulis, showing refringent sporestage of Marteilia refringens (star). Scale bar 50µm (H&E).(RD Adlard)Fig.M.3.4.1.1a. Tissue imprint from Saccostreacommercialis (Sydney rock oyster) heavily infectedby Marteilia sydneyi (arrows) (QX disease).Scale bar 250 µm (H&E).(RD Adlard)Fig.M.3.2a. Digestive duct of a European oyster,Ostrea edulis, showing infection of distalportion of the epithelial cells by plasmodia (arrows)of Marteilia refringens. Scale bar 15 µm(H&E).Fig.M.3.4.1.1b. Oil immersion of tissue squashpreparation of spore stages of Marteilia sydneyifrom Sydney rock oyster (Saccostreacommercialis) with magnified inset showing twospores within the sporangium. Scale bar 50 µm(H&E).126

M.3 Marteiliosis(Marteilia refringens, M. sydneyi)M.3.4 Diagnostic MethodsMore detailed methods for diagnosis can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.M.3.4.1 PresumptiveM.3.4.1.1 Tissue Imprints (Level II)As described under M.3.3.1.2, tissue imprintsmay also be used for presumptive diagnoses(Fig.M.3.4.1.1a,b). For first-time diagnoses backup tissues should be fixed for histology and EMconfirmatory diagnosis.M.3.4.1.2 Histopathology (Level II)Histology techniques as described underM.3.3.2.1, may be used. For first-time diagnosesa back up tissue specimen fixed for EMis recommended, as described below.M.3.4.2 ConfirmatoryM.3.4.2.1 Transmission Electron Microscopy(TEM) (Level III)TEM tissue preparation involves fixing tissueseither in 1G4F (M.1.3.3.3) or small (< 1 mmcubed) sub-samples of infected tissue in 2-3%glutaraldehyde mixed and buffered for ambientfiltered seawater. Ideally, fixation in 2-3%gluteraldehyde should not exceed 1 hr, sincelonger storage may induce membranous artifacts.Tissues should be fixed in 1G4F for 12-24 hrs.Following primary fixation, rinse tissues in a suitablebuffer and post-fix in 1-2% osmium tetroxide(OsO 4= osmic acid - highly toxic). Secondaryfixation should be complete within 1 hr. TheOsO 4fixative must also be rinsed with buffer/filtered(0.22 µm) seawater prior to dehydrationand resin-embedding.Post-fixed tissues can be stored in a compatiblebuffer or embedded post-rinsing in a resin suitablefor ultramicrotome sectioning. Screening of1 µm sections melted onto glass microscopeslides with 1% toluidine blue solution is onemethod of selecting the tissue specimens foroptimum evidence of possible Marteilia spp. Ultrathinsections are then mounted on copper grids(with or without formvar coating), and stained withlead citrate + uranyl acetate (or equivalent EMstain).Marteilia refringens plasmodia contain striatedinclusions, eight sporangial primordia, with upto four spores to each mature sporangium.Marteilia sydneyi has a thick layer of concentricmembranes surrounding the mature spore,lacks striated inclusions in the plasmodia, formseight to sixteen sporangial primordia in eachplasmodium and each sporangium containstwo (rarely three) spores.M.3.4.2.2 In situ Hybridization (Level III)In situ hybridization (Level III) techniques areunder development but not yet available commercially.Information on the current status ofthese and related molecular probe techniquesmay be obtained from IFREMER Laboratory atLa Tremblade, France (OIE 2000a, Annex MAI).M.3.5 Modes of TransmissionMarteilia refringens transmission appears to berestricted to periods when water temperaturesexceed 17°C. High salinities may impedeMarteilia spp. multiplication within the host tissues.Marteilia sydneyi also has a seasonal periodof transmission with infections occurringgenerally from mid- to late-summer (January toMarch). Heavy mortalities and sporulation occurall year round. The route of infection and life-cycleoutside the mollusc host are unknown. Since ithas not been possible to transmit the infectionexperimentally in the laboratory, an intermediatehost is suspected. This is reinforced by recentobservations showing spores do not survive morethan 7-10 days once isolated from the oyster.Cold temperatures prolong survival (35 days at15°C). Spore survival within fish or birds was limitedto 2 hrs, suggesting they are an unlikely modeof dispersal or transmission.M.3.6 Control MeasuresNone known. High salinities appear to suppressclinical manifestation of the disease, however,no eradication attempts have been successful,to date. Prevention of introduction or transfer ofoysters or mussels from Marteilia spp. enzooticwaters into historically uninfected waters is recommended.M.3.7 Selected ReferencesAnderson, T.J., R.D. Adlard, and R.J.G. Lester.1995. Molecular diagnosis of Marteilia sydneyi(Paramyxea) in Sydney rock oysters,Saccostrea commercialis (Angas). J. Fish Dis.18(6): 507-510.127

M.3 Marteiliosis(Marteilia refringens, M. sydneyi)Auffret, M. and M. Poder. 1983. Studies ofMarteilia maurini, parasite of Mytilus edulisfrom the north coasts of Brittany. Revue desTravaux de l’Institut des P?ches Maritimes,Nantes 47(1-2): 105-109.Berthe, F.C.J., M. Pernas, M. Zerabib, P. Haffner,A. Thebault, and A.J. Figueras. 1998. Experimentaltransmission of Marteilia refringenswith special consideration of its life cycle. Dis.Aquat. Org. 34(2): 135-144.Berthe, F.C.J., F. Le Roux, E. Peyretaillade, P.Peyret, D. Rodriguez, M. Gouy, and C.P.Vivares. 2000. Phylogenetic analysis of thesmall subunit ribosomal RNA of Marteiliarefringens validates the existence of PhylumParamyxea (Desportes and Perkins, 1990).J. Eukaryote Microbiol. 47(3): 288-293.Comps, M. 1983. Morphological study of Marteiliachristenseni sp.n., parasite of Scrobiculariapiperata P. (Mollusc, Pelecypod). Revue desTravaux de l’Institut des P?ches MaritimesNantes 47(1-2): 99-104.Hine, P.M. and T. Thorne. 2000. A survey of someparasites and diseases of several species ofbivalve mollusc in northern Western Australia.Dis. Aquat. Org. 40(1): 67 -78.Kleeman, S.N. and R.D. Adlard. 2000. Moleculardetection of Marteilia sydneyi, pathogen ofSydney rock oysters. Dis. Aquat. Org. 40(2):137-146.Montes, J., M.A. Longa, A. Lama, and A. Guerra.1998. Marteiliosis of Japanese oyster(Crassostrea gigas) reared in Galicia NWSpain. Bull. Europ. Soc. Fish Pathol. 18(4):124-126.Moyer, M.A., N.J. Blake, and W.S. Arnold. 1993.An ascetosporan disease causing mass mortalityin the Atlantic calico scallop, Argopectengibbus (Linnaeus, 1758). J. Shellfish Res.12(2): 305-310.Norton, J.H., F.O. Perkins, and E. Ledua. 1993.Marteilia-like infection in a giant clam, Tridacnamaxima, in Fiji. Journal of Invertebrate Pathology61(3): 328-330.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Renault, T., N. Cochennec, and B. Chollet. 1995.Marteiliosis in American oysters Crassostreavirginica reared in France. Dis. Aquat. Org.23(3): 161-164.Robledo, J.A.F. and A. Figueras. 1995. The effectsof culture-site, depth, season and stocksource on the prevalence of Marteiliarefringens in cultured mussels (Mytilusgalloprovincialis) from Galicia, Spain. J.Parasitol. 81(3): 354-363.Roubal, F.R., J. Masel, and R.J.G. Lester. 1989.Studies on Marteilia sydneyi, agent of QX diseasein the Sydney rock oyster, Saccostreacommercialis, with implications for its life-cycle.Aust. J. Mar. Fresh. Res. 40(2): 155-167.Villalba, A., S.G. Mourelle, M.C. Lopez, M.J.Caraballal, and C. Azevedo. 1993. Marteiliasisaffecting cultured mussels Mytilusgalloprovincialis of Galicia (NW Spain). 1. Etiology,phases of the infection, and temporaland spatial variability in prevalence. Dis. Aquat.Org. 16(1): 61-72.Villalba, A., S.G. Mourelle, M.J. Carballal, andC. Lopez. 1997. Symbionts and diseases offarmed mussels Mytilus galloprovincialisthroughout the culture process in the Rias ofGalicia (NW Spain). Dis. Aquat. Org. 31(2):127-139.Wesche, S.J., R.D. Adlard, and R.J.G. Lester.1999. Survival of spores of the oyster pathogenMarteilia sydneyi (Protozoa, Paramyxea)as assessed using fluorogenic dyes. Dis.Aquat. Org. 36(3): 221-226.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.128

M.4 MIKROCYTOSIS(MIKROCYTOS MACKINI,M. ROUGHLEYI)M.4.1 Background InformationM.4.1.1 Causative AgentsMikrocytosis is caused by two species of parasitesof uncertain taxonomic affinity. Mikrocytosmackini is reponsible for Denman Island Disease(Microcell disease) of Pacific oysters(Crassostrea gigas), and Mikrocytos roughleyi isresponsible for Australian Winter Disease (WinterDisease, Microcell Disease) of Sydney rockoysters, Saccostrea glomerata. More informationabout the disease can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000a).M.4.1.2 Host RangeMikrocytos mackini naturally infects Crassostreagigas (Pacific oysters). Ostrea edulis (Europeanoysters), O. conchaphila (= O. lurida) (Olympiaoyster) and Crassostrea virginica (American oysters)growing in enzootic waters are also susceptibleto infection. Saccostrea glomerata(Crassostrea commercialis, Saccostreacommercialis) (Sydney rock oyster) is the onlyknown host for Mikrocytos roughleyi.M.4.1.3 Geographic DistributionMikrocytos mackini is restricted to specific localitiesaround Vancouver Island and southwestcoast of the Pacific coast of Canada. The parasiteis limited to waters with temperatures below12?C. Mikrocytos roughleyi occurs in central tosouthern New South Wales, and at Albany andCarnarvon, Western Australia.M.4.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System (1999-2000)No positive report during the reporting periodsfor 1999 and 2000. In Australia, last year of occurrencewas 1996 (in New South Wales andWestern Australia. Most countries have no informationabout occurrence of the disease (OIE1999, OIE 2000b).M.4.2 Clinical AspectsMikrocytos mackini initiates focal infections of thevesicular connective tissue cells. This elicitshaemocyte infiltration and abscess formation.Grossly visible pustules (Fig.M.4.2a), abscesslesions and tissue ulcers, mainly in the mantle,may correspond to brown scar formation on theadjacent surface of the inner shell. However,such lesions are not always present. Small cells,1-3 µm in diameter, are found (rarely) aroundthe periphery of advanced lesions, or in connectivetissue cells in earlier stages of diseasedevelopment. Severe infections appear to berestricted to oysters over 2 years old.Mikrocytos roughleyi induces a systemic intracellularinfection of the haemocytes (never theconnective tissue cells) which may result in focallesions in the gills, connective, gonadal anddigestive tract.M.4.3 Screening MethodsMore detailed methods for screening can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.M.4.3.1 PresumptiveM.4.3.1.1 Gross Observations (Level I)Slowed growth, gaping and mortalities ofCrassostrea gigas and Saccostrea glomeratashould be considered suspect for Mikrocytosis.Gross signs are not pathogen specific and requireLevel II examination, at least for first-timeobservations.M.4.3.1.2 Cytological Examination andTissue Imprints (Level II)Heart impressions (dabs) can be made onto aclean microscope slide and air-dried. Once dry,the slide is fixed in 70% methanol. Quick andeffective staining can be achieved with commerciallyavailable blood-staining (cytological) kits,using the manufacturer’s instructions. Thestained slides are then rinsed (gently) under tapwater, allowed to dry and cover-slipped using asynthetic resin mounting medium. Intracytoplasmicparasites in the haemocytes will match thedescriptions given above for histology. This techniqueis more applicable to M. roughleyi than M.mackini.Tissue sections through mantle tissues (especiallyabscess/ulcer lesions, where present) arecut and excess water removed with blotting paper.The cut section is dabbed onto a clean microscopeslide, fixed for 2-3 minutes in 70%methanol and stained. Quick and effective stainingcan be achieved using a commercially availableblood-staining (cytological) kits, using themanufacturer’s instructions. The stained slides129

M.4 MIKROCYTOSIS(Mikrocytos mackini, M. roughleyi)(SM Bower)are then rinsed (gently) under tap water, allowedto dry and cover-slipped using a synthetic resinmounting medium.The parasite morphology is as described for histology(M.4.3.2.1), although colouration mayvary with the stain chosen. Initial screening witha haematoxylin or trichrome stain, as used forhistology, may assist familiarisation with tissueimprint characteristics prior to using a dip-quickmethod. An observation time of 10 mins underoil immersion is considered sufficient for screeningpurposes.Fig.M.4.2a. Gross abscess lesions (arrows) inthe mantle tissues of a Pacific oyster(Crassostrea virginica) severely infected byMikrocytos mackini (Denman Island Disease).(SM Bower)Fig.M.4.3.2.1a. Histological section throughmantle tissue abscess corresponding to thegross lesions pictured in Fig.M.4.2a, in a Pacificoyster (Crassostrea gigas) infected byMikrocytos mackini (H&E).(SM Bower)Fig.M.4.3.2.1b. Oil immersion of Mikrocytosmackini (arrows) in the connective tissue surroundingthe abscess lesion pictured inFig.M.4.3.2.1a. Scale bar 20 µm (H&E).M.4.3.2 ConfirmatoryM.4.3.2.1 Histopathology (Level II)It is recommended that at least two dorso-ventralsections (2-3 mm) through each oyster beexamined using oil immersion for screening purposes.Sections from oysters >2 yrs (or >30 mmshell height) should be fixed immediately in afast fixative such as 1G4F. Davidsons or 10%buffered formalin may be used for smaller orwhole oysters (see M.1.3.3.3) but these fixativesare not optimal for subsequent confirmatoryElectron Microscopy (EM) diagnosis(M.4.4.2.1), or species identification, if required.Also smaller oysters are not the recommendedsize-group for Mikrocytos screening. Sectionsthrough pustules, abscess or ulcer lesionsshould selected where present. Several standardstains (e.g., haematoxylin-eosin) enabledetection of Mikrocytos spp.Mikrocytos mackini appears as 2-3 µm intracellularinclusions in the cytoplasm of the vesicularconnective tissues immediately adjacentto the abscess-like lesions (Fig. M.4.3.2.1a,b).It may also be observed in muscle cells and,occasionally, in haemocytes or free, within thelesions. It is distinguished from Bonamia by aneccentric nucleus and from M. roughleyi by theconsistent absence of a cytoplasmic vacuole,and the presence of a mitochondrion in M.roughleyi. These features will not be clear underoil and require confirmation using 1 micronresin sections or TEM (described below). Neitherof these techniques, however, are practicalfor screening purposes.Mikrocytos roughleyi measures 1-3 µm in diameterand occurs exclusively in thehaemocytes. A cytoplasmic vacuole may ormay not be present. When present, it displacesthe nucleus peripherally. Nucleolar structuresmay or may not be visible under oil immersionresolution for this intracellular parasite.130

M.4 Mikrocytosis(Mikrocytos mackini, M. roughleyi)M.4.4 Diagnostic MethodsMore detailed methods for diagnosis can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.M.4.4.1 PresumptiveM.4.4.1.1 Histopathology and Tissue Imprints(Level II)Histology (M.4.3.2.1) may be used, however, forfirst-time diagnoses, EM confirmation is recommended(M.4.4.2.2). Tissue imprints may also beused for presumptive diagnoses, where theydemonstrate the features described underM.4.3.1.2.M.4.4.2 ConfirmatoryM.4.4.2.1 Histopathology and Tissue Imprints(Level II)Histology (M.4.3.2.1) and tissue imprints(M.4.3.1.2) may be used, however, for first-timediagnoses, EM confirmation is recommended(M.4.4.2.2).M.4.4.2.2 Transmission Electron Microscopy(TEM) (Level III)Tissues should be fixed in 1G4F for 12-24 hours.Following primary fixation, rinse tissues in a suitablebuffer and post-fix in 1-2% osmium tetroxide(OsO 4= osmic acid - highly toxic). Secondaryfixation should be complete within 1 hour. TheOsO 4fixative must also be rinsed with buffer/filtered(0.22 microns) seawater prior to dehydrationand resin-embedding.Post-fixed tissues can be stored in a compatiblebuffer or embedded post-rinsing in a resin suitablefor ultramicrotome sectioning. Screening of1 micron sections melted onto glass microscopeslides with 1% toludine blue solution is onemethod of selecting the tissue specimens foroptimum evidence of putative Mikrocytos spp.Ultrathin sections are then mounted on coppergrids (with or without formvar coating), andstained with lead citrate + uranyl acetate (orequivalent EM stain).Mikrocytos mackini is distinguished ultrastructurally(as well as by tissue location and host species)from Bonamia spp. by the location of thenucleolus. In M. mackini it is in the centre of thenucleus, while in B. ostreae it is eccentric.Mikrocytos mackini also lacks mitochondria.The ultrastructural characteristics of Mikrocytosroughleyi have not been published, however, itis distinguished from M. mackini by the presenceof a cytoplasmic vacuole (along with completelydifferent geographic, host and tissue locations!).M.4.5 Modes of TransmissionMikrocytos mackini transmission appears restrictedto early spring (April-May) following periodsof 3-4 months at water temperatures

M.4 Mikrocytosis(Mikrocytos mackini, M. roughleyi)Mikrocytos mackini (sp. n.) and Mikrocytosroughleyi (sp. n.). Fish. Bull. 86(3): 581 -594.Hervio, D., S.M. Bower, and G.R. Meyer. 1995.Life-cycle, distribution and lack of host specificityof Mikrocytos mackini, the cause ofDenman Island disease of Pacific oysters(Crassostrea gigas). J. Shellfish Res. 14(1):228 (abstract).Hervio, D., S.M. Bower, and G.R. Meyer. 1996.Detection, isolation and experimental transmissionof Mikrocytos mackini, a microcellparasite of Pacific oysters Crassostrea gigas(Thunberg). J. Inverteb. Pathol. 67(1): 72-79.Lester, R.J.G. 1990. Diseases of cultured molluscsin Australia. Advances in Tropical Aquaculture:Workshop, Tahiti, French Polynesia Feb.20 – Mar. 4, 1989. Actes de colloquesIFREMER 9: 207-216.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Smith, I.R., J.A. Nell, and R.D. Adlard. 2000. Theeffect of growing level and growing method onwinter mortality, Mikrocytos roughleyi, in diploidand triploid Sydney rock oysters,Saccostrea glomerulata. Aquac. 185(3-4): 197-205.132

M.5 PERKINSOSIS(PERKINSUS MARINUS, P. OLSENI)M.5.1 Background InformationM.5.1.1 Causative AgentsPerkinsosis is caused by two species of protistanparasite belonging to the phylum Apicomplexa(although recent nucleic acid investigations suggesta possible affiliation with the dinoflagellates).Perkinsus marinus is responsible for “Dermo”disease in Crassostrea virginica (American oysters)and Perkinus olseni causes perkinsosis inmany bivalve species in tropical and subtropicalwaters. Other perkinsiid species are known toinfect clams in Europe (Perkinsus atlanticus) andthe eastern USA (Perkinsus spp.), as well asJapanese (Yesso) scallops, Patinopectenyessoensis in Pacific Canada (Perkinsusqugwadi). The taxonomic relationship betweenthese and the two species listed as ‘notifiable’ byOIE is currently under investigation. More informationabout the disease can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000a).M.5.1.2 Host RangePerkinsus marinus (formerly known asDermocystidium marinum and Labyrinthomyxamarinus) infects Crassostrea virginica (Americanoysters). Experimental infection to C. gigas (Pacificoysters) is possible, but they appear moreresistant than C. virginica. Perkinsus olsenishows a strong rDNA similarity to Perkinsusaltanticus of Ruditapes decussatus and the speciationwithin this genus, as mentioned underM.5.1.1, is currently under nucleic acid investigation.Recognised hosts of P. olseni are theabalone species: Haliotis rubra, H. cyclobates,H. scalaris and H. laevigata. More than 50 othermolluscan species harbour Perkinsus spp., aswell as other possibly related species, withoutapparent harmful effects (e.g., in Arca clams[Fig.M.5.1.2a] and Pinctada pearl oysters[Fig.M.5.1.2b]).M.5.1.3 Geographic DistributionPerkinsus marinus is found along the east coastof the United States from Massachusetts toFlorida, along the Gulf of Mexico coast to Venezuela,and in Puerto Rico, Cuba and Brazil. Ithas also been introduced into Pearl Harbour,Hawaii. Range extension into Delaware Bay, NewJersey, Cape Cod and Maine are attributed torepeated oyster introductions and increased winterwater temperatures. Perkinsus olseni occursin South Australia. Other species occur in Atlanticand Pacific oceans and the MediterraneanSea.M.5.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System (1999-2000)P. marinus was not reported in Australia during1999 and 2000 reporting periods; P. olseni likewisenot reported in 1999 and 2000 (last year ofoccurrence in South Australia in 1997, and in1995 in New South Wales and Western Australia).Suspected in Korea RO for reporting period1999 and 2000. In New Zealand, positively reportedfrom April to December 2000. Perkinsusolseni occurs in wild populations of New Zealandcockles, Austrovenus stutchburyi (FamilyVeneridae) and two other bivalve species,Macomona liliana (Family Tellinidae) and Barbatianovae-zelandiae (Family Arcidae). These speciesoccur widely in the coast of New Zealand.Affected locations have been the Waitemata andKaipara Harbours but the organism is probablyenzootic in the warmer waters of northern NewZealand (OIE 1999, OIE 2000a).M.5.2 Clinical AspectsThe effects of Perkinsus marinus on Crassostreavirginica range from pale appearance of the digestivegland, reduced condition indices, severeemaciation, gaping, mantle retraction, retardedgonadal development and growth and occasionalabscess lesions. Mortalities of up to 95% haveoccurred in infected C. virginica stocks.Proliferation of Perkinsus olseni results in disruptionof connective and epithelial tissues and somehost species show occasional abscess formation.Pustules up to 8 mm in diameter in affectedHaliotis spp. reduce market value and have beenassociated with heavy losses in H. laevi gata.M.5.3 Screening MethodsMore detailed methods for screening can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.M.5.3.1 PresumptiveM.5.3.1.1 Gross Observations (Level I)Slowed growth, gaping and mortalities ofCrassostrea virginica and Haliotis spp., as wellas other mollusc species in Perkinsus-enzooticwaters should be considered suspect forPerkinsiosis. Gross signs are not pathogen-specificand require Level II examination, at least forfirst time observations.133

M.5 Perkinsosis(Perkinsus marinus, P. olseni)(PM Hine)(SE McGladdery)Fig.M.5.1.2a. Arca clam showing a Perkinsuslikeparasite within the connective tissue. Magnifiedinsert shows details of an advanced ‘schizont’like stage with trophozoites showingvacuole-like inclusions. Scale bar 100 µm.(H&E).(PM Hine)Fig.M.5.3.2.1b. Schizont (‘rosette’) stages ofPerkinsus marinus (arrows), the cause of‘Dermo’ disease in American oyster(Crassostrea virginica) digestive gland connectivetissue. Scale bar 30 µm (H&E).(SE McGladdery)Fig.M.5.1.2b. Pinctada albicans pearl oystershowing a Perkinsus-like parasite. Magnifiedinsert shows details of a ‘schizont’-like stagecontaining ‘trophozoites’ with vacuole-like inclusions.Scale bar 250 µm (H&E).(SM Bower)Fig.5.3.2.2. Enlarged hypnospores of Perkinsusmarinus stained blue-black with Lugol’s iodinefollowing incubation in fluid thioglycollate medium.Scale bar 200 µm.M.5.3.2 ConfirmatoryM.5.3.2.1 Histopathology (Level II)Fig.M.5.3.2.1a. Trophozoite (‘signet-ring’)stages of Perkinsus marinus (arrows), the causeof ‘Dermo’ disease in American oyster(Crassostrea virginica) connective tissue. Scalebar 20 µm (H&E).It is recommended that at least two dorso-ventralsections through each oyster be examinedusing oil immersion for screening purposes. Sectionsfrom oysters >2 yrs (or >30 mm shell height)should be fixed immediately in a fast fixative suchas 1G4F. Davidson’s or 10% buffered formalinmay be used for smaller or whole oysters (seeM.1.3.3.3) but these fixatives are not optimal forsubsequent confirmatory Electron Microscopy(EM) diagnosis (M.5.4.2.1), or species identification,if required. Sections through pustules,abscess or ulcer lesions should be selected,where present. Several standard stains (e.g.,134

M.5 Perkinsosis(Perkinsus marinus, P. olseni)haematoxylin-eosin) enable detection ofPerkinsus spp.Perkinsus marinus infections are usually systemic,with trophozoites occuring in the connectivetissue of all organs. Immature trophozoites(meronts, merozoites or aplanospores) measure2-3 µm in diameter. “Signet-ring” stages are maturetrophozoites, measuring 3-10 µm in diameter,each with a visible eccentric vacuole displacingthe nucleus and cytoplasm peripherally(Fig.M.5.3.2.1a). The “rosette” stage(tomonts, sporangia or schizonts) measure 4-15 µm in diameter and can contain 2, 4, 8, 16or 32 developing trophozoites (Fig.M.5.3.2.1b).Perkinsus olseni shows the same developmentalstages although the trophozoite stages arelarger ranging from 13-16 µm in diameter. Dueto host and parasite diversity, however, morphologicalfeatures cannot be considered specific.M.5.3.2.2 Fluid Thioglycollate Culture(Level II)Tissue samples measuring 5-10 mm are excised(select lesions, rectal and gill tissues) and placedin fluid thioglycollate medium containing antibiotics.Incubation temperature and time varies perhost species and environment. The standard protocolfor P. marinus is 22-25˚C for 4-7 days inthe dark. Warmer temperatures can be used forP. olseni.The cultured parasites expand in size to 70-250?m in diameter. Following incubation, the tissuesand placed in a solution of 1:5 Lugol’s iodine:waterfor 10 minutes. The tissue is then teased aparton a microscope slide and examined, using lowpower on a light microscope, for enlargedhypnospores with walls that stain blue-black(Fig.M.5.3.2.2).M.5.4 Diagnostic MethodsMore detailed methods for diagnostics can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.M.5.4.1 PresumptiveM.5.4.1.1 Histopathology (Level II)M.5.4.1.2 Fluid Thioglycollate MediumCulture (Level II)Fluid thioglycollate medium culture (Level II) mayalso be used for presumptive diagnosis(M.5.3.2.2).M.5.4.2 ConfirmatoryM.5.4.2.1 Transmission Electron Microscopy(TEM) (Level III)TEM is required to detect the species-specificultrastructure of the zoospore stage of development(collected from zoospore release from culturedaplanospores [prezoosporangia]). Tissuepreparation involves fixing concentratedzoospores or zoosporangia (produced by placingthe aplanospores into filtered ambient seawater,where they develop into zoosporangia andproduce hundreds of motile zoospores) in 2-3%glutaraldehyde mixed and buffered for ambientfiltered seawater. Oyster tissues can also be fixedin 1G4F for 12-24 hrs. Following primary fixation,rinse tissues in a suitable buffer and postfixin 1-2% osmium tetroxide (OsO 4= osmic acid- highly toxic). Secondary fixation should be completewithin 1 hr. The OsO 4fixative must also berinsed with buffer/filtered (0.22 µm) seawaterprior to dehydration and resin-embedding.Post-fixed tissues can be stored in a compatiblebuffer or embedded post-rinsing in a resin suitablefor ultramicrotome sectioning. Screening of1 ?m-thick sections melted onto glass microscopeslides with 1% toluidine blue solution isone method of selecting the tissue specimensfor optimum evidence of putative Perkinsus spp.Such pre-screening should not be necessary forconcentrated zoospore or zoosporangia preparations.Ultrathin sections are then mounted oncopper grids (with or without formvar coating),and stained with lead citrate + uranyl acetate (orequivalent EM stain).The anterior flagellum of Perkinsus marinuszoospores is ornamented with a row of hair- andspur-like structures. The posterior flagellum issmooth. An apical complex is present, consistingof a conoid, sub-pellicular microtubules,rhoptries, rectilinear micronemes and conoidassociatedmicronemes. Large vacuoles are alsopresent at the anterior end of the zoospore.Histology (M.5.3.2.1), may be used. However,for first-time diagnoses a back up tissue specimenfixed for EM is recommended (M.5.4.2.1).135

M.5 Perkinsosis(Perkinsus marinus, P. olseni)M.5.5 Modes of TransmissionProliferation of Perkinsus spp. is correlated withwarm water temperatures (>20˚C) and this coincideswith increased clinical signs and mortalities.Effects appear cumulative with mortalitiespeaking at the end of the warm water seasonin each hemisphere. The infective stage isa biflagellate zoospore which transforms intothe feeding trophozoite stage after entering thehost’s tissues. These multiply by binary fissionwithin the host tissues. Perkinsus marinusshows a wide salinity tolerance range. Perkinsusolseni is associated with full strength salinityenvironments.Direct transmission of Perkinsus spp. has beendemonstrated by exposure of susceptible hoststo infected hosts, including cross-species transmissionfor P. olseni. There is currently no evidenceof cross-genus transmission of P. marinus.M.5.6 Control MeasuresNone known for Perkinsus spp. Most effortsagainst P. marinus have concentrated on developmentof resistant (tolerant) stocks of oysters.These currently show potential for surviving inenzootic areas, but are not recommended for usein non-enzootic areas due to their potential to actas sub-clinical carriers of the pathogen. Somesuccess has been achieved, however, in preventingP. marinus infection of hatchery-reared larvaland juvenile oysters using filtration and UV sterilizationof influent water. The almost ubiquitousoccurrence of Perkinsus in many bivalve speciesaround mainland Australia makes control byrestriction of movements impractical.M.5.7 Selected ReferencesAlemida, M., F.C. Berthe, A. Thebault, and M.T.Dinis. 1999. Whole clam culture as a quantitativediagnostic procedure of Perkinsusatlanticus (Apicomplexa, Perkinsea) in clamsRuditapes decussatus. Aquac. 177(1-4): 325-332.Blackbourn, J., S.M. Bower, and G.R. Meyer.1998. Perkinsus qugwadi sp. nov. (incertaecedis), a pathogenic protozoan parasite ofJapanese scallops, Patinopecten yesssoensis,cultured in British Columbia, Canada. Can. J.Zool. 76(5): 942-953.Bower, S.M., J. Blackbourn, and G.R. Meyer.1998. Distribution, prevalence and pathogenicityof the protozoan Perkinsus qugwadiin Japanese scallops, Patinopectenyesssoensis, cultured in British Columbia,Canada. Can. J. Zool. 76(5): 954-959.Bower, S.M., J. Blackbourne, G.R. Meyer, andD.W. Welch. 1999. Effect of Perkinsusqugwadi on various species and strains ofscallops. Dis. Aquat. Org. 36(2): 143-151.Canestri-Trotti, G., E.M. Baccarani, F. Paesanti,and E. Turolla. 2000. Monitoring of infectionsby Protozoa of the genera Nematopsis,Perkinsus and Porospora in the smooth venusclam Callista chione from the north-westernAdriatic Sea (Italy). Dis. Aquat. Org. 42(2):157-161.Cook, T., M. Folli, J. Klinck, S.E. Ford, and J.Miller. 1998. The relationship between increasingsea-surface temperature and the northwardspread of Perkinsus marinus (Dermo) diseaseepizootics in oysters. Estuarine, Coastal andShelf Science 46(4): 587 -597.Fisher, W.S., L.M. Oliver, L., W.W. Walker, C.S.Manning and T.F. Lytle. 1999. Decreased resistanceeastern oysters (Crassostreavirginica) to a protozoan pathogen (Perkinsusmarinus) after sub-lethal exposure to tributyltinoxide. Mar. Environ. Res. 47(2): 185-201.Ford, S.E., R. Smolowitz, and M.M. Chintala.2000. Temperature and range extension byPerkinsus marinus. J. Shellfish Res. 19(1): 598(abstract).Ford, S.E., Z. Xu, and G. Debrosse. 2001. Useof particle filtration and UV radiation to preventinfection by Haplosporidium nelsoni (MSX) andPerkinsus marinus (Dermo). Aquac. 194(1-2):37-49.Hine, P.M. and T. Thorne. 2000. A survey of someparasites and diseases of several species ofbivalve mollusc in northern Western Australia.Dis. Aquat. Org. 40(1): 67 -78.Kotob, S.I., S.M. McLaughlin, P. van Berkum,and M. Faisal. 1999. Discrimination betweentwo Perkinsus spp. isolated from the soft shellclam, Mya arenaria, by sequence analysis oftwo internal transcribed spacer regions and the5.8S ribosomal RNA gene. Parasitol. 119(4):363-368.Kotob, S.I., S.M. McLaughlin, P. van Berkum, P.and M. Faisal. 1999. Characterisation of twoPerkinsus spp. from the soft shell clam, Mya136

M.5 Perkinsosis(Perkinsus marinus, P. olseni)arenaria, using the small subunit ribosomalRNA gene. J. Eukaryotic Microbiol. 46(4):439-444.McLaughlin, S.M. and M. Faisal. 1998. In vitropropagation of two Perkinsus spp. from thesoft shell clam Mya arenaria. Parasite 5(4):341-348.McLaughlin, S.M. and M. Faisal. 1998. Histopathologicalalternations associated withPerkinsus spp. infection in the soft shell clamMya arenaria. Parasite 5(4): 263-271.McLaughlin, S.M. and M. Faisal. 1999. A comparisonof diagnostic assays for detection ofPerkinsus spp. in the soft shell clam Myaarenaria. Aquac. 172(1-2): 197-204.McLaughlin, S.M. and M. Faisal. 2000. Prevalenceof Perkinsus spp. in Chesapeake Baysoft-shell clams, Mya arenaria Linnaeus, 1758,during 1990-1998. J. Shellfish Res. 19(1): 349-352.Nickens, A.D., E. Wagner, and J.F. LaPeyre.2000. Improved procedure to count Perkinsusmarinus in eastern oyster hemolymph. J.Shellfish Res. 19(1): 665 (abstract).clams naturally infected with Perkinsusatlanticus. Fish and Shellfish Immunol. 10(7):597-609.Park, K-I., K-S. Choi, and J-W. Choi. 1999. Epizootiologyof Perkinsus sp. found in Manilaclam, Ruditapes philippinarum in KomsoeBay, Korea. J. Kor. Fish. Soc. 32(3): 303-309.Robledo, J.A.F., J.D. Gauthier, C.A. Coss, A.C,Wright, G.R. Vasta. 1999. Species -specificityand sensitivity of a PCR-based assay forPerkinsus marinus in the eastern oyster,Crassostrea virginica: A comparison with thefluid thioglycollate assay. J. Parasitol. 84(6):1237-1244.Robledo, J.A.F., C.A. Coss, and G.R. Vasta.2000. Characterization of the ribosomal RNAlocus of Perkinsus atlanticus and developmentof a polymerase chain reaction -based diagnosticassay. J. Parasitol. 86(5): 972-978.Yarnall, H.A., K.S. Reece, N.A. Stokes, and E.M.Burreson. 2000. A quantitative competitivepolymerase chain reaction assay for the oysterpathogen Perkinsus marinus. J. Parasitol.86(4): 827-837.O’Farrell, C.L., J.F. LaPeyre, K.T. Paynter, andE.M. Burreson. 2000. Osmotic tolerance andvolume regulation in in vitro cultures of theoyster pathogen Perkinsus marinus. J. ShellfishRes. 19(1): 139-145.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Ordas, M.C. and A. Figueras. 1998. In vitro cultureof Perkinsus atlanticus, a parasite of thecarpet shell clam Ruditapes decussatus. Dis.Aquat. Org. 33(2): 129-136.Ordas, M., A. Ordas, C. Beloso, and A. Figueras.2000. Immune parameters in carpet shell137

M.6 HAPLOSPORIDIOSIS(HAPLOSPORIDIUM COSTALE,H. NELSONI)M.6.1 Background Information(PM Hine)M.6.1.1 Causative AgentsHaplosporidiosis is caused by two species ofprotistan parasite belonging to the phylumHaplosporidia. Haplosporidium nelsoni (syn.Minchinia nelsoni) is responsible for “MSX” (multinucleatesphere X) disease in Crassostreavirginica (American oysters) and Haplosporidiumcostale (Minchinia costale) causes “SSO” (seasideorganism) disease in the same species.More information about the disease can be foundin the OIE Diagnostic Manual for Aquatic AnimalDiseases (OIE 2000a).M.6.1.2 Host RangeBoth Haplosporidium nelsoni and H. costalecause disease in Crassostrea virginica (Americanoysters). Recently a Haplosporidium sp. fromCrassostrea gigas (Pacific oyster) has been identifiedas H. nelsoni using DNA sequencing ofsmall sub-unit ribosomal DNA.Fig.M.6.1.3b. Oil immersion magnification ofthe operculated spore stage of theHaplosporidium-like parasite in the gold-lippedpearl oyster Pinctada maxima from north WesternAustralia. Scale bar 10 µm. (H&E).(PM Hine)M.6.1.3 Geographic DistributionHaplosporidium nelsoni occurs in American oystersalong the Atlantic coast of the United Statesfrom northern Florida to Maine. Enzootic areasappear limited to Delaware Bay, ChesapeakeBay, Long Island Sound and Cape Cod.Haplosporidium nelsoni has been found in C. gigasfrom California and Washington on the Pacificcoast of the USA, and Korea, Japan andFrance.(PM Hine)Fig.M.6.1.3c. Haemocyte infiltration activity inthe connective tissue of a Sydney rock oyster(Saccostrea cucullata) containing spores of aHaplosporidium-like parasite (arrow). Scale bar0.5 mm. (H&E).(PM Hine)Fig.M.6.1.3a. Massive connective tissue anddigestive tubule infection by an unidentifiedHaplosporidium-like parasite in the gold-lippedpearl oyster Pinctada maxima from north WesternAustralia. Scale bar 0.5 mm (H&E).Fig.M.6.1.3d. Oil immersion magnification ofHaplosporidium-like spores (arrow) associatedwith heavy haemocyte infiltration in a Sydneyrock oyster (Saccostrea cucullata). Scale bar10 µm. (H&E).138

M.6 Haplosporidiosis(Haplosporidium costale, H. nelsoni)Haplosporidium costale has been reportedsolely from C. virginica from the Atlantic coastof the United States and has a small subunitrDNA distinct from that of H. nelsoni.Hapolosporidium costale also has a narrowerdistribution, ranging from Long Island Sound,New York, to Cape Charles, Virginia.Similar agents have been reported from hatchery-rearedpearl oysters, Pinctada maxima,(Fig.M.6.1.3a,b) and the rock oyster, Saccostreacucullata (Fig.M.6.1.3c,d), from north WesternAustralia.M.6.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System (1999-2000)No information or no positive report for this diseasein any country for the reporting periods 1999and 2000 (OIE 1999, OIE 2000b).M.6.2 Clinical AspectsHaplosporidium nelsoni occurs extracellularlyin the connective tissue and digestive gland epithelia.It is often associated with a visible brownreddiscolouration of gill and mantle tissues.Sporulation of H. nelsoni is prevalent in juvenileoysters (1-2 yrs) but sporadic in adults and occursexclusively in the epithelial tissues of thedigestive tubules. Sporulation of H. costale occursthroughout the connective tissues.Haplosporidum nelsoni infections appear andcontinue throughout the summer (mid-May to theend of October). Gradual disruption of the digestivegland epithelia is associated with weakeningand cumulative mortalities of oysters. A secondwave of mortalities may occur in early springfrom oysters too weak to survive over-wintering.Holding in vivo for up to 2 weeks in 10 ppt salinityseawater at 20˚C kills the parasite but not thehost. H. nelsoni does not cause disease at

M.6 Haplosporidiosis(Haplosporidium costale, H. nelsoni)(SE McGladdery)(SE McGladdery)Fig.M.6.3.1.2a. Plasmodia (black arrows) andspores (white arrows) of Haplosporidiumcostale, the cause of SSO disease, throughoutthe connective tissue of an American oyster(Crassostrea virginica). Scale bar 50 µm.(SE McGladdery)Fig.M.6.4.2.2b. Oil immersion magnification ofMSX spores in the digestive tubule epitheliumof an American oyster Crassostrea virginica.Scale bar 25 µm. (H&E).techniques used are the same as described forconfirmatory diagnosis (M.6.4.2.2).M.6.4 Diagnostic MethodsMore detailed methods for diagnostics can befound in the OIE Diagnostic Manual for AquaticAnimal Diseases (OIE 2000a), at http://www.oie.int or selected references.M.6.4.1 PresumptiveM.6.4.1.1 Gross Observations (Level I)Fig.M.6.3.1.2b. Plasmodia (black arrows) andspores (white arrows) of Haplosporidiumnelsoni, the cause of MSX disease, throughoutthe connective tissue and digestive tubules ofan American oyster (Crassostrea virginica).Scale bar 100 µm.(SE McGladdery)The only presumptive diagnosis would be grossobservations of cumulative mortalities of Americanoysters in early spring and late summer inareas (12-25 ppt salinity) with an established historyof MSX epizootics. Such presumptive diagnosisrequires confirmation via another diagnostictechnique (histology). Likewise, summermortalities of the same oyster species in waterswith a history of SSO disease may be presumedto be due to SSO. Both must be confirmed, however,since infection distributions for bothspecies of Haplosporidium may overlap.M.6.4.2 ConfirmatoryM.6.4.2.1 Cytological Examination andTissue Imprints (Level II)Fig.M.6.4.2.2a. Oil immersion magnification ofSSO spores in the connective tissue of anAmerican oyster Crassostrea virginica. Scalebar 15 µm.Positive cytological or tissue imprints (M.6.4.1.1)can be considered confirmatory where collectedfrom susceptible oyster species and areas witha historic record of the presence ofHaplosporidium spp. infections.140

M.6 Haplosporidiosis(Haplosporidium costale, H. nelsoni)M.6.4.2.2 Histopathology (Level II)Positive histological sections can be consideredconfirmatory where collected from susceptibleoyster species and areas with a historicrecord of the presence of Haplosporidium spp.infections.It is recommended that at least two dorso-ventralsections through each oyster be examinedusing oil immersion for screening purposes. Sectionsfrom oysters >2 yrs (or >30 mm shellheight) should be fixed immediately in a fastfixative such as 1G4F. Davidson’s or 10% bufferedformalin may be used for smaller or wholeoysters (see M.1.3.3.3) but these fixatives arenot optimal for subsequent confirmatory ElectronMicroscopy (EM) diagnosis (M.6.4.2.3), orspecies identification, if required. Several standardstains (e.g., haemotoxylin-eosin) enabledetection of Haplosprodium spp..Haplosporidium spp. infections are usually systemicand characterised by massive infiltrationby hyalinocyte haemocytes (agranularhaemocytes with a low cytoplasm: nucleoplasmratio). The sporoplasm of spores of H. costalewhich are smaller than those of MSX and oftenmasked by the intense haemocyte infiltration response,can be differentially stained using amodified Ziehl-Nielsen stain. Sporocyts of H.costale occur in the connective tissues(Fig.M.6.3.1.2a), measure approximately 10-25µm in diameter and contain oval, operculate,spores approximately 3 µm in size(Fig.M.6.4.2.2a). Sporocysts of H. nelsoni occurin the digestive tubule epithelia and measure20-50 µm in diameter. The operculatespores of MSX measure 4-6 x 5-8 µm(Fig.M.6.4.2.2b). In C. gigas spores may alsooccur in other tissues. Older foci of infection inboth oyster species may be surrounded byhaemocytes and necrotic tissue debris. A similarinfectious agent occurs in the pearl oysterPinctada maxima in north Western Australia(Fig.M.6.1.3a,b). The spore size of thisHaplosporidium resembles H. nelsoni but differsfrom MSX infections in both C. virginicaand C. gigas by being found exclusively in theconnective tissue.The plasmodial stages of both H. costale and H.nelsoni are as described under M.6.3.1.2.M.6.4.2.3 Transmission Electron Microscopy(TEM) (Level III)TEM is required for confirmation of species-specificultrastructure of the spores - especially inareas enzootic for both disease agents. Tissuesare fixed in 2-3% glutaraldehyde mixed andbuffered for ambient filtered seawater. Oystertissues can also be fixed in 1G4F for 12-24 hrs.Following primary fixation, rinse tissues in asuitable buffer and post-fix in 1-2% osmiumtetroxide (OsO 4= osmic acid - highly toxic).Secondary fixation should be complete within1 hr. The OsO 4fixative must also be rinsed withbuffer/filtered (0.22 µm) seawater prior to dehydrationand resin-embedding.Post-fixed tissues can be stored in a compatiblebuffer or embedded post-rinsing in a resinsuitable for ultramicrotome sectioning. Screeningof 1 micron sections melted onto glassmicroscope slides with 1% toluidine blue solutionis one method of selecting the tissue specimensfor optimum evidence of Haplosporidiumplasmodia or spores.M.6.4.2.4 In situ Hybridization (Level III)DNA-probes for both species of Haplosporidiumhave been produced at the Virginia Institute ofMarine Science (VIMS), College of William andMary, Gloucester, Virginia, USA. These are notyet commercially available, but labelled probesmay be obtained for experienced users, orsamples may be sent to VIMS 1 for in situ hybridizationanalysis.M.6.5 Modes of TransmissionNeither parasite has been successfully transmittedunder laboratory conditions and one (or more)intermediate host(s) is/are suspected.M.6.6 Control MeasuresNone are known for Haplosporidium spp.. Mostefforts have concentrated on development of resistantstocks of oysters. These currently showpotential for survival in enzootic areas, but arenot recommended for use in non-enzootic areasdue to their potential as sub-clinical carriers ofthe pathogen. Some success has also beenachieved in preventing infection of hatcheryrearedlarval and juvenile oysters through filtrationand UV radiation of influent water.1Attention Dr. N. Stokes, Virginia Institute of Marine Science, College of William and Mary, Gloucester Point, Virginia 23062,USA. (E-mail: stokes@vims.edu).141

M.6 Haplosporidiosis(Haplosporidium costale, H. nelsoni)M.6.7 Selected ReferencesAndrews, J.D. 1967. Interaction of two diseasesof oysters in natural waters. Proc. Nat. Shellfish.Assoc. 57: 38-49.Andrews, J.D. 1982. Epizootiology of late summerand fall infections of oysters byHaplosporidium nelsoni, and comparison to theannual life cycle of Haplosporidium costalis, atypical haplosporidian. J. Shellfish Res. 2: 15-23.Andrews, J.D. and M. Castagna. 1978. Epizootiologyof Minchinia costalis in susceptibleoysters in seaside bays of Virginia’s easternshore, 1959-1976. J. Inverteb. Pathol. 32:124-138.Andrews, J.D., J.L. Wood, and H.D. Hoese.1962. Oyster mortality studies in Virginia: III.Epizootiology of a disease caused byHaplosporidium costale, Wood and Andrews.J. Insect Pathol. 4(3): 327-343.Barber, B.J., S.E. Ford, and D.T.J. Littlewood.1991. A physiological comparison of resistantand susceptible oysters Crassostrea virginica(Gmelin) exposed to the endoparasiteHaplosporidium nelsoni (Haskin, Stauber &Mackin). J. Exper. Mar. Biol. Ecol. 146: 101-112.Burreson, E.M. 1997. Molecular evidence for anexotic pathogen: Pacific origin ofHalposporidium nelsoni (MSX), a pathogen ofAtlantic oysters, p. 62. In: M. Pascoe (ed.). 10thInternational Congress of Protozoology, TheUniversity of Sydney, Australia, Monday 21July - Friday 25 July 1997, Programme & Abstracts.Business Meetings & Incentives,Sydney. (abstract).Burreson, E.M., M.E. Robinson, and A. Villalba.1988. A comparison of paraffin histology andhemolymph analysis for the diagnosis ofHaplosporidium nelsoni (MSX) in Crassostreavirginica (Gmelin). J. Shellfish Res. 7: 19-23.Burreson, E.M., N.A. Stokes, and C.S. Friedman.2000. Increased virulence in an introducedpathogen: Haplosporidium nelsoni (MSX) in theeastern oyster Crassostrea virginica. J. Aquat.Anim. Health 12(1): 1-8.Comps, M. and Y. Pichot. 1991. Fine spore structureof a haplosporidan parasitizingCrassostrea gigas: taxonomic implications.Dis. Aquat. Org. 11: 73-77.Farley, C.A. 1967. A proposed life-cycle ofMinchinia nelsoni (Haplosporida,Haplosporidiidae) in the American oysterCrassostrea virginica. J. Protozool. 22(3):418-427.Friedman, C.S., D.F. Cloney, D. Manzer, andR.P. Hedrick. 1991. Haplosporidiosis of thePacific oyster, Crassostrea gigas. J. Inverteb.Pathol. 58: 367-372.Fong, D., M.-Y. Chan, R. Rodriguez, C.-C. Chen,Y. Liang, D.T.J. Littlewood, and S.E. Ford.1993. Small subunit ribosomal RNA gene sequenceof the parasitic protozoanHaplosporidium nelsoni provides a molecularprobe for the oyster MSX disease. Mol.Biochem. Parasitol. 62: 139-142.Ford, S.E. 1985. Effects of salinity on survivalof the MSX parasite Haplosporidium nelsoni(Haskin, Stauber and Mackin) in oysters. J.Shellfish Res. 5(2): 85-90.Ford, S.E. 1992. Avoiding the transmission ofdisease in commercial culture of molluscs, withspecial reference to Perkinsus marinus(Dermo) and Haplosporidium nelsoni (MSX).J. Shellfish Res. 11: 539-546.Ford, S.E. and H.H. Haskin. 1987. Infection andmortality patterns in strains of oystersCrassostrea virginica selected for resistanceto the parasite Haplosporidium nelsoni (MSX).J. Protozool. 73(2): 368-376.Ford, S.E. and H.H. Haskin. 1988. Managementstrategies for MSX (Haplosporidium nelsoni)disease in eastern oysters. Amer. Fish. Soc.Spec. Pub. 18: 249?256.Ford, S.E., Z. Xu, and G. Debrosse. 2001. Useof particle filtration and UV radiation to preventinfection by Haplosporidium nelsoni (MSX) andPerkinsus marinus (Dermo) in hatchery-rearedlarval and juvenile oysters. Aquac. 194(1-2):37-49.Hine, P.M. and T. Thorne. 2000. A survey of someparasites and diseases of several species ofbivalve mollusc in northern Western Australia.Dis. Aquat. Org. 40(1): 67-78.Katkansky, S.C. and R.W. Warner. 1970. Sporulationof a haplosporidian in a Pacific oyster(Crassostrea gigas) in Humboldt Bay, California.J. Fish. Res. Bd. Can. 27(7): 1320-1321.142

M.6 Haplosporidiosis(Haplosporidium costale, H. nelsoni)Kern, F.G. 1976. Sporulation of Minchinia sp.(Haplosporida, Haplosporidiidae) in the Pacificoyster Crassostrea gigas (Thunberg) fromthe Republic of Korea. J. Protozool. 23(4):498-500.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Renault, T., N.A. Stokes, B. Chollet, N.Cochennec, F.C. Berthe, A. Gerard, A. andE.M. Burreson. 2000. Haplosporidiosis in thePacific oyster Crassostrea gigas from theFrench Atlantic coast. Dis. Aquat. Org. 42(3):207-214.Stokes, N.A. and E.M. Burreson. 1995. A sensitiveand specific DNA probe for the oysterpathogen Haplosporidium nelsoni. J. EukaryoticMicrobiol. 42: 350-357.Wolf, P.H. and V. Spague. 1978. An unidentifiedprotistan parasite of the pearl oyster Pinctadamaxima, in tropical Australia. J. Inverteb.Pathol. 31: 262-263.143

M.7 MARTEILIOIDOSIS(MARTEILIOIDES CHUNGMUENSIS,M. BRANCHIALIS)M.7.1 Background InformationM.7.1.1 Causative AgentsMarteilioidosis is caused by two species of parasites,belonging to the protistan PhylumParamyxea. Marteilioides chungmuenis is responsiblefor oocyte infections of Pacific oysters(Crassostrea gigas) and Marteilioides branchialisinfects the gills of Saccostrea glomerata (syn.Crassostrea commercialis, Saccostreacommercialis).The techniques used are the same as describedfor confirmatory disease diagnosis (M.7.4.2.1).Presence of histological inclusions, as describedunder M.7.4.2.1, can be considered confirmatoryfor Marteilioides spp., during screening.(MS Park and DL Choi)M.7.1.2 Host RangeThe Pacific oyster Crassostrea gigas is infectedby Marteilioides chungmuensis. Marteilioidesbranchialis infects the Sydney rock oyster,Saccostrea commercialis.M.7.1.3 Geographic DistributionMarteilioides chungmuensis infects C. gigas inJapan and Korea. Marteilioides branchialis isfound in Australia (New South Wales).M.7.2 Clinical AspectsMarteilioides chungmuensis infects the cytoplasmof mature oocytes and significant proportions ofthe reproductive output of a female oyster canbe affected. The infected eggs are released orretained within the follicle, leading to visible distentionof the mantle surface (Fig.M.7.2a, b).Prevalences of up to 8.3% have been reportedfrom Korea. Marteilioides branchialis causes focallesions on the gill lamellae and, in conjunctionwith infections by Marteilia sydneyi (M.3), isassociated with significant mortalities of Sydneyrock oysters being cultured in trays during theautumn.M.7.3 Screening MethodsM.7.3.1 PresumptiveM.7.3.1.1 Gross Observations (Level I)Marteilioides branchialis causes focal patches (1-2 mm in diameter) of discolouration and swellingon the gill lamellae. The presence of such lesionsin Sydney rock oysters in the Austral autumnshould be treated as presumptive Marteilioidosis.M.7.3.2 ConfirmatoryM.7.3.2.1 Histopathology (Level II)Fig.M.7.2a,b. a. Gross deformation of mantletissues of Pacific oyster (Crassostrea gigas)from Korea, due to infection by the protistanparasite Marteiloides chungmuensis causingretention of the infected ova within the ovaryand gonoducts; b. (insert) normal mantle tissuesof a Pacific oyster.(MS Park)Fig.M.7.4.2.1. Histological section through theovary of a Pacific oyster (Crassostrea gigas) withnormal ova (white arrows) and ova severely infectedby the protistan parasite Marteiliodeschungmuensis (black arrows). Scale bar 100µm.M.7.4 Diagnostic MethodsM.7.4.1 PresumptiveM.7.4.1.1 Gross Observations (Level I)As for M.7.3.1.1, focal patches (1-2 mm in diameter)of discolouration and swelling on the gilllamellae of Sydney rock oysters in the Australautumn can be treated as presumptive positivesfor M. branchialis.144

M.7 Marteilioidosis(Marteilioides chungmuensis,M. branchialis)M.7.4.1.2 Histopathology (Level II)For first-time diagnoses a back up tissue specimenfixed for EM is recommended (M.7.4.2.3).M.7.4.2 ConfirmatoryM.7.4.2.1 Histopathology (Level II)Positive histological sections can be consideredconfirmatory where collected from susceptibleoyster species and areas with a historic recordof the presence of Marteilioides spp. infections.It is recommended that at least two dorso-ventralsections through each oyster be examinedusing oil immersion for screening purposes. Sectionsfrom oysters >2 yrs (or >30 mm shell height)should be fixed immediately in a fast fixative suchas 1G4F. Davidsons or 10% buffered formalinmay be used for smaller or whole oysters (seeM.1.3.3.3) but these fixatives are not optimal forsubsequent confirmatory Electron Microscopy(EM) diagnosis (M.6.4.2.3), or species identification,if required. Several standard stains (e.g.,haemotoxylin-eosin) enable detection ofMarteilioides spp..Marteilioides chungmuensis is located in the cytoplasmof infected ova (Fig.M.7.4.2.1). Stem(primary) cells contain secondary cells. Thesemay, in turn, contain developing sporonts, givingrise to a single tertiary cell by endogenous budding.Each tertiary cell forms a tricellular sporeby internal cleavage.Marteilioides branchialis causes epithelial hyperplasiaand granulocyte infiltration at the site ofinfection. Uninucleate primary cells contain twoto six secondary cells (some may contain up to12) in the cytoplasm of epithelial cells, connectivetissue cells and occasionally the infiltratinghaemocytes within the lesion.M.7.4.2.2 Transmission Electron Microscopy(TEM) (Level III)TEM is required for confirmation of species-specificultrastructure of these parasites. Tissues arefixed in 2-3% glutaraldehyde mixed and bufferedfor ambient filtered seawater. Tissues can alsobe fixed in 1G4F for 12-24 hours. Following primaryfixation, rinse in a suitable buffer and postfixin 1-2% osmium tetroxide (OsO 4= osmic acid- highly toxic). Secondary fixation should be completewithin 1 hour. The OsO 4fixative must alsobe rinsed with buffer/filtered (0.22 microns) seawaterprior to dehydration and resin-embedding.Post-fixed tissues should be stored in a compatiblebuffer or embedded post-rinsing in aresin suitable for ultramicrotome sectioning.Screening of 1 micron sections melted ontoglass microscope slides with 1% toludine bluesolution is one method of selecting the tissuespecimens for optimum evidence of putativeMarteilioides spp. Ultrathin sections are thenmounted on copper grids (with or withoutformvar reinforced support) for staining withlead citrate + uranyl acetate or equivalent EMstain.Marteilioides branchialis is differentiated from theother Marteilioides spp. by the presence of twoconcentric cells (rather than three) within thespore. In addition M. chungmuensis in C. gigascontains only two to three sporonts per primary/stem cell compared with two-six (or up to 12) forM. branchialis. Multivesicular bodies resemblingthose of Marteilia spp. are present in M.branchialis stem cells, but absent from those ofM. chungmuensis.M.7.5 Modes of TransmissionUnknown.M.7.6 Control MeasuresNone known.M.7.7 Selected ReferencesAnderson, T.J. and R.J.G. Lester. 1992.Sporulation of Marteilioides branchialis n.sp.(Paramyxea) in the Sydney rock oyster,Saccostrea commercialis: an electronmicroscope study. J. Protozool. 39(4): 502-508.Anderson, T.J., T.F. McCaul, V. Boulo, J.A.F.Robledo, and R.J.G. Lester. 1994. Light andelectron immunohistochemical assays onparamyxea parasites. Aquat. Living Res. 7(1):47-52.Elston, R.A. 1993. Infectious diseases of thePacific oyster Crassostrea gigas. Ann. Rev.Fish Dis. 3: 259-276.Comps, M., M.S. Park, and I. Desportes. 1986.Etude ultrastructurale de Marteilioideschungmuensis n.g. n.sp., parasite desovocytes de l’hu?tre Crassostrea gigas Th.Protistol. 22(3): 279-285.145

M.7 Marteilioidosis(Marteilioides chungmuensis,M. branchialis)Comps, M., M.S. Park, and I. Desportes. 1987.Fine structure of Marteilioides chungmuensisn.g. n.sp., parasite of the oocytes of theoyster Crassostrea gigas. Aquac. 67(1-2):264-265.Hine, P.M. and T. Thorne. 2000. A survey ofsome parasites and diseases of severalspecies of bivalve mollusc in northernWestern Australia. Dis. Aquat. Org. 40(1): 67-8.146

M.8 IRIDOVIROSIS(OYSTER VELAR VIRUS DISEASE)M.8.1Background InformationM.8.3.2 ConfirmatoryM.8.1.1 Causative AgentsOyster Velar Virus Disease (OVVD) (Iridovirosis)is caused by an icosahedral DNA virus with morphologicalsimilarities to the Iridoviridae.M.8.1.2 Host RangeCrassostrea gigas (Pacific oyster) larvae are thedocumented host species, although similar viralagents have been associated with gill disease(“Maladie des Branchies”) and haemocyte infectionsin Portugese oysters (Crassostrea angulata)and C. gigas.M.8.1.3 Geographic DistributionInfections have been reported solely from twohatcheries in Washington State, but are believedto have a ubiquitious distribution throughout juvenileC. gigas production, with clinical manifestationonly under sub-optimal growing conditions.M.8.2 Clinical AspectsOVVD causes sloughing of the velar epitheliumof larvae >150µm in length, and can cause upto 100% mortality under hatchery conditions.The larvae can not feed, weaken and die.M.8.3 Screening MethodsM.8.3.1 PresumptiveGenerally-speaking, since this is an opportunisticinfection, only clinical infections will demonstratedetectable infections – as described underM.8.4.M.8.3.1.1 Wet Mounts (Level I)Wet mounts of veliger larvae which demonstratesloughing of ciliated epithelial surfaces can beconsidered to be suspect for OVVD. As withgross observations, other opportunistic pathogens(bacteria and Herpes-like viruses) may beinvolved, so Level II/III diagnostics are required.M.8.3.1.2 Histopathology (Level II)Using the techniques described under M.8.4.2.1,detection the features described in that sectioncan be considered to be presumptively positivefor OVVD. Such inclusions require TEM (LevelIII) (M.8.4.2.2) for confirmatory diagnosis, at leastfor first time observations.M.8.3.2.1 Transmission Electron Microscopy(Level III)As described under M.8.4.2.2.M.8.4 Diagnostic MethodsM.8.4.1 PresumptiveM.8.4.1.1 Gross Observations (Level I)Slowed growth, cessation of feeding and swimmingin larval Crassostrea gigas should be consideredsuspect for OVVD. Gross signs are notpathogen specific and require Level II examination(M.8.4.2), at least for first time observations.M.8.4.1.2 Wet Mounts (Level I)As described under M.8.3.1.1. For first-time diagnosesa back up tissue specimen fixed for TEMis recommended (M.8.4.2.2).M.8.4.1.3 Histopathology (Level II)As described under M.8.4.2.1.M.8.4.1.4 Transmission Electron Microscopy(Level III)As described under M.8.4.2.2.M.8.4.2 ConfirmatoryM.8.4.2.1 Histopathology (Level II)Where larvae have a history of OVVD, detectionof inclusions and ciliated epithelial pathology, asdescribed below, can be considered confirmatoryfor the disease. However, it should be notedthat other microbial infections can induce similarhistopathology and electron microscopy is theideal confirmatory technique (M.8.4.2.2).Larvae must be concentrated by centrifugationor filtration into a pellet prior to embedding. Thisis best achieved post-fixation in Davidson’s,1G4F or other fixative. Although paraffin embeddingis possible, resin embedding is recommendedfor optimal sectioning. Paraffin permitssectioning down to 3 µm using standard microtome.Resin embedded tissue can be sectioneddown to 1 µm thick, but requiresspecialised microtomes and/or block holdersand specialised staining.147

M.8 Iridovirosis(Oyster Velar Virus Disease)Standard stains (e.g., haemotoxylin-eosin) willdetect intracytoplasmic inclusion bodies in ciliatedvelar epithelial cells. Early inclusion bodiesare spherical, but become more irregular as theviruses proliferate. Inclusion bodies may also bedetected in oesophageal and oral epithelia or,more rarely, in mantle epithelial cells.M.8.4.2.2 Transmission Electron Microscopy(TEM) (Level III)TEM is required to visualise the causative virusesin situ in gill tissue sections of concentrated ‘pellets’of larvae. Fixation in 2-3% glutaraldehydemixed and buffered for ambient filtered seawatershould not exceed 1 hour to reduce artifacts. Tissuescan also be fixed in 1G4F for 12-24 hrs.Following primary fixation, rinse in a suitablebuffer and post-fix in 1-2% osmium tetroxide(OsO 4= osmic acid - highly toxic). Secondaryfixation should be complete within 1 hr. The OsO 4fixative must also be rinsed with buffer/filtered(0.22 µm) seawater prior to dehydration andresin-embedding.Post-fixed tissues should be stored in a compatiblebuffer or embedded post-rinsing in a resinsuitable for ultramicrotome sectioning. Screeningof 1 micron sections melted onto glass microscopeslides with 1% toludine blue solution isone method of selecting the best specimens forultrathin sectioning. Ultrathin sections aremounted on copper grids (with or without formvarreinforced support) for staining with lead citrate+ uranyl acetate or equivalent EM stain.Icosahedral viral particles (228 +/- 7 nm in diameter)with a bi-laminar membrane capsid shouldbe evident to confirm Iridoviral involvement.M.8.5 Modes of TransmissionThe disease appeared in March-May at affectedhatcheries. Direct transmission between moribundand uninfected larvae is suspected.M.8.6 Control MeasuresNone known except for reduced stocking densities,improved water exchange and generalhatchery sanitation methods (tank and line disinfection,etc.).M.8.7 Selected ReferencesComps, M. and N. Cochennec. 1993. A Herpeslikevirus from the European oyster Ostreaedulis L. J. Inverteb. Pathol. 62: 201-203.Elston, R.A. 1979. Virus-like particles associatedwith lesions in larval Pacific oysters(Crassostrea gigas). J. Inverteb. Pathol. 33:71-74.Elston, R.A. 1993. Infectious diseases of thePacific oyster, Crassostrea gigas. Ann. Rev.Fish Dis. 3: 259-276.Elston, R. 1997. Special topic review: bivalvemollusc viruses. Wor. J. Microbiol. Biotech. 13:393-403.Elston, R.A. and M.T. Wilkinson. 1985. Pathology,management and diagnosis of oyster velarvirus disease (OVVD). Aquac. 48: 189-210.Farley, C.A. 1976. Epizootic neoplasia in bivalvemolluscs. Prog. Exper. Tumor Res. 20: 283-294.Farley, C.A., W.G. Banfield, G. Kasnic Jr. andW.S. Foster. 1972. Oyster Herpes-type virus.Science 178: 759-760.Hine, P.M., B. Wesney and B.E. Hay. 1992. Herpesvirus associated with mortalities amonghatchery-reared larval Pacific oystersCrassostrea gigas. Dis. Aquat. Org. 12: 135-142.Le Deuff, R.M., J.L. Nicolas, T. Renault and N.Cochennec. 1994. Experimental transmissionof a Herpes-like virus to axenic larvae of Pacificoyster, Crassostrea gigas. Bull. Eur.Assoc. Fish Pathol.14: 69-72.LeDeuff, R.M., T. Renault and A. G?rard. 1996.Effects of temperature on herpes-like virusdetection among hatchery-reared larval Pacificoyster Crassostrea gigas. Dis. Aquat. Org. 24:149-157.Meyers, T.R. 1981. Endemic diseases of culturedshellfish of Long Island, New York: adult andjuvenile American oysters (Crassostreavirginica) and hard clams(Mercenariamercenaria). Aquac. 22: 305-330.Nicolas, J.L., M. Comps and N. Cochennec.1992. Herpes-like virus infecting Pacific oysterlarvae, Crassostrea gigas. Bull. Eur. Assoc.Fish Pathol. 12: 11-13.Renault, T., N. Cochennec, R.M. Le Deuff andB. Chollet. 1994. Herpes-like virus infectingJapanese oyster (Crassostrea gigas) spat.Bull. Eur.Assoc. Fish Pathol. 14: 64 -66.148

Annex M.AI. OIE Reference Laboratoryfor Molluscan DiseasesDiseaseMollusc pathogensExpert/LaboratoryDr. F. BerthIFREMERLaboratoire de Genetique Aquaculture et PathologieBP 133, 17390 La TrembladeFRANCETel: 33(0)5 46.36.98.36Fax: 33 (0)5 46.36.37.51E-mail: fberthe@ifremer.fr149

Annex M.AII. List of Regional Resource Expertsfor Molluscan Diseases Asia-Pacific 1DiseaseBonamiosisMarteiliosisMolluscan DiseasesExpertDr. Brian JonesSenior Fish PathologistFisheries WAAdjunct Professor, Muresk Institutec/o Animal Health Labs3 Baron-Hay CourtSouth Perth WA 6151AUSTRALIATel: 61-8-9368-3649Fax: 61-8-9474-1881E-mail: bjones@agric.wa.gov.auMarteilioisis/MicrocytosisDr. Robert D. AdlardQueensland MuseumPO Box 3300, South Brisbane,Queensland 4101AUSTRALIATel: +61 7 38407723Fax: +61 7 38461226E-mail: RobertAd@qm.qld.gov.au;http://www.qmuseum.qld.gov.auDr. Sarah N. KleemanAquatic Animal BiosecurityAnimal BiosecurityGPO Box 858Canberra ACT 2601AUSTRALIATel: +61 2 6272 3024Fax: +61 2 6272 3399E-mail: sarah.kleeman@affa.gov.auProfessor R.J.G. LesterDepartment of Microbiology and ParasitologyThe University of Queensland, BrisbaneAUSTRALIA 4072.Tel: +61-7-3365-3305,Fax: +61-7-3365-4620E-mail: R.Lester@mailbox.uq.edu.auhttp://www.biosci.uq.edu.au/micro/academic/lesterlester.htmDr. Dong Lim ChoiPathology Division#408-1, Shirang-ri, Kijang-upKijang-gun, Busan 619-902KOREA ROTel: +82-51-720-2493Fax: +82-51-720-2498E-mail: dlchoi@haema.nfrda.re.krDr. Mi-Seon ParkPathology Division#408-1, Shirang-ri, Kijang-upKijang-gun, Busan 619-902KOREA RO 619-902Tel: +82-51-720-2493Fax: +82-51-720-2498E-mail: parkms@nfrdi.re.kr1The experts included in this list has previously been consulted and agreed to provide valuable information and health adviseconcerning their particular expertise.150

Annex M.AII. List of Regional Resource Experts forMolluscan Diseases Asia-PacificList of Experts Outside Asia-Pacific Region 2Dr. Jie HuangYellow Sea Fisheries Research InstituteChinese Academy of Fishery Sciences106 Nanjing RoadQingdao, Shandong 266071PEOPLE’S REPUBLIC of CHINATel: 86 (532) 582 3062Fax: 86 (532) 581 1514E-mail: aqudis@public.qd.sd.cnMr. Arthur de VeraFish Health SectionBureau of Fisheries and Aquatic ResourcesArcadia Building, 860 Quezon AvenueQuezon City, Metro ManilaPHILIPPINESFax: (632) 3725055Tel: (632) 4109988 to 89E-mail: azdv@edsamail.com.phDr. Paul Michael HineAquatic Animal DiseasesNational Centre for Disease InvestigationMAF Operations, P.O. Box 40-742Upper HuttNEW ZEALANDTel: +64-4-526-5600Fax: +64-4-526-5601E-mail: hinem@maf.govt.nzMolluscan DiseasesDr. Susan BowerDFO Pacific Biological Station3190 Hammond Bay RoadNanaimo, British ColumbiaV9R 5K6CANADATel: 250-756-7077Fax: 250-756-7053E-mail: bowers@dfo-mpo.gc.caDr. Sharon E. McGladderyOceans and Aquaculture Science200 Kent Street (8W160)Ottawa, Ontario, K1A 0E6CANADATel: 613-991-6855Fax: 613-954-0807E-mail: mcgladderys@dfo-mpo.gc.ca2These experts outside the Asia-Pacific region has supported the regional programme on aquatic animal health and agreed toassist further in providing valuable information and health advise on molluscan diseases.151

Annex M.AIII. List of Useful Diagnostic Manuals/Guides/Keys to Molluscan Diseases• Australian Aquatic Animal Disease – Identification Field Guide (1999) by Alistair Herfortand Grant RawlinInformation: AFFA Shopfront – Agriculture, Fisheries and Forestry – AustraliaGPO Box 858, Canberra, ACT 2601Tel: (02) 6272 5550 or free call: 1800 020 157Fax: (02) 6272 5771E-mail: shopfront@affa.gov.au• Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish byBower, SE McGladdery and IM Price (1994)Information: Dr. Susan BowerDFO Pacific Biological Station3190 Hammond Bay RoadNanaimo, British ColumbiaV9R 5K6CANADATel: 250-756-7077Fax: 250-756-7053E-mail: bowers@dfo-mpo.gc.ca• Mollusc Diseases: Guide for the Shellfish Farmer. 1990. by R.A. Elston. Washington SeaGrant Program, University of Washington Press, Seattle. 73 pp.• A Manual of the Parasites, Pests and Diseases of Canadian Atlantic Bivalves. 1993. bySE McGladdery, RE Drinnan and MF Stephenson.Information: Dr. Sharon McGladderyOceans and Aquaculture Science200 Kent Street (8W160)Ottawa, Ontario, K1A 0E6Tel: 613-991-6855Fax: 613-954-0807E-mail: mcgladderys@dfo-mpo.gc.ca152

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Internal and External Anatomy ofa Penaeid Shrimpstomacheye stalkoesophagusantenna pereiopodshepatopancreasheartpleopodsInternal and external anatomy of a penaeid shrimp.hindgutabdominalsegmentanus154

SECTION 4 -CRUSTACEAN DISEASESInternal and External Anatomy of a Penaeid ShrimpSECTION 4 - CRUSTACEAN DISEASESC.1 GENERAL TECHNIQUESC.1.1 Gross ObservationsC.1.1.1 BehaviourC.1.1.1.1 GeneralC.1.1.1.2 MortalitiesC.1.1.1.3 FeedingC.1.1.2 Surface ObservationsC.1.1.2.1 Colonisation and ErosionC.1.1.2.2 Cuticle Softening, Spots and DamageC.1.1.2.3 ColourC.1.1.2.4 Environmental ObservationsC.1.1.3 Soft-Tissue SurfacesC.1.2 Environmental ParametersC.1.3 General ProceduresC.1.3.1 Pre-collection PreparationC.1.3.2 Background InformationC.1.3.3 Sample Collection for Health SurveillanceC.1.3.4 Sample Collection for Disease DiagnosisC.1.3.5 Live Specimen Collection for ShippingC.1.3.6 Preservation of Tissue SamplesC.1.3.7 Shipping Preserved SamplesC.1.4 Record-KeepingC.1.4.1 Gross ObservationsC.1.4.2 Environmental ObservationsC.1.4.3 Stocking RecordsC.1.5 ReferencesVIRAL DISEASES OF SHRIMPC.2 Yellowhead Disease (YHD)C.3 Infectious Hepatopancreas and HaematopoieticNecrosis (IHHN)C.4 White Spot Disease (WSD)C.4a Bacterial White Spot Syndrome (BWSS)C.5 Baculoviral Midgut Gland Necrosis (BMN)C.6 Gill-Associated Virus (GAV)C.7 Spawner Mortality Syndrome("Midcrop mortality syndrome")C.8 Taura Syndrome (TS)C.9 Nuclear Polyhedrosis Baculovirosis (NPD)BACTERIAL DISEASE OF SHRIMPC.10 Necrotising Hepatopancreatitis (NH)FUNGAL DISEASE OF CRAYFISHC.11 Crayfish Plague154157157157157157158158158158158160160160160160162162162162164165165165165166166167173178183186189192194201207211C.AIANNEXESOIE Reference Laboratories forCrustacean Diseases215155

Section 4 - Crustacean DiseasesC.AIIC.AIIIList of Regional Resource Experts for CrustaceanDiseases in the Asia-PacificList of Useful Manuals/Guide to CrustaceanDiseases in Asia-Pacific216219List of National Coordinators(NCs)Members of Regional Working Group (RWG) andTechnical Support Services (TSS)List of Figures221225230156

C.1 GENERAL TECHNIQUESGeneral crustacean health advice and othervaluable information are available from the OIEReference Laboratories, Regional ResourceExperts in the Asia-Pacific, FAO and NACA.A list is provided in Annexes F.AI and AII, andup-to-date contact information may be obtainedfrom the NACA Secretariat in Bangkok(E-mail:naca@enaca.org) . Other useful guidesto diagnostic procedures which provide valuablereferences for crustacean diseases arelisted in Annex F.AIII.C.1.1 Gross ObservationsGross observations of clinical signs in shrimpcan be easily made at the farm or pond sideusing little, if any, equipment. Although, in mostcases, such observations are insufficient for adefinite diagnosis, such information is essentialfor preliminary compilation of a strong “casedescription” (or case history). Accurate anddetailed gross observations also help with initiationof an action plan which can effectivelyreduce losses or spread of the disease, e.g.,destruction or isolation of affected stocks, treatmentsor alterations to husbandry practices (i.e.,feeding regimes, stocking densities, pondfertilisation, etc.). These can all be started whilewaiting for more conclusive diagnostic results.C.1.1.1 Behaviour (Level 1)C.1.1.1.1 GeneralAbnormal shrimp behaviour is often the first signof a stress or disease problem. Farmers or farmworkers, through daily contact with their stocks,rapidly develop a subconscious sense of when“something is wrong”. This may be subtlechanges in feeding behaviour, swimming movementor unusual aggregations. Even predatoractivity can provide clues to more “hidden”changes such as when fish- or shrimp-eatingbirds congregate round affected ponds.Record-keeping (see C.1.4) can provide valuableadditional evidence that reinforces suchobservations and can indicate earlier dateswhen problems started to appear. It is importantthat farmers and workers on the farm, aswell as field support staff, get to know the “normal”(healthy) behaviour of their stocks. Sincesome species and growing environments maydemonstrate or evoke subtle differences inbehaviour, these should be taken into account,especially if changing or adding species, orwhen information gathered from a differentgrowing environment is used. Where anychange from normal behaviour affects morethan small numbers of random individuals, thisshould be considered cause for concern andwarrants investigation.Some clues to look out for in shrimp stocks include:• unusual activity during the daytime - shrimpstend to be more active at night and stick todeeper water during the day• swimming at or near pond surface or edges- often associated with lethargy (shrimpswimming near the surface may attractpredatory birds)• increased feed consumption followed bygoing off-feed• reduction or cessation of feeding• abnormal feed conversion ratios, length/weight ratios• general weakening - lethargy (note: lethargyis also characteristic in crustaceans when thewater temperature or dissolved oxygen levelsare low, so these possibilities should beeliminated as potential causes before diseaseinvestigations are started)C.1.1.1.2 MortalitiesMortalities that reach levels of concern to a producershould be examined for any patterns inlosses, such as:• relatively uniform mortalities throughout asystem should be examined immediately andenvironmental factors determined (ideallywith pre-mortality records - see C.1.4)• apparently random, or sporadic mortalitiesmay indicate a within-system or stock problems.If the following conditions exist - (a)no history of stock-related mortalities, (b) allstock originate from the same source, and(c) there have been no changes to the rearingsystem prior to mortality problems -samples of affected and unaffected shrimpshould be submitted for laboratory examination(Level II or III), as appropriate, andsupported by gross observations and stockhistory (see C.1.4)• mortalities that spread suggest an infectiouscause and should be sampled immediately.Affected shrimp should be kept as faraway as possible from unaffected shrimp untilthe cause of the mortalities can be established.157

C.1 General TechniquesC.1.1.1.3 FeedingAbsence of feeding behaviour and lack of feedin the gut are good indicators of potential problems.Daily gut content checks can be madeon shrimp caught in feeding trays or bowls(where used) or, less frequently, from samplestaken to determine growth. Ideally examinationof feeding behaviour should be made every 1-2 weeks, even in extensive farming systems.Feeding behaviour is most easily checked byplacing feed in a tray or bowl (Fig.C.1.1.1.3a)and seeing how quickly the shrimp respond,ideally after the shrimp have not been fed for atleast a few hours. It is important that the feedused is attractive to the shrimp as poorly formulated,old or badly stored feeds may not beattractive to the shrimp. Gut contents can bechecked by holding the shrimp against a lightto show the gut in the tail segments(Fig.C.1.1.1.3b). If these are empty, especiallyjust after providing feed, it may indicate eitherof the following conditions: i) underfeeding, orii) onset of cessation of feeding (anorexia).Where possible, feed records (see C.1.4) shouldbe maintained to determine normal feed consumptionpatterns (i.e., feeding activity byhealthy shrimp), which can be compared with“suspect” feeding activity. In many cases ofchronic loss, daily feed consumption patternsmay remain stable or oscillate over periods ofseveral weeks. These can be detected by makinga graph of daily feed consumption or bycomparing daily feed consumption in the recordbook over an extended period (e.g. 3-4 weeks).C.1.1.2 Surface Observations (Level 1)C.1.1.2.1 Colonisation and ErosionColonisation of the shell (cuticle) and gills of acrustacean is an on-going process that is usuallycontrolled by grooming. The presence ofnumerous surface organisms (e.g. “parasites”- which damage their host; or “commensals” -that do not adversely impact their host) suggestssub-optimal holding conditions or a possibledisease problem. Apparent wearing away(erosion) of the cuticle or appendages (legs, tail,antennae, rostrum) (Fig.C.1.1.2.1a), or loss ofappendages, with or without blackening (melanization)are also highly indicative of a diseaseproblem. Breakage of the antennae is an earlywarning sign. In healthy penaeid shrimp, theseshould extend approximately 1/3 past thelength of the body (when bent back along thebody line). Likewise, erosion or swelling of thetail (uropods and telson), with or without blackening,is an early sign of disease(Fig.C.1.1.2.1b).C.1.1.2.2 Cuticle Softening, Spots andDamageSoftening of the shell (Fig.C.1.1.2.2a andFig.C.1.1.2.2b), other than during a moult, mayalso indicate the presence of infection. Damageor wounds to the shell provide an opportunityfor opportunistic infections (mainly bacterialand fungal) to invade the soft-tissues andproliferate, which can seriously impact thehealth of the shrimp.Certain diseases, such as White Spot Disease,directly affect the appearance of the shell, however,few changes are specific to a particularinfection. In the case of white spots on the cuticle,for example, recent work (Wang et al. 2000)has shown that a bacteria can produce signssimilar to those produced by White Spot Disease(see C.4) and Bacterial White Spot Syndrome(see C.4a).C.1.1.2.3 ColourShrimp colour is another good indicator ofhealth problems. Many crustaceans becomemore reddish in color when infected by a widerange of organisms, or when exposed to toxicconditions (Fig.C.1.1.2.3a), especially those thataffect the hepatopancreas. This is thought tobe due to the release of yellow-orange (carotenoid)pigments that are normally stored in thehepatopancreas. This red colour is not specificfor any single condition (or groups of infections),however, so further diagnosis is needed.Yellowish coloration of the cephalothorax isassociated with yellowhead disease (see C.2)and overall reddening can be associated withgill associated virus infections (see C.6), whitespot disease or bacteria, as described above,or bacterial septicemia (see C.10). In somecases, the colour changes are restricted to extremities,such as the tail fan or appendages(Fig.C.1.1.2.3b), and these should be examinedclosely.It should be noted that some shrimpbroodstock, particularly those from deeperwaters, can be red in colour (thought to be dueto a carotenoid-rich diet). This does not appearto be related to health and its normality can beestablished through familiarisation with the speciesbeing grown. Under certain conditions,some crustaceans may turn a distinct blue158

C.1 General Techniques(P Chanratchakool)(P Chanratchakool/MG Bondad-Reantaso)aFig.C.1.1.1.3a. Behaviour observation ofshrimp PL in a bowl.(P Chanratchakool)bFig.C.1.1.2.2a,b. Shrimp with persistent softshell.Fig.C.1.1.1.3b. Light coloured shrimp with fullguts from a pond with healthy phytoplankton.(P Chanratchakool)(P Chanratchakool)Fig.C.1.1.2.3a. Abnormal blue and red discoloration.(P Chanratchakool)Fig.C.1.1.2.1a. Black discoloration of damagedappendages.(P Chanratchakool)Fig.C.1.1.2.3b.Red discolorationof swollen appendage.>Fig.C.1.1.2.1b. Swollen tail due to localizedbacterial infection.159

C.1 General Techniquescolour. This has been shown to be due to lowlevels of a carotenoid pigment in the hepatopancreas(and other tissues), which may be inducedby environmental or toxic conditions.Normal differences in colouration (light to dark)within a species may be due to other environmentalvariables. For example, Penaeusmonodon grown in low salinities, are often muchpaler than P. monodon grown in brackish-wateror marine conditions. These variations donot appear to be related to general health.C.1.1.2.4 Environmental ObservationsShrimp with brown gills or soft shells (or a representativesub-sample), should be transferredto a well aerated aquarium with clean sea waterat the same salinity as the pond from whichthey came. They should be observed every 1-2hrs over 1 day. If the shrimp return to normalactivity within a few hours, check environmentalparameters in the rearing pond(s).C.1.1.3 Soft-Tissue Surfaces (Level 1)A readily observable change to soft tissues isfouling of the gill area (Fig. C.1.1.3a), sometimesaccompanied by brown discoloration (Fig.C.1.1.3b) (see C.1.1.2.4). This can be due todisease and should trigger action since it reducesthe shrimp’s ability to take up oxygenand survive.Removal of the shell in the head region ofshrimp allows gross examination of the organsin this region, particularly the hepatopancreas(Fig.C.1.1.3c). In some conditions, the hepatopancreasmay appear discoloured (i.e., yellowish,pale, red), swollen or shrunken, comparedwith healthy shrimp. If the hepatopancreas isgently teased out of the shell, the mid-gut willbecome visible and permit direct examinationof colour (dark - feeding; light/white/yellow -mucoid, empty or not feeding - see C.1.1.1.3).This information is useful for determining thehealth of the shrimp and if infectious diseaseagents are present.C.1.2 Environmental Parameters(Level 1)Environmental conditions can have a significanteffect on crustacean health, both directly (withinthe ranges of physiological tolerances) and indirectly(enhancing susceptibility to infectionsor their expression). Examples include changesto dissolved oxygen levels and/or pH which maypromote clinical expression of previously latentyellowhead disease (see C.2) and white spotdisease (see C.4) or the effect of salinity on theexpression of necrotising hepatopancreatitis(see C.10). This is especially important for speciesgrown under conditions that bear littleresemblance to the wild situation. Water temperature,salinity, turbidity, fouling and planktonblooms (Fig. C.1.2 a,b,c and d) are all importantfactors. Rapid changes in conditions,rather than gradual changes, are particularly importantas potential triggers for disease. Therefore,the farm manager and workers, should attemptto keep pond rearing conditions withinthe optimum range for the species and as constantas possible within that range. High stockingrates are common in aquaculture but predisposeindividuals to stress so that even minorchanges in environmental conditions mayprecipitate disease. In addition, many smallchanges do not, on their own, affect shrimphealth. However, when several of these smallchanges occur simultaneously, results can befar more severe.C.1.3 General ProceduresC.1.3.1 Pre-collection Preparation(Level I)The diagnostic laboratory which will be receivingthe sample should be consulted to ascertainthe best method of transportation (e.g., onice, preserved in fixative, whole or tissuesamples). The laboratory will also indicate ifboth clinically affected, as well as apparentlyhealthy individuals, are required for comparativepurposes. As noted under C.1.3.3 andC.1.3.4, screening and disease diagnosis oftenhave different sample-size requirements.The laboratory should be informed of exactlywhat is going to be sent (i.e., numbers, sizeclassesor tissues) and the intended date of collectionand delivery, as far in advance as possible.For screening healthy animals, samplesizes are usually larger so more lead time is requiredby the laboratory. Screening can be alsobe planned ahead of time, based on predicteddates of shipping post-larvae (PL) orbroodstock, which means the shipper has moretime to notify the laboratory well in advance. Incases of disease outbreaks and significantmortalities, there may be less opportunity foradvance warning for the laboratory. However,the laboratory should still be contacted prior toshipment or hand-delivery of any diseasedsamples (for the reasons given under C.1.3.4).Some samples may require secured packagingor collection by designated personnel, ifthere are national or international certification160

C.1 General Techniques(P Chanratchakool)(P Chanratchakool)Fig.C.1.1.3a. Severe fouling on the gills.(P Chanratchakool)aFig.C.1.1.3b. Brown discolouration of the gills.b(P Chanratchakool)cFig.C.1.1.3c. Shrimp on left side with smallhepatopancreas.(V Alday de Graindorge and TW Flegel)Fig.C.1.2a, b, c. Examples of different kinds ofplankton blooms (a- yellow/green colouredbloom; b- brown coloured bloom; c- blue-greencoloured bloom.(P Chanratchakool)Fig.C.1.2d. Dead phytoplankton.>Fig. C.1.3.6. Points of injection of fixative.161

C.1 General Techniquesrequirements or risk of disease spread via transportof the sample to an area non-endemic fora suspected disease.Pre-collection discussions with the diagnosticlaboratory can significantly speed up processingand diagnosis of a sample (days to weeks)since it allows preparation of the required diagnosticmaterials in advance of arrival of thesample(s) and ensures that emergency samplesare scheduled in for rapid diagnosis.C.1.3.2 Background Information (Level 1)All samples submitted for diagnosis should includeas much supporting information as possibleincluding:• Gross observations and a history of environmentalparameters (as described under C.1.1and C.1.2)• Approximate prevalence and pattern of mortality(acute or chronic/sporadic cumulativelosses)• History and origin of affected population• If the stock is not local, their origin(s) anddate(s) of transfer should be included• Details of feed, consumption rates and anychemical treatments usedThe above information provides valuable backgrounddetails which can help focus attentionon possible handling stress, changes in environmentor infectious agents as the primarycause of any health problems.C.1.3.3 Sample Collection for Health SurveillanceThe most important factors associated withcollection of specimens for surveillance are:• sample numbers that are high enough toensure adequate pathogen detection (seeC.1.3.1 and Table C.1.3.3). Check the numberof specimens required by the laboratorybefore collecting the sample(s) and ensurethat each specimen is intact. Larger numbersare generally needed for screening purposes,compared to numbers required for diseasediagnosis;• susceptible species are sampled;• samples include age- or size-groups that aremost likely to manifest detectable infections.Such information is given under the specificdisease sections; and• samples are collected during the seasonwhen infections are known to occur. Suchinformation is also given under the specificdisease sections.As mentioned under C.1.3.1, check whether ornot designated personnel are required to do thecollection, or if secured packaging is necessary,or whether samples are being collected to meetnational or international certification requirements.C.1.3.4 Sample Collection for DiseaseDiagnosisAll samples submitted for disease diagnosisshould include as much supporting informationas possible, as described under C.1.3.2, withparticular emphasis on:• rates and levels of mortality compared with“normal” levels for the time of year;• patterns of mortality (random/sporadic,localised, spreading, widespread);• history and origin(s) of the affectedpopulation(s); and• details of feed used, consumption rates andany chemical treatments.As in C.1.3.2, the above information will helpclarify whether or not an infectious agent is involvedand will enable to focus the investigativeprocedures required for an accurate diagnosis.This information is also critical for laboratoriesoutside the region or areas where thesuspected disease is endemic. Under such circumstances,the laboratory may have to preparefor strict containment and sterile disposalof all specimen shipping materials and wasteproducts, in order to prevent escape from thelaboratory.Wherever possible, check the number of specimensrequired by the laboratory for diagnosticexamination before collecting the sample(s).Also check with the laboratory to see whetheror not they require specimens showing clinicalsigns of disease only, or sub-samples of bothapparently healthy individuals and clinically affectedspecimens from the same pond/site. Thelatter option is usually used where a diseaseoutbreakor other problem is detected for thefirst time. Comparative samples can help pinpointabnormalities in the diseased specimens.C.1.3.5 Live Specimen Collection forShipping (Level 1)Once the required sample size is determined,the crustaceans should be collected from thewater. This should take place as close to shippingas possible to reduce possible mortalitiesduring transportation (especially important formoribund or diseased samples). Wherever pos-162

C.1 General TechniquesPrevalence (%)Population Size 0.5 1.0 2.0 3.0 4.0 5.0 10.050 46 46 46 37 37 29 20100 93 93 76 61 50 43 23250 192 156 110 75 62 49 25500 314 223 127 88 67 54 261000 448 256 136 92 69 55 272500 512 279 142 95 71 56 275000 562 288 145 96 71 57 2710000 579 292 146 96 72 29 27100000 594 296 147 97 72 57 271000000 596 297 147 97 72 57 27>1000000 600 300 150 100 75 60 30Table C.1.3.3 1 . Sample sizes needed to detect at least one infected host in a population of agiven size, at a given prevalence of infection. Assumptions of 2% and 5% prevalences are mostcommonly used for surveillance of presumed exotic pathogens, with a 95% confidence limit.sible, ensure that each specimen is intact.As noted under C.1.3.1, inform the laboratoryof the estimated time of arrival of the sampleso they can have the materials required to processprepared before the samples arrive. Thisshortens the time between removal from thepond and preparation of the specimens for examination.The crustaceans should be packed in seawaterin double plastic bags with the airspace inthe bag filled with oxygen. The bags should besealed tightly with rubber bands or rubber ringsand packed inside a foam box. A small amountof ice may be added to keep the water cool,especially if a long transport time is expected.This box is then taped securely and may bepackaged inside a cardboard carton. Checkwith the diagnostic laboratory about packingrequirements. Some laboratories have specificpackaging requirements for diseased organisms.Samples submitted for certification purposesmay have additional shipping and collectionrequirements (see C.1.3.3).Label containers clearly:“LIVE SPECIMENS, STORE AT ___ to ___˚C, DONOT FREEZE”(insert temperature tolerance range of shrimpbeing shipped)If being shipped by air also indicate:“HOLD AT AIRPORT AND CALL FOR PICK-UP”• Clearly indicate the name and telephonenumber of the contact person responsible forpicking up the package at the airport or receivingit at the laboratory.• Where possible, ship early in the week toavoid arrival during the weekend which maylead to loss through improper storage ofsamples.• Inform the contact person as soon as theshipment has been sent and, where appropriate,give them the name of the carrier, theflight number, the waybill number and theestimated time of arrival.(Note: Some airlines have restrictions on shippingof seawater or preserved samples. It is agood idea to check with local airlines if they dohave any special requirements)1Ossiander, F.J. and G. Wedermeyer. 1973. Journal Fisheries Research Board of Canada 30:1383-1384.163

C.1 General TechniquesC.1.3.6 Preservation of Tissue Samples(Level 2)In some cases, such as locations remote froma diagnostic laboratory or where transport connectionsare slow, it may not be possible to providea live shrimp sample. Since freezing is usuallyinadequate for most diagnostic techniques(histology, bacteriology, mycology, etc.), specimensshould be fixed (chemical preservationto prevent tissue breakdown and decay) on site.This makes the sample suitable for subsequenthistological examination, in situ hybridization,PCR or electron microscopy, but will preventroutine bacteriology, mycology, virology or othertechniques requiring live micro-organisms. Diagnosticneeds should therefore be discussedwith the laboratory prior to collectingthe sample.The best general fixative for penaeid shrimp isDavidson’s fixative.330 ml 95% ethanol220 ml 100% formalin (37% w/v formaldehydein aqueous solution)115 ml glacial acetic acid335 ml distilled water.Mix and store at room temperature.(It should be noted, however, that formalin residuescan interfere with the PCR process.Samples for PCR analysis should be fixed in70% ethanol.)For any preservation procedure, it is essentialto remember that the main digestive organ ofthe shrimp (the hepatopancreas) is very importantfor disease diagnosis, but undergoes rapidautolysis (tissue digestion by digestive juicesreleased from the dying hepatopancreatic cells)immediately after death. This means that thepre-death structure of the hepatopancreas israpidly lost (turns to mush). Delays of even afew seconds in fixative penetration into this organcan result in the whole specimen beinguseless for diagnosis, thus, specimens must beimmersed or injected with fixative while stillalive. Dead shrimp, even when preserved onice (or frozen) are of no use for subsequent fixation.In tropical areas, it is best to use cold fixativethat has been stored in the freezer or kepton ice, as this helps arrest autolysis and secondarymicrobial proliferation, as the tissues arepreserved.Larvae and early post larvae (PL) should beimmersed directly in a minimum of 10 volumesof fixative to one volume of shrimp tissue. This10:1 ratio is critical for effective preservation.Attempts to cut costs by using lower ratios offixative to tissue can result in inadequate preservationof tissues for processing.For PL that are more than approximately 20 mmin length, use a fine needle to make a small,shallow incision that breaks and slightly lifts thecuticle in the midline of the back, at the cuticularjunction between the cephalothorax and firstabdominal segment. This allows the fixative topenetrate the hepatopancreas quickly.For larger PL’s, juveniles and adults, the fixativeshould be injected directly into the shrimp,as follows:• Place the shrimp briefly in ice water to sedatethem• Using surgical rubber gloves and protectiveeyeglasses, immediately inject the fixative(approximately 10% of the shrimp’s bodyweight) at the following sites (Fig. C.1.3.6):º hepatopancreasº region anterior to the hepatopancreasº anterior abdominal region, andº posterior abdominal region.Be careful to hold the shrimp so the angle ofinjection is pointed away from your body, sincefixative can sometimes spurt back out of aninjection site when the needle is removed andmay injure the eyes. It is also best to brace theinjection hand against the forearm of the handholding the shrimp, to avoid over penetrationof the needle into that hand. The hepatopancreasshould receive a larger proportion of theinjected fixative than the abdominal region. Inlarger shrimp it is better to inject the hepatopancreasat several points. All signs of lifeshould cease and the colour should change atthe injection sites.Immediately following injection, slit the cuticlewith dissecting scissors along the side of thebody from the sixth abdominal segment to thecuticle overlying the “head region” (cephalothorax).From there, angle the cut forward and upwarduntil it reaches the base of the rostrum.Avoid cutting too deeply into the underlying tissue.Shrimp over 12 g should be transverselydissected, at least once, posterior of the abdomen/cephalothoraxjunction and again midabdominally.The tissues should then be immersedin a 10:1 volume ratio of fixative to tissue,at room temperature. The fixative can bechanged after 24-72 hr to 70% ethanol, for longtermstorage.164

C.1 General TechniquesC.1.3.7 Shipping Preserved Samples(Level 1)For shipping, remove specimens from ethanolstorage, wrap in paper towel saturated with50% ethanol and place in a sealed plastic bag.There should be no free liquid in the bag. Sealand place within a second sealed bag. In mostcountries, small numbers of such specimenscan be sent to diagnostic laboratories by airmail.However, some countries or transportcompanies (especially air couriers) have strictregulations regarding shipping any chemicals,including fixed samples for diagnostic examination.Check with the post office or carrierbefore collecting the samples to ensure theyare processed and packed in an appropriateand acceptable manner. All sample bags shouldbe packed in a durable, leak-proof container.Label containers clearly with the name and telephonenumber of the contact person responsiblefor picking up the package at the airportor receiving it at the laboratory.If being shipped by air indicate - “HOLD AT AIR-PORT AND CALL FOR PICK-UP”.Where possible, ship early in the week to avoidarrival during the weekend which may lead toloss through improper storage of samples. Informthe contact person as soon as the shipmenthas been sent and, where appropriate,give them the name of the carrier, the flight number,the waybill number and the estimated timeof arrival.C.1.4 Record-Keeping (Level 1)Record-keeping is essential for effective diseasemanagement. For crustaceans, many ofthe factors that should be recorded on a regularbasis are outlined in sections C.1.1 andC.1.2. It is critical to establish and record normalbehaviour and appearance to compare withobservations during disease events.C.1.4.1 Gross Observations (Level 1)These could be included in routine logs of crustaceangrowth which, ideally, would be monitoredon a regular basis either by sub-samplingfrom tanks or ponds, or by “best-guess” estimatesfrom surface observations.For hatchery operations, the minimum essentialinformation which should be recorded/logged include:• feeding activity and feed rates• growth/larval staging• mortalities• larval conditionThese observations should be recorded on adaily basis for all stages, and include date, time,tank, broodstock (where there are more thanone) and food-source (e.g., brine shrimp culturebatch or other food-source). Dates andtimes for tank and water changes should alsobe noted, along with dates and times for pipeflushing and/or disinfection. Ideally, these logsshould be checked regularly by the person responsiblefor the site/animals.Where possible, hatcheries should invest in amicroscope and conduct daily microscopicexaminations of the larvae. This will allow themto quickly identify problems developing withtheir stocks, often before they become evidentin the majority of the population.For pond sites, the minimum essential observationswhich need to be recorded/logged include:• growth• feed consumption• fouling• mortalityThese should be recorded with date, site locationand any action taken (e.g., sample collectionfor laboratory examination). It is importantto understand that rates of change for theseparameters are essential for assessing thecause of any disease outbreak. This means levelshave to be logged on a regular and consistentbasis in order to detect patterns over time.Ideally, these logs should be checked regularlyby the person responsible for the site/animals.C.1.4.2 Environmental Observations(Level 1)This is most applicable to open ponds. Theminimum essential data that should be recordedare:• temperature• salinity• pH• turbidity (qualitative evaluation or secchi disc)• algal bloom(s)• human activity (treatments, sorting, pondchanges, etc.)• predator activity165

C.1 General TechniquesAs with C.1.4.1, types and rates of changes inthese parameters prior to any disease outbreaksare extremely important in assessing thecause of the outbreak. Although helpful, datarecorded on the day of specimen collection aremuch less useful than continuous records.Thus, the importance of keeping careful, regularand continuous records, regardless of the“expected” results, cannot be overstressed.Frequency of record-keeping will vary with siteand, possibly, season. For example, more frequentmonitoring may be required during unstableweather, compared to seasons with extended,stable, conditions.Human and predator activity should be loggedon an “as it happens” basis.C.1.4.3 Stocking Records (Level 1)All movements of crustaceans into and out of ahatchery and pond/site should be recorded.These should include:• the exact source of the broodstock or larvaeand any health certification history (e.g., resultsof any tests carried out prior to/on arrival)• condition on arrival• date, time and person responsible for receivingdelivery of the stock• date, time and destination of stock shippedout of the hatcheryEdition. Aquatic Animal Health Research Institute.Department of Fisheries. Bangkok,Thailand. 152p.Chanratchakool, P., J.F. Turnbull, S. Funge-Smith and C. Limsuan. 1995. Health Managementin Shrimp Ponds. Second Edition.Aquatic Animal Health Research Institute.Department of Fisheries. Bangkok, Thailand.111p.Lightner, D.V. 1996. A Handbook of ShrimpPathology and Diagnostic Procedures forDiseases of Cultured Penaeid Shrimp. WorldAquaculture Society, Baton Rouge, LA. 304p.Ossiander, F.J. and G. Wedermeyer. 1973. Computerprogram for sample size required todetermine disease incidence in fish populations.J. Fish. Res. Bd. Can. 30: 1383-1384.Wang,Y.G., K. L. Lee, M. Najiah, M. Shariff andM. D. Hassan. 2000. A new bacterial whitespot syndrome (BWSS) in cultured tigershrimp Penaeus monodon and its comparisonwith white spot syndrome (WSS) causedby virus. Dis. Aquat.Org. 41:9-18.In addition, all movements of stocks within ahatchery, nursery or grow out site should belogged with the date for tracking purposes if adisease situation arises.Where possible, animals from different sourcesshould not be mixed. If mixing is unavoidable,keep strict records of when mixing occurred.C.1.5 ReferencesAlday de Graindorge, V. and T.W. Flegel. 1999.Diagnosis of shrimp diseases with emphasison black tiger prawn, Penaeus monodon.Food and Agriculture Organization of theUnited Nations (FAO), Multimedia Asia Co.,Ltd, BIOTEC, Network of Aquaculture Centresin Asia Pacific (NACA) and SoutheastAsian Chapter of the World Aquaculture Society(WAS). Bangkok, Thailand. (InteractiveCD-ROM format).Chanratchakool, P., J.F. Turnbull, S.J. Funge-Smith, I.H. MacRae and C. Limsuan.1998.Health Management in Shrimp Ponds. Third166

VIRAL DISEASES OF SHRIMPC.2 YELLOWHEAD DISEASE (YHD) 1C.2.1 Background InformationC.2.2Clinical AspectsC.2.1.1 Causative AgentYellowhead disease (YHD) is caused byYellowhead Virus (YHV) (also reported in olderliterature as Yellowhead Baculovirus - YBV andYellowhead Disease Baculovirus - YHDBV). Itis now known not to be a member of theBaculoviridae. YHV is a single stranded RNA,rod shaped (44 ± 6 X 173 ±13 nm), envelopedcytoplasmic virus, likely related to viruses in theFamily Coronaviridae. Agarose gel electrophoresisindicates a genome size of approximately 22Kilobases. Lymphoid organ virus (LOV) and gillassociated virus (GAV) (see C.6) of Penaeusmonodon in Australia are related to the YHV complexviruses, although, of the two, only GAV isknown to cause mortality. More detailed informationabout the disease can be found in the OIEDiagnostic Manual for Aquatic Animal Diseases(OIE 2000a) and Lightner (1996).C.2.1.2 Host RangeNatural infections occur in Penaeus monodon,but experimental infections have been shown inP. japonicus, P. vannamei, P. setiferus, P. aztecus,P. duorarum and P. stylirostris. Penaeusmerguiensis, appear to be resistant to disease(but not necessarily infection). Palaemonstyliferus has been shown to be a carrier of viablevirus. Euphausia spp. (krill), Acetes spp. andother small shrimp are also reported to carry YHDviruses.C.2.1.3Geographic DistributionYHD affects cultivated shrimp in Asia includingChina PR, India, Philippines and Thailand.YHD has been reported from cultured shrimp inTexas and one sample has been reported to bepositive for YHV by antibody assay (Loh et al.1998).C.2.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System (1999-2000)YHD was reported in Malaysia in June, in thePhilippines in January to March and July; in SriLanka in January and suspected for the wholeyear of 1999 in Thailand. For the reporting periodfor the year 2000, India reported it in Octoberand it was suspected for the whole year inThailand and Sri Lanka (OIE 1999, OIE 2000b).Gross signs of disease (Fig.C.2.2) and mortalityoccur within 2 to 4 days following an intervalof exceptionally high feeding activity thatends in abrupt cessation of feeding. Mortalitiescan reach 100% within 3-5 days. Diseasedshrimp aggregate at the edges of the ponds ornear the surface. The hepatopancreas becomesdiscoloured which gives the cephalothorax ayellowish appearance, hence the name of thedisease. The overall appearance of the shrimpis abnormally pale. Post-larvae (PL) at 20-25days and older shrimp appear particularly susceptible,while PL

C.2 Yellowhead Disease (YHD)(TW Flegel)(DV Lightner)Fig.C.2.2. Gross sign of yellow head disease(YHD) are displayed by the three Penaeusmonodon on the left.a(DV Lightner)bFig.C.2.3.1.4c. Histological section of the gillsfrom a juvenile P. monodon with YHD. A generalizeddiffuse necrosis of cells in the gill lamellaeis shown, and affected cells display pyknoticand karyorrhectic nuclei (arrows). A few largeconspicuous, generally spherical cells with basophiliccytoplasm are present in thesection.These cells may be immaturehemocytes, released prematurely in responseto a YHV-induced hemocytopenia. Mayer-Bennett H&E. 1000x magnification.Fig.C.2.3.1.4a,b. Histological section of the lymphoid organ of a juvenile P. monodon with severeacute YHD at low and high magnification.A generalized, diffuse necrosis of LO cells isshown. Affected cells display pyknotic and karyorrhecticnuclei. Single or multiple perinuclearinclusion bodies, that range from pale to darklybasophilic, are apparent in some affected cells(arrows). This marked necrosis in acute YHDdistinguishes YHD from infections due to Taurasyndrome virus,which produces similar cytopathologyin other target tissues but not in theLO. Mayer-Bennett H&E. 525x and 1700x magnifications,respectively.C.2.3.1PresumptiveThere are no gross observations (Level I) or histopathological(Level II) diagnostic techniqueswhich can provide presumptive detection ofYHD in sub-clinical shrimp.168

C.2 Yellowhead Disease (YHD)C.2.3.2ConfirmatoryC.2.3.2.1 Reverse Transcriptase-PolymeraseChain Reaction Assay (Level III)For certification of YHV infection status ofbroodstock and fry, reverse transcriptase-polymerasechain reaction (RT-PCR) technology isrecommended.There are several commercially available RT-PCR kits now available to screen haemolymphfrom broodstock shrimp and PL tissues for evidenceof YHV RNA.C.2.4 Diagnostic MethodsMore detailed information on methods for diagnosiscan be found in the OIE Diagnostic Manualfor Aquatic Animal Diseases (OIE 2000a), at http://www.oie.int, or at selected references.C.2.4.1PresumptiveC.2.4.1.1 Gross Observations (Level 1)YHD can be suspected when an abnormal increasein feeding rates is followed by a sharpcessation in feeding. Moribund shrimp may appearnear the surface or edges of grow out pondsand show slow swimming behaviour in responseto stimuli. These may also show pale overall bodycolouration, a yellowish cephalothorax, pale gillsand hepatopancreas. YHD should be suspectedunder such circumstances, especially for P.monodon, and samples collected for confirmatorydiagnosis.C.2.4.1.2 Gill Squash (Level II)Fix whole shrimp, or gill filaments, in Davidson’sfixative overnight 2 . Wash gill filament in tap waterto remove the fixative and stain with Mayer-Bennett’s H&E. Clear in xylene and, using a finepair of needles (a stereo microscope is helpful),break off several secondary filaments and replacethe main filament in xylene for permanentreference storage in a sealed vial. Mountsecondary filaments, coverslip and use lightpressure to flatten the filaments as much aspossible, making them easy to see through. Thissame procedure can be used on thin layers ofsubcuticular tissue.Moderate to large numbers of deeply basophilic,evenly stained, spherical, cytoplasmic inclusionsapproximately 2 mm in diameter orsmaller are presumptive for YHD, along withsimilar observations from haemolymph smears.As with tissue sections and wet-fixed gill filaments,these slides can be kept as a permanentrecord.C.2.4.1.3 Haemolymph Smears (Level II)Smears that show moderate to high numbers ofblood cells with pycnotic and karyorrhexic nuclei,with no evidence of bacteria, can be indicativeof early YHD. It is important that no bacteriaare present, since these can produce similarhaemocyte nucleus changes. Such changes aredifficult to see in moribund shrimp because ofthe loss of blood cells so grossly normal shrimpshould be sampled for these signs from the samepond where the moribund shrimp were obtained.The haemolymph is collected in a syringe containingtwice the haemolymph volume of 25%formalin or modified Davidson’s fixative (i.e., withthe acetic acid component replaced by water orformalin). The blood cell suspension is mixedthoroughly in the syringe, the needle removedand a drop placed onto a microscope slide.Smear and air dry the preparation before stainingwith H&E and eosin or other standard bloodstains. Dehydrate, mount and coverslip. The resultsshould be consistent with the gill wholemounts (above) or histopathology of tissue sections,in order to make a presumptive YHD diagnosis.C.2.4.1.4 Histopathology (Level II)Fix moribund shrimp from a suspected YHD outbreakin Davidson’s fixative and process for standardH&E stain. Most tissues where haemolympis present may be infected, however, principalsites include the lymphoid organ (Oka organ)(Fig.C.2.3.1.4a,b), hepatopancreatic interstitialcells (not tubule epithelial cells), heart, midgutmuscle and connective tissue (but not epithelialcells), stomach sub-cuticulum and gill tissues(Fig.C.2.3.1.4c). Light microscopy should revealmoderate to large numbers of deeply basophilic,evenly stained, spherical, cytoplasmic inclusions,approximately 2 mm in diameter(smaller in ectodermal and mesodermal tissues).Moribund shrimp show systemic necrosisof gill and stomach sub-cuticular cells, with2If more rapid results are required, fixation can be shortened to 2 hours by substituting the acetic acid component of Davidson’sfixative with 50% concentrated HCl (this should be stored no more than a few days before use). After fixation, wash thoroughlyand check that the pH has returned to near neutral before staining. Do not fix for longer periods or above 25 o C as this may resultin excessive tissue damage that will make interpretation difficult or impossible.169

C.2 Yellowhead Disease (YHD)intense basophilic cytoplasmic inclusions (H&Estaining) due to phagocytosed nuclei and viralinclusions. In the lymphoid organ, high numbersof karyorrhexic and pyknotic basophilicinclusions are found in matrix cells of the normaltubules. On the other hand, similar inclusions–are found only in lymphoid organ spheroidswith Rhabdovirus of Penaeid Shrimp (RPS)described from Hawaii and Lymphoidal ParvolikeVirus (LPV, LOV) described from Australia;Lymphoid Organ Vacuolisation Virus (LOVV) inP. vannamei in Hawaii and the Americas; andTaura Syndrome Virus (TSV) in P. vannamei, P.stylirostris and P. setiferus from central andsouth America. Gill Associated Virus (GAV) inAustralian P. monodon; a Yellow-Head-Disease-Like Virus (YHDLV) in P. japonicus from TaiwanProvince of China produce similar histopathologyto YHV.C.2.4.2 ConfirmatoryIn cases where results from presumptive screeningindicate possible YHD infection, but confirmationof the infectious agent is required (e.g.,first time finding or presence of other pathogenicfactors), bioassay (see C.2.4.2.1), electron microscopy(see C.2.4.2.2) and molecular techniques(see C.2.4.2.3-5) are required.C.2.4.2.1 Bioassay (Levels I-II)The simplest bioassay method is to allow na?veshrimp (± 10 g wet weight) to feed on carcassesof suspect shrimp. Alternatively, preparehomogenates of gill tissues from suspect shrimp.Centrifuge solids into a loose pellet, decant andfilter (0.45 - 0.22 mm) the supernatant. Exposena?ve juvenile Penaeus monodon (± 10 g wetweight) to the supernatant Infected shrimp shouldevoke clinical signs in the na?ve shrimp within24-72 hours and 100% mortality will generallyoccur within 3-5 days. Infections should be confirmedby histology of gills and haemolymph.C.2.4.2.2 Transmission Electron Microscopy(TEM) (Level III)For TEM, the most suitable tissues of moribundshrimp suspected to be infected by YHD arethe lymphoid organ and gills. Fix tissues in 2.5%glutaraldehyde, 2% paraformaldehyde in cacodylatebuffer and post-fix in 1% osmiumtetroxide, prior to dehydration and embeddingin Spurr’s resin. 50nm sections are mounted onCu-200 grids and should be stained with uranylacetate/70% methanol and Reynold’s leadcitrate. Diagnosis of YHV is confirmed by thepresence of non-occluded, enveloped, rodshapedparticles, 150-200 x 40-50 nm in sizein the perinuclear or cytoplasmic area of thetarget tissues or within cytoplasmic vesicles.Non-enveloped, filamentous forms measuring

C.2 Yellowhead Disease (YHD)• infected individuals and their offspring bedestroyed in a sanitary manner• associated equipment and rearing water aredisinfected• exclude potential carriers of YHD by screeningPL pre-stocking in ponds• prevention of exposure to potential carriers,post-stocking, can be achieved by filtrationor prior treatment in storage ponds of waterused for water exchanges.• avoidance of rapid changes in pH or prolongedperiods of low ( 9 should be avoided. Changes in salinityapparently do not trigger outbreaks.• avoid fresh aquatic feeds in grow-out ponds,maturation units and hatchery facilities, unlessthe feed is subjected to prior sterilization(gamma radiation) or pasteurization (i.e.,holding at 7˚C for 10 min).If an outbreak occurs, it is recommended that theaffected pond be treated with 30 ppm chlorine tokill the shrimp and potential carriers. The deadshrimp and other animals should be removed andburied or burned. If they cannot be removed, thepond should be thoroughly dried before restocking.If the outbreak pond can be emergency harvested,the discharge water should be pumpedinto an adjacent pond for disinfection with chlorineand holding for a minimum of 4 days beforedischarge. All other waste materials should beburied or burned. Harvesting personnel shouldchange clothing and shower at the site with waterthat will be discharged into the treatment pond.Clothing used during harvesting should be placedin a specific container to be sent for chlorine treatmentand laundering. Equipment, vehicles andrubber boots and the outside of shrimp containersshould be disinfected with chlorine and thedischarge water run into the treatment pond.Neighbours should be notified of any YHD outbreakand control efforts, and advised not carryout any water exchange for at least 4 days followingdischarge from the pond used for disinfection.Processing plants receiving emergencyharvested shrimp should be notified that thespecific lot of shrimp is YHV infected and appropriatemeasures should be taken at the plantto avoid transfer of the disease via transportcontainers and processing wastes. Prohibitionof introduction of living shrimp from YHV andGAV enzootic areas into historically uninfectedareas is recommended.C.2.6 Selected ReferencesKhanobdee, K., C. Soowannayan, T.W. Flegel,S. Ubol, and B. Withyachumnarnkul. 2001.Evidence for apoptosis correlated with mortalityin the giant black tiger shrimp Penaeusmonodon infected with yellow head virus.Dis. Aquat. Org. (in press).Lightner, D.V. 1996. A Handbook of Shrimp Pathologyand Diagnostic Procedures for Diseaseof Cultured Penaeid Shrimp. WorldAquaculture Society, Baton Rouge, LA. 304p.Loh, P.C., E.C.B. Nadala, Jr., L.M. Tapay, andY. Lu. 1998. Recent developments in immunologically-basedand cell culture protocolsfor the specific detection of shrimp viralpathogens, pp. 255-259. In: Flegel T.W. (ed)Advances in Shrimp Biotechnology. NationalCenter for Genetic Engineering and Biotechnology,Bangkok, Thailand.Lu, Y., L.M. Tapay, and P.C. Loh. 1996. Developmentof a nitrocellullose-enzyme immunoassayfor the detection of yellow-head virusfrom penaeid shrimp. J. Fish Dis. 19(1):9-13.Nadala, E.C.B. Jr., L.M. Tapay, S. Cao, and P.C.Loh. 1997. Detection of yellowhead virus andChinese baculovirus in penaeid shrimp by thewestern blot technique. J. Virol. Meth.69(1-2): 39-44.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Spann, K.M., J.E. Vickers, and R.J.G. Lester.1995. Lymphoid organ virus of Penaeusmonodon from Australia. Dis. Aquat. Org.23(2): 127-134.Spann, K.M., J.A. Cowley, P.J. Walker, andR.J.G. Lester. 1997. A yellow-head-like virusfrom Penaeus monodon cultured in Australia.Dis.Aquat. Org. 31(3): 169-179.171

C.2 Yellowhead Disease (YHD)Wang, C.S., K.F.J.Tang, G.H. Kou, S.N. Chen.1996. Yellow head disease like virus infectionin the Kuruma shrimp Peneaus japonicuscultured in Taiwan. Fish Pathol. 31(4): 177-182.Wongteerasupaya, C., V. Boonsaeng, S.Panyim,A.Tassanakajon,B.Withyachumnarnkul, and T.W. Flegel.1997. Detection of yellow-head virus (YHV)of Penaeus monodon by RT-PCR amplification.Dis. Aquat. Org. 31(3): 181-186.172

C.3 INFECTIOUS HYPODERMAL ANDHAEMATOPOIETIC NECROSIS (IHHN)C.3.1 Background InformationC.3.1.1 Causative AgentInfectious Hypodermal and Hematopoietic Necrosis(IHHN) is caused by a non-enveloped icosahedralvirus, Infectious Hypodermal and HematopoieticNecrosis Virus (IHHNV), averaging 22 nmin diameter, with a density of 1.40 g/ml in CsCl,containing linear ssDNA with an estimated sizeof 4.1 kb, and a capsid that has four polypeptideswith molecular weights of 74, 47, 39, and37.5 kD. Because of these characteristics,IHHNV has been classified as a member of thefamily Parvoviridae. More detailed informationabout the disease can be found at OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000a) and Lightner (1996).C.3.1.2 Host RangeIHHNV infects a wide range of penaeid shrimps,but does not appear to infect other decapod crustaceans.Natural infections have been reportedin Penaeus vannamei, P. stylirostris, P.occidentalis, P. monodon, P. semisulcatus, P.californiensis and P. japonicus. Experimental infectionshave also been reported for P. setiferus,P. aztecus and P. duorarum. Penaeus indicusand P. merguiensis appear to be refractory toIHHNV infection.C.3.1.3 Geographic DistributionIHHN occurs in wild and cultured penaeid shrimpsin Central America, Ecuador, India, Indonesia,Malaysia, Philippines, Peru, Taiwan Province ofChina, and Thailand. Although IHHNV has beenreported from cultured penaeid shrimp from mostregions of the western hemisphere and in wildpenaeids throughout their geographic range alongthe Pacific coast of the Americas (Peru to northernMexico), it has not been found in penaeidson the Atlantic side of the Americas. IHHNV hasbeen reported in cultured penaeid shrimp fromGuam, French Polynesia, Hawaii, Israel and NewCaledonia. An IHHN-like virus has also been reportedfrom Australia.C.3.1.4 Asia-Pacific Quarterly Aquatic AnimalDisease reporting System (1999-2000)The disease was suspected in India during the2nd quarter reporting period for 1999 and 1stquarter reporting period for 2000 (OIE 1999, OIE2000b).C.3.2 Clinical AspectsPenaeus stylirostris. Infection by IHHNV causesacute epizootics and mass mortality (> 90%) inP. stylirostris. Although vertically infected larvaeand early postlarvae do not become diseased,juveniles >35 days old appear susceptibleshowing gross signs followed by massmortalities. In horizontally infected juveniles, theincubation period and severity of the diseaseappears size and/or age dependent, with youngjuveniles always being the most severely affected(Fig.C.3.2a). Infected adults seldomshow signs of the disease or mortalities.Penaeus vannamei. The chronic disease, “runtdeformity syndrome” (RDS) (Fig.C.3.2b,c), iscaused by IHHNV infection of P. vannamei. Juvenileswith RDS show wide ranges of sizes,with many smaller than average (“runted”)shrimp. Size variations typically exceed 30%from the mean size and may reach 90%.Uninfected populations of juvenile P. vannameiusually show size variations of < 30% of themean. Similar RDS signs have been observedin cultured P. stylirostris.C.3.3 Screening MethodsMore detailed information on methods forscreening IHHN can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000a), at http://www.oie.int, or at selected references.C.3.3.1 PresumptiveThere are no gross signs (Level I) or histologicalfeatures (Level II) that can be used to indicatepresumptive infection by IHHNV in subclinicalcarriers.C.3.3.2ConfirmatoryMolecular methods are required to detectIHHNV in sub-clinical carriers.C.3.3.2.1 Dot Blot Hybridization (Level III)Haemolymph samples or a small appendage(pleiopod) can be used for dot blot testing.Commercial dot blot hybridization kits for IHHNare now available.173

C.3 Infectious Hypodermal AndHaematopoietic Necrosis (IHHN)(DV Lightner)Fig.C.3.2a. A small juvenile Penaeus stylirostrisshowing gross signs of acute IHHN disease.Visible through the cuticle, especially on the abdomen,are multifocal white to buff colored lesionsin the cuticular epithelium or subcutis (arrows).While such lesions are common in P.stylirostris with acute terminal IHHN disease,they are not pathognomonic for IHHN disease.(DV Lightner)(DV Lightner)(DV Lightner)Fig.C.3.2c.Lateral viewof juvenile P.vannamei(preserved inDavidson’sAFA) showinggross signs ofI H H N V -caused RDS.Cuticular abnormalitiesofthe sixth abdominalsegmentand tailfan are illustrated.Fig.C.3.2b. Dorsal view of juvenile P. vannamei(preserved in Davidson’s AFA) showing grosssigns of IHHNV-caused RDS. Cuticular abnormalitiesof the sixth abdominal segment andtail fan are illustrated.Fig.C.3.4.1.2b. A low magnification photomicrograph(LM) of an H&E stained section of ajuvenile P. stylirostris with severe acute IHHNdisease. This section is through the cuticularepithelium and subcuticular connective tissuesjust dorsal and posterior to the heart. Numerousnecrotic cells with pyknotic nuclei or withpathognomonic eosinophilic intranuclear inclusionbodies (Cowdry type A) are present (arrows).Mayer-Bennett H&E. 830x magnification.(DV Lightner)Fig.C.3.4.1.2a. A high magnification of gillsshowing eosinophilic intranuclear inclusions(Cowdry type A inclusions or CAIs) that arepathognomonic for IHHNV infections. Mayer-Bennett H&E. 1800x magnification.>174

C.3 Infectious Hypodermal AndHaematopoietic Necrosis (IHHN)C.3.3.2.2 Polymerase Chain Reaction(PCR) (Level III)The same tissue samples described in C.3.3.2.1can be used for non-lethal screening of nonclinicalbroodstock and juveniles of susceptiblespecies, using PCR.C.3.4 Diagnostic MethodsMore detailed information on methods for diagnosisof IHHN can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000b), at http://www.oie.int, or at selected references.C.3.4.1 PresumptiveC.3.4.1.1 Gross Observations (Level I)Gross signs are not IHHN specific. Acute infectionsof juvenile P. stylirostris may result in amarked reduction in food consumption, followedby changes in behaviour and appearance.The shrimp may rise slowly to the watersurface, become motionless and then roll-over,and slowly sink (ventral side up) to the bottom.This behavior may continue for several hoursuntil the shrimp become too weak to continue,or are cannibalised by healthier siblings. By thisstage of infection white or buff-coloured spots(which differ from the white spots that occur inWSD - C.4) in the cuticular epidermis, especiallyat the junction of the abdominal tergalplates, resulting in a mottled appearance. Thismottling may later fade in P. stylirostris. MoribundP. stylirostris may further develop a distinctlybluish colour and opaque abdominalmusculature. Although P. monodon is frequentlyfound to be infected with IHHNV, it does notgenerally appear to cause any major clinicaldisease in the species. Juvenile shrimp (P.vannamei and P. stylirostris) with RDS displaybent or deformed rostrums, wrinkled antennalflagella, cuticular roughness, and other cuticulardeformities. They also show a high percentage(30-90%) of stunted growth (“runt shrimp”)compared with less than 30% below averagesize in uninfected populations.C.3.4.1.2 Histopathology (Level II)Infected cells occur in the gills (Fig.C.3.4.1.2a),epidermal (Fig.C.3.4.1.2b) and hypodermal epitheliaof fore and hindgut, nerve cord and nerveganglia, as well as mesodermal haematopoieticorgans, antennal gland, gonads, lymphoid organ,and connective tissue. Eosinophilic (withH&E stain) intranuclear, Cowdry type A inclusionbodies (CAIs) provide a presumptive diagnosisof IHHNV infection. Infected nuclei areenlarged with a central eosinophilic inclusionsometimes separated from the marginatedchromatin by an unstained ringwhen tissues arepreserved with acetic acid containing fixatives.Since IHHNV intranuclear inclusion bodies canbe confused with developing intranuclear inclusionbodies due to White Spot Disease, electronmicroscopy (C.3.4.2.2) or in situ hybridizationassays of suspect sections with IHHNVspecificDNA probes (C.3.4.2.3-5) may be requiredfor definitive diagnosis. Basophilicstrands may be visible within the CAIs and cytoplasmicinclusion bodies may also be present.C.3.4.2ConfirmatoryC.3.4.2.1 Bioassay (Levels I/II)Prevalence and severity of IHHNV infectionsmay be “enhanced” in a quarantined populationby holding the suspect shrimp in crowdedor other stressful conditions (low dissolved oxygen,elevated water temperature, or elevatedammonia or nitrite). These conditions may encourageexpression of low grade IHHNV infectionsand transmission from sub-clinical carriersto uninfected shrimp. This increase in prevalenceand severity can enhance detection usingscreening methods.Indicator shrimp (0.1-4.0 gm juvenile P.stylirostris) can also be used to assess the presenceof IHHNV by cohabitation, feeding ofminced carcasses or injection with cell-freehomogenates from suspect shrimp.C.3.4.2.2 Transmission Electron Microscopy(TEM) (Level III)Negative stain preparations of purified virusshow non-enveloped, icosahedral virions, 20-22 nm in diameter. Transmission electron microscopicpreparations show intranuclear inclusionscontaining virions 17-26 nm in diameter.Viral particles are also present in the cytoplasmwhere they assemble and replicate. Chromatinstrands (that may be visible as basophilic inclusionsunder light microscopy) are a prominentfeature of IHHNV intranuclear inclusionbodies. Paracrystalline arrays of virions correspondto cytoplasmic inclusion bodies that maybe detected under light microscopy.175

C.3 Infectious Hypodermal AndHaematopoietic Necrosis (IHHN)C.3.4.2.3 Dot Blot Hybridization (Level III)As described in C.3.3.2.1.C.3.4.2.2 Polymerase Chain Reaction(Level III)As described in C.3.3.2.2.C.3.4.2.5 In situ Hybridization (Level III)IHHNV-specific DNA probes are now availablefor in situ hybridization confirmation of histologicaland/or electron microscopic observation.C.3.5 Modes of TransmissionSome members of populations of P. stylirostrisand P. vannamei, which survive IHHNV infectionsand/or epizootics, may carry sub-clinicalinfections for life which may be passed horizontallyto other stocks, or vertically, if used asbroodstock.C.3.6 Control MeasuresEradication methods for IHHNV can be appliedto certain aquaculture situations. These methodsare dependent upon eradication of infectedstocks, disinfection of the culture facility, avoidanceof re-introduction of the virus (from othernearby culture facilities, wild shrimp, etc.), andre-stocking with IHHNV-free post-larvae thathave been produced from IHHNV-freebroodstock.C.3.7 Selected ReferencesBell, T.A. and D.V. Lightner, D.V. 1984. IHHN virus:Infectivity and pathogenicity studies inPenaeus stylirostris and Penaeus vannamei.Aquac. 38: 185-194.Bray, W.A., A.L. Lawrence, and J.R. Leung-Trujillo. 1994. The effect of salinity on growthand survival of Penaeus vannamei, with observationson the interaction of IHHN virusand salinity. Aquac. 122(2-3): 133-146.Browdy, C.L., J.D. Holloway, Jr., C.O. King, A.D.Stokes, J.S. Hopkins, and P.A. Sandifer. 1993.IHHN virus and intensive culture of Penaeusvannamei: Effects of stocking density andwater exchange rates. Crus. Biol.13(1): 87-94.Carr, W.H., J.N. Sweeney, L. Nunan, D.V.Lightner, H.H. Hirsch, and J.J. Reddington.1996. The use of an infectious hypodermaland hematopoietic necrosis virus gene probeserodiagnostic field kit for screening of candidatespecific pathogen-free Penaeusvannamei broodstock. Aquac.147(1-2): 1-8.Castille, F.L., T.M. Samocha, A.L. Lawrence, H.He, P. Frelier, and F. Jaenike. 1993. Variabilityin growth and survival of early postlarvalshrimp (Penaeus vannamei Boone 1931).Aquac. 113(1-2): 65-81.Karunasagar, I. and I. Karunasagar. 1996.Shrimp diseases and control. AquacultureFoundation of India, Madras, India 1996: 63-67Lightner, D.V. 1996. A Handbook of ShrimpPathology and Diagnostic Procedures forDisease of Cultured Penaeid Shrimp. WorldAquaculture Society, Baton Rouge, LA. 304p.Lu, Y., P.C. Loh, and J.A. Brock. 1989. Isolation,purification and characterisation ofinfectious hypodermal and hematopoietic necrosisvirus (IHHNV) from penaeid shrimp. J.Virol. Meth. 26: 339-344.Mari, J., J.R. Bonami, and D.V. Lightner. 1993.Partial cloning of the genome of infectioushypodermal and hematopoietic necrosis virus,an unusual parvovirus pathogenic forpenaeid shrimps - diagnosis of the diseaseusing a specific probe. J. Gen. Vir.74(12):2637-2643.Nunan, L.M., B. Poulos, and D.V. Lightner. 1994.Detection of the infectious hypodermal andhematopoietic necrosis virus (IHHNV) inPenaeus shrimp tissue homogenate andhemolymph using polymerase chain reaction(PCR). International Symposium on AquaticAnimal Health: Program and Abstracts. Universityof California, School of VeterinaryMedicine, Davis, CA, USA. 1994: P-62.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.176

C.3 Infectious Hypodermal AndHaematopoietic Necrosis (IHHN)OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Owens, L., I.G. Anderson, M. Kenway, L. Trott,and J.A.H. Benzie. 1992. Infectious hypodermaland haematopoietic necrosis virus(IHHNV) in a hybrid penaeid prawn from tropicalAustralia. Dis. Aquat. Org. 14: 219-228.Poulos, B.T., D.V. Lightner, B. Trumper, and J.R.Bonami. 1994. Monoclonal antibodies to apenaeid shrimp parvovirus, infectious hypodermaland hematopoeitic necrosis virus(IHHNV). J. Aquat. Anim. Health 6(2): 149-154.177

C.4 WHITE SPOT DISEASE (WSD) 3C.4.1 Background InformationC.4.1.1 Causative AgentThe causative agent of white spot disease(WSD) is the white spot syndrome virus (WSSV)or white spot virus (WSV), a double strandedDNA (dsDNA) virus. In initial reports, WSV wasdescribed as a non-occluded baculovirus butsubsequent analysis of WSV-DNA sequencesdoes not support this contention. The virusesin this complex have recently been shown tocomprise a new group with the proposed nameof Nimaviridae (Van Hulten et al. 2001). In theliterature, however, several names have beenused to describe the virus, including baculoviralhypodermal and haematopoietic necrosis(HHNBV), Shrimp Explosive Epidemic Disease(SEED), China virus disease, rod-shapednuclear virus of Penaeus japonicus (RV-PJ);systemic ectodermal and mesodermalbaculovirus (SEMBV), white spot baculovirus(WSBV) and white spot syndrome virus (WSSV).More detailed information about the diseasecan be found in the OIE Manual for AquaticAnimal Diseases (OIE 2000a) and Lightner(1996).C.4.1.2 Host RangeWhite spot disease has a wide spectrum of hosts.Outbreaks were first reported from farmedPenaeus japonicus in Japan and natural infectionshave subsequently been observed in P.chinensis, P. indicus, P. merguiensis, P. monodon,P. setiferus, P. stylirostris, and P. vannamei. Inexperimental studies, WSD is also lethal to P.aztecus, P. duodarum and P. setiferus.C.4.1.3 Geographic DistributionWSD was first reported in Taiwan Province ofChina and China mainland between 1991-1992,and in Japan in 1993 from shrimp imported fromChina PR. Later outbreaks have been reportedfrom elsewhere in Asia including China PR, India,Indonesia, Korea RO, Malaysia, TaiwanProvince of China, Thailand, and Vietnam. Inaddition to the Asian countries listed above,farmed shrimp exhibiting the gross signs and histologyof WSD have been reported in the USAand Latin America.gua, Panama, Peru and USA (Subasinghe etal. 2001).C.4.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System (1999-2000)WSD was reported by Bangladesh, China PR,India, Indonesia, Japan, Korea RO, Malaysia,Philippines, Taiwan Province of China, SriLanka, and Thailand; and suspected in Pakistanduring the reporting period for the year1999. In year 2000, Bangladesh, India, Japan,Korea RO, Malaysia, Philippines, Sri Lanka,Thailand and Vietnam reported positive occurrenceof the disease (NACA/FAO 2000a,b,c; OIE1999, OIE 2000a,b).C.4.2 Clinical AspectsWSD outbeaks are often characterised by highand rapid mortality of infected populations,usually shortly after the first appearance of theclinical signs. Acutely affected shrimp demonstrateanorexia and lethargy, have a loose cuticlewith numerous white spots (about 0.5 to2.0 mm in diameter) on the inside surface ofthe carapace (Fig.C.4.2a,b). These spots arewithin the cuticle structure and cannot be removedby scraping. Moribund shrimp may alsoshow a pink to red discolouration. Susceptibleshrimp species displaying these clinical signsare likely to undergo high levels of mortality.Pathology is associated with systemic destructionof the ectodermal and mesodermal tissuesof the gills and sub-cuticular tissues.C.4.3 Screening MethodsMore detailed information on methods forscreening for WSD can be found in the OIE DiagnosticManual for Aquatic Animal Diseases(OIE 2000a), at http://www.oie.int, or in selectedreferences.C.4.3.1 PresumptiveThere are no gross observations (Level I) or histopathological(Level II) diagnostic techniqueswhich can provide presumptive detection ofWSD in sub-clinical shrimp.As of 1999, WSD has been reported in at leastnine countries in the Americas: Columbia, Ecuador,Guatemala, Honduras, Mexico, Nicara-3White spot disease (WSD) is now classified as an OIE Notifiable Disease (OIE 2000a).178

C.4 White Spot Disease (WSD)(DV Lightner)(DV Lightner)Fig.C.4.2a. A juvenile P. monodon with distinctivewhite spots of WSD.(DV Lightner/P. Saibaba)Fig.C.4.3.3.1.2a. Histological section from thestomach of a juvenile P.chinensis infected withWSD. Prominent intranuclear inclusion bodiesare abundant in the cuticular epithelium andsubcuticular connective tissue of the organ (arrows).(DV Lightner)Fig.C.4.2b. Carapace from a juvenile P.monodon with WSD. Calcareous deposits onthe underside of the shell account for the whitespots.C.4.3.2ConfirmatoryC.4.3.2.1 Nested PCR of Tissues andHaemolymph (Level III)The protocol described by Lo et al (1996, 1998)is the recommended procedure for nested PCRof tissues and haemolymph. There are alsocommercially available kits for detection of WSDin sub-clinical carriers using PCR-based techniques.C.4.3.2.2 Polymerase Chain Reaction(PCR) of Postlarvae (Level III)From a nursery or hatchery tank containing 100000 postlarvae (PL) or more, sample approximately1000 PL from each of 5 different points.Fig.C.4.3.3.1.2b. Section of the gills from a juvenileP. chinensis with WSBV. Infected cellsshow developing and fully developed intranuclearinclusion bodies of WSBV (arrows).Mayer-Bennett H&E. 900x magnification.Pool the samples in a basin, gently swirl thewater and select an assay sample from livingPL collected at the center of the basin. A sampleof 150 PL is required to give a 95% confidenceof detecting an infection at 2% prevalence inthe population (see Table C.1.3.3 of C.1 GeneralTechniques).For PL 11 and older, exclude shrimp eyes fromany tissue samples, since these inhibit the PCRprocess. Follow the procedures from the recommendedsource for nested PCR given underC.4.3.2.1.179

C.4 White Spot Disease (WSD)C.4.3.2.3 Dot Blot Hybridization (Level III)Details on dot blot hybridisation techniques anddetection kit availability are provided in the OIEDiagnostic Manual (OIE 2000a).C.4.3.2.4 In situ Hybridization (Level III)Details on in situ hybridization techniques anddetection kit availability are provided in the OIEDiagnostic Manual (OIE 2000a).C.4.4 Diagnostic MethodsMore detailed information on methods for diagnosisof WSD can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000a), at http://www.oie.int, or in selected references.C.4.4.1PresumptiveC.4.4.1.1 Gross Observations (Level I)WSD outbreaks are generally preceded by cessationof feeding followed, within a few days,by the appearance of moribund shrimp swimmingnear the surface at the edge of rearingponds. These shrimp exhibit white inclusionsembedded in the cuticle and often show reddishdiscolouration of the body. The cuticularinclusions range from minute spots to discsseveral mm in diameter that may coalesce intolarger plaques. They are most easily observedby removing the cuticle from the cephalothorax,scraping away any attached tissue andholding the cuticle up to the light. The appearanceof white spots in the cuticle can be causedby other conditions. In particular, Wang et al.,2000, report a condition called bacterial whitespot syndrome (BWSS) which can easily bemistaken for WSD (see C.4a). Therefore, histopathologicalexamination is required for confirmatorydiagnosis.C.4.4.1.2 Rapid Squash Mount Preparations(Level II)Two types of rapid squash mount preparationsthat can be used for presumptive diagnosis ofWSD: i) fresh, unstained wet mounts fixed in10% formalin solution and viewed by dark fieldmicroscopy with a wet-type condenser, and ii)fixed tissues stained with H&E.For method ii) fix whole shrimp or gill filamentsin Davidson’s fixative overnight. If more rapidresults are required, fixation can be shortenedto 2 hrs by changing the acetic acid in theDavidson’s fixative to 50% concentrated HCl(this should not be stored longer than a few daysbefore use). After fixation, wash the tissues thoroughlyand ensure pH is near neutral beforestaining. Do not fix for longer periods, or above25 o C, as this can cause tissue damage that willmake interpretation difficult or impossible. Stainwith Meyer’s H&E and dehydrate to xylene (orequivalent clearing solution). Place a gill filamenton a microscope slide tease off severalsecondary filaments. Replace the main filamentin a sealed vial filled with xylene as a permanentback-up reference. Being careful not tolet the secondary gill filaments dry, tease apartand remove any large fragments or particlesfrom the slide. Add a drop of mounting fluidand a cover glass, using light pressure to flattenthe tissue as much as possible. The sameprocedure can be used for thin layers of subcuticulartissue.Examine under a compound microscope at 40xmagnification for moderate to large numbersof hypertrophied nuclei with basophilic, centrally-positioned,inclusions surrounded bymarginated chromatin. The whole mount slidescan also be kept as permanent records.C.4.4.1.3 Histopathology (Level II)Moribund shrimp from a suspected WSD outbreakshould be fixed in Davidson’s fixative andstained with haematoxylin and eosin (H&E). Thehistopathology of WSD is distinctive, and canprovide a conclusive diagnosis. However, firsttime detection or detection in species not previouslyreported to be susceptible, requiremolecular assay or electron microscopy demonstrationof a viral aetiology.Moribund shrimp with WSV show systemicdestruction of ectodermal and mesodermal tissues.Nuclei of infected cells are hypertrophiedand when stained with haematoxylin and eosinshow lightly to deeply basophilic central inclusionssurrounded by marginated chromatin.These intranuclear inclusions can also be seenin squash mounts of gills or sub-cuticular tissue(see C.4.4.1.2), or in tissue sections. Thebest tissues for examination are the subcuticulartissue of the stomach (Fig.C.4.3.3.1.2a),cephalothorax or gill tissues (Fig.C.4.3.3.1.2b).C.4.4.2 ConfirmatoryA definitive diagnosis can be accomplished bypolymerase chain reaction (PCR) technology180

C.4 White Spot Disease (WSD)(single-step or nested), in situ hybridization,Western blot analysis (detailed protocols canbe found in OIE (2000a) or electron microscopy(TEM).C.4.4.2.5 Transmission Electron Microscopy(TEM) (Level III)The most suitable tissues for TEM examinationare subcuticular tissues, gills and pereiopodsthat have been pre-screened by histology(C.4.4.1.3) or rapid-stain tissue squashes(C.4.4.1.2) which show signs of hypertrophiednuclei with Cowdry A-type inclusions or marginatedchromatin surrounding a basophilic inclusionbody. Fix tissues for at least 24h in a10:1 fixative to tissue volume ration of 6%gluteraldehyde at 4°C and buffered with sodiumcacodylate or phosphate solution to pH7. Forlonger term storage, reduce gluteraldehyde to0.5-1.0% concentration. Post-fix in 1% osmiumtetroxide, and stain with uranyl acetate and leadcitrate (or equivalent TEM stain). WSD virionsare rod-shaped to elliptical with a trilaminarenvelope and measure 80-120 x 250-380 nm.C.4.4.2.6 Negative Stain Electron Microscopy(Level III)Negative stain preparations from shrimphaemolymph may show virions with unique, taillikeappendages within the hypertrophied nucleiof infected cells, but no evidence of occlusionbodies.C.4.5 Modes of TransmissionWild broodstock and fry used to stock rearingponds are known to carry WSV, as are numerousother crustaceans and even aquatic insectlarvae. Molecular techniques have been usedto confirm infection of non-penaeid carriers ofWSV and transmission studies show that thesecan transmit WSV to shrimp.C.4.6 Control MeasuresThere are no known treatments for shrimp infectedwith WSV, however, a number of preventativemeasures are recommended to reducespread.At facilities used for the production of PL, it isrecommended that wild broodstock bescreened for WSD by nested PCR. Any infectedindividuals, and their offspring, should be destroyedin a sanitary manner and all contaminatedequipment and rearing water be disinfected.It is also recommended that broodstockP. monodon be tested for WSD after spawningto increase the probability of viral detection.At grow-out, PL should be screened for freedomfrom WSV by nested PCR using sufficientlylarge numbers of PL to ensure detection of significantinfections. A biased sampling regime,which selects weaker animals for testing, canfurther increase the probability of detecting infectedbatches.During cultivation, it is suspected that rapidchanges in water temperature, hardness andsalinity, or reduced oxygen levels (

C.4 White Spot Disease (WSD)to avoid transfer of the disease via transportcontainers and processing wastes. Preventionof introduction of live shrimp from WSV enzooticareas into historically uninfected areas or areasdefined as free from the disease is recommended.C.4.7 Selected ReferencesChou, H.Y., C.Y. Huang, C.H. Wang, H.C.Chiang and C.F. Lo. 1995. Pathogenicity of abaculovirus infection causing white spot syndromein cultured penaeid shrimp in Taiwan.Dis. Aquat. Org. 23: 165-173.Inouye, K, S. Miwa, N. Oseko, H. Nakano, T.Kimura, K. Momoyama and M. Hiraoka.1994. Mass mortalities of cultured Kurumashrimp Penaeus japonicus in Japan in 1993:electron microscopic evidence of the causativevirus. Fish Pathol. 29:149-158.Lightner, D.V. 1996. A Handbook of ShrimpPathology and Diagnostic Procedures forDisease of Cultured Penaeid Shrimp. WorldAquaculture Society, Baton Rouge, LA. 304p.Lo, C.F., Y.S. Chang, C.T. Cheng, and G.H.Kou 1998. PCR monitoring of cultured shrimpfor white spot syndrome virus (WSSV) infectionin growout ponds. In: Flegel T.W. (ed)Advances in shrimp biotechnology, pp. 281-286. National Center for Genetic Engineeringand Biotechnology. Bangkok, Thailand.Lo, C.F., J.H. Leu , C.H. Ho, C.H. Chen, S.E.Peng, Y.T. Chen, C.M. Chou, , P.Y. Yeh , C.J.Huang, H.Y. Chou, C.H. Wang, and G.K. Kou.1996. Detection of baculovirus associatedwith white spot syndrome (WSBV) in penaeidshrimps using polymerase chain reaction.Dis. Aquat. Org. 25: 133-141.Network of Aquaculture Centres in Asia-Pacificand Food and Agriculture Organization of theUnited Nations. 2000a. Quarterly Aquatic AnimalDisease Report (Asia and Pacific Region),2000/1, January-March 2000. FAO ProjectTCP/RAS/6714. Bangkok, Thailand. 57p.Network of Aquaculture Centres in Asia-Pacificand Food and Agriculture Organization of theUnited Nations. 2000b. Quarterly AquaticAnimal Disease Report (Asia and Pacific Region),2000/2, April-June 2000. FAO ProjectTCP/RAS/6714. Bangkok, Thailand. 59p.Network of Aquaculture Centres in Asia-Pacificand Food and Agriculture Organization of theUnited Nations. 2000c. Quarterly Aquatic AnimalDisease Report (Asia and Pacific Region),2000/3, July-September 2000. FAO ProjectTCP/RAS/6714. Bangkok, Thailand. 57p.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Subasinghe, R.P., M.G. Bondad-Reantaso, andS.E. McGladdery. 2001. Aquaculture development,health and wealth. In: R.P.Subasinghe, P. Bueno, M.J. Phillips, C.Hough, S.E. McGladdery & J.R. Arthur, eds.Aquaculture in the Third Millennium. TechnicalProceedings of the Conference onAquaculture in the Third Millennium,Bangkok, Thailand, 20-25 February 2000.NACA, Bangkok and FAO, Rome. (in press)Van Hulten, M.C. , J. Witteveldt, S. Peters, N.Kloosterboer, R. Tarchini, M. Fiers, H.Sandbrink, R.K. Lankhorst, and J.M. Vlak.2001 The white spot syndrome virus DNAgenome sequence. Virol. 286 (1):7-22.Wang, C.H., C.F. Lo, J.H. Leu, C.M. Chou, M.C.Tung, C.F. Chang, M.S. Su and G.H. Kou.1995. Purification and genomic analysis ofbaculoviruses associated with white spotsyndrome (WSBV) of Penaeus monodon. Dis.Aquat. Org. 23:239-242.Wongteerasupaya, C., J.E. Vickers, S.Sriurairatana, G.L. Nash, A. Akarajamorn, V.Boonsaeng, S. Panyim, A. Tassanakajon, B.Withyachumnarnkul and T.W. Flegel. 1995.A non-occluded, systemic baculovirus thatoccurs in cells of ectodermal and mesodermalorigin and causes high mortality in theblack tiger prawn, Penaeus monodon. Dis.Aquat. Org. 21:69-77.182

C.4a BACTERIAL WHITE SPOTSYNDROME (BWSS)Bacterial White Spot Syndrome (BWSS) is arecently described condition which affectsPenaeus monodon. It is, as yet, poorly understoodcondition and is included in the AsiaDiagnostic Guide due to the possibility of diagnosticconfusion with viral White Spot Disease(WSD).C.4a.1 Background InformationSince 1993, white spot disease virus (WSDV)has caused massive losses to the shrimp industryin Asia and Latin America. Recently, anotherdisease syndrome showing similar gross clinicalsigns of white spots, has been detected and reportedas “bacterial white spot syndrome’’(BWSS) (Wang et al., 1999, 2000). The similargross clinical signs have also caused confusionduring PCR-based screening for WSD since,shrimp with apparent WSDV clinical signs, givenegative results. The clinical effects of BWSS,appear far less significant than those of WSDinfection, although it has been suggested thatsevere infections may reduce moulting andgrowth.C.4a.1.1 Causative Agent(s)The bacterium Bacillus subtilis has been suggestedas the possible causative agent due toits association with the white spots (Wang et al.,2000) but no causal relationship has been demonstrated,nor have infectivity studies been conducted.Vibrio cholerae is also often isolated insignificant numbers and similar white spots havebeen described in farmed shrimp in Thailand asa result of exposure to high pH and alkalinity inponds in the absence of the White Spot virus orbacterial colonisation of the spots, indicating thatthe bacterial involvement may be secondary. Thelack of certainty as to the causative agent andthe possibility of secondary involvement of bacterianeeds to be addressed through further research.Until the bacterial etiology is clearly demonstrated,bacteria cannot be definitively regardedas the causative agent.C.4a.1.2 Host RangeTo date, the syndrome has only been reported incultured Penaeus monodon.C.4a.1.3 Geographic DistributionBWSS was first detected from a shrimp (Penaeusmonodon) farm in Malaysia in 1998 (Wang et al.1999, 2000). This remains the only confirmedreport of the condition.C.4a.2Clinical AspectsDull white spots are seen on the carapace andall over the body but are more noticeable whenthe cuticle is peeled away from the body. Thewhite spots are rounded and not as dense asthose seen in WSD (Fig.C.4a.2). Wet mountmicroscopy reveals the spots as opaque brownishlichen-like lesions with a crenellated margin(although this is also the case with spots in theearly stages of WSD and cannot be used as adistinctive diagnostic feature). The spot centeris often eroded and even perforated. During theearly stage of infection, shrimp are still active,feeding and able to moult – at which point thewhite spots may be lost. However, delayedmoulting, reduced growth and low mortalitieshave been reported in severely infected shrimp(Wang et al., 2000).C.4a.3Screening MethodsThere are no reported methodologies availableto screen for sub-clinical infections, sinceBWSS appears to be an opportunistic infection.C.4a.4Diagnostic MethodsC.4a.4.1 PresumptiveC.4a.4.1.1 Gross Observations (Level I)The presence of white spots on shrimp cuticleswithout significant mortality.C.4a.4.1.2 Wet Mounts (Level I)If cuticular spots are detected in P. monodon,which show an opaque brownish lichen-likeappearance with a crenallated margin and thecenter shows signs of erosion and/or perforation,along with extensive bacterial involvement,such infections could be attributable to BWSS.Such infections should be confirmed as beingnegative for WSD.C.4a.4.1.2 Polymerase Chain Reaction(PCR) (Level III)Negative WSDV-PCR results from samplesshowing gross clinical signs attributed to WSD,may be suggestive of the alternate aetiology ofBWSS.183

C.4a Bacterial White SpotSyndrome (BWSS)(M. Shariff)C.4a.4.2 ConfirmatoryC.4a.4.2.1 Histopathology (Level II)Fig. C.4a.2. Penaeus monodon dense whitespots on the carapace induced by WSD.(M. Shariff/ Wang et al. 2000 (DAO 41:9-18))Histological examinations should be conductedto ensure that the soft-tissues associated withthe cuticular lesions do not show signs of theWSDV characteristic endodermal and mesodermalintranuclear inclusian bodies. In the caseof BWSS, bacteria will be the primary microbialforeign particle and this should be in primaryassociation with the cuticular lesionsthemselves.C.4a.4.2.2 Scanning Electron Microscopy(TEM) (Level III)The presence of spot lesions (Fig. C.4a.4.2.1a,b)together with numerous bacteria (Fig.C.4a.4.2.2c) under scanning electron microscopywill confirm BWSS.C.4a.5 Modes of TransmissionaSince bacteria are only localized on the bodysurface, the mode of transmission is thoughtto be through the rearing water. However, thishas yet to be demonstrated using transmissionstudies.C.4a.6 Control MeasuresbFig. C.4a.4.2.2a, b. Bacterial white spots(BWS), which are less dense than virus-inducedwhite spots. Note some BWS have a distinctwhitish marginal ring and maybe with or withouta pinpoint whitish dot in the center(M. Shariff/ Wang et al. 2000 (DAO 41:9-18))Although the exact aetiology is unknown, somemeasures may help to reduce the risk of BWSS.Build up of high bacterial density in rearingwater should be avoided. Changing water frequentlyis recommended . Indiscriminate useof probiotics containing Bacillus subtilis shouldalso be avoided until the relationship betweenthis bacteria and the BWSS syndrome is betterunderstood. It has been claimed that BWSS inshrimp ponds can be treated with quick lime(CaO) at 25 ppm, however, this is still under investigationand the use of quicklime may itselfcause problems due to rapid increases in pondwaterpH (see C.4.6).>Fig. C.4a.4.2.2c. Presence of large number ofbacteria attached to exposed fibrillar laminaeof the endocuticle.184

C.4a Bacterial White SpotSyndrome (BWSS)C.4a.7 Selected ReferencesWang, Y.G., M. Shariff, K.L. Lee and M.D.Hassan. 1999. A review on diseases ofcultured shrimp in Malaysia. Paper waspresented at Workshop on Thematic Reviewon Management Strategies for MajorDiseases in Shrimp Aquaculture, 28-30November 1999, Cebu, Philippines. WB,NACA, WWF and FAO.Wang, Y. G., K.L. Lee, M. Najiah, M. Shariffand M.D. Hassan. 2000. A new bacterialwhite spot syndrome (BWSS) in cultured tigershrimp Penaeus monodon and its comparisonwith white spot syndrome (WSS) causedby virus. Dis. Aquat. Org. 41: 9-18.185

C.5 BACULOVIRAL MIDGUT GLANDNECROSIS (BMN)C.5.1Background InformationC.5.3.1 PresumptiveC.5.1.1Causative AgentThe pathogen responsible for Baculoviral MidgutGland Necrosis (BMN) disease is Baculoviralmidgut gland necrosis virus (BMNV), a non-occludedgut-infecting baculovirus, whose non-envelopednucleocapsid measures 36 by 250 nm;enveloped virions measures ~ 72 by ~ 310 nm.More detailed information about the disease canbe found in the OIE Diagnostic Manual for AquaticAnimal Diseases (2000a), and Lighter (1996).Techniques suitable for presumptive screeningof asymptomatic animals at Levels I or II arenot available.C.5.3.2 ConfirmatoryC.5.3.2.1 Histopathology (Level II)Histopathology as described for C.5.4.2.1 is thestandard screening method recommended byOIE (2000a).C.5.1.2Host RangeC.5.4 Diagnostic MethodsBMN was observed as natural infections inPenaeus japonicus, P. monodon and P. plebejus(Fig.C.5.1.2a); and as experimental infections inP. chinensis and P. semisulcatus.C.5.1.3Geographic DistributionBMN has occurred in the Kyushu and Chugokuarea of Japan since 1971. BMN-like virus (nonoccluded,type C baculovirus) has also been reportedin P. japonicus in Korea RO and fromP. monodon in the Philippines and possibly inAustralia and Indonesia.C.5.1.4 Asia-Pacific Quarterly AquaticAnimal Diseases Reporting System (1999-2000)For the reporting year 1999, no positive reportfrom Japan (1992 was last year of occurrence).The disease was suspected in Korea RO fromJanuary to September 1999 and whole year of2000 (OIE 1999, OIE 2000a).C.5.2Clinical AspectsIn Japan, BMN is considered to be one of themajor problems in hatcheries where it infects larvaeand early postlarval stages causing highmortalities. The apparent white turbidity of thehepatopancreas is caused by necrosis of hepatopancreastubule epithelium and possibly also themucosal epithelium. Larvae float inactively butlater stages (late PL) tend to show resistance tothe disease.C.5.3Screening MethodsMore detailed information on methods for screeningBMN can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE 2000a),at http://www.oie.int, or at selected references.More detailed information on methods for diagnosiscan be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000a), at http://www.oie.int, or at selected references.C.5.4.1 PresumptiveC.5.4.1.1 Gross Observations (Level 1)Morbid or larvae heavily infected with BMNVshows a cloudy midgut gland, easily observableby the naked eye.C.5.4.1.2 Wet-Mount Technique (Level II)Hypertrophied nuclei in fresh squashes (viewedunder dark-field microscopy) or in stainedsmears of hepatopancreas (using light microscopy)are demonstrated in BMNV infectedsamples. When viewed under dark-field illuminationequipped with a wet-type condenser, theinfected nuclei appear white against the darkbackground. This is due to the increased reflectedand diffracted rays produced by numerousvirus particles in the nucleus. Samples fixedin 10% formalin also give same results.C.5.4.2 ConfirmatoryC.5.4.2.1 Histopathology (Level II)Samples are fixed in Davidson’s fixative, stainedwith standard H&E and examined under brightfield microscopy. Infected shrimps show greatlyhypertropied nuclei (Fig.C.5.4.2.1a) inhepatopancreatic epithelial cells undergoingnecrosis. Infected nuclei show diminishednuclear chromatin, marginated chromatin (Fig.C.5.4.2.1b, c) and absence of occlusion bodiescharacteristic of Baculovirus penaei (BP) (seealso Fig. C.9.3.2.3a,b – section C.9) and186

C.5 Baculoviral Midgut GlandNecrosis (BMN)(DV Lightner)(DV Lightner)bFig.C.5.1.2a. Section of the hepatopancreas ofP. plebejus displaying several hepatopancreascells containing BMN-type intranuclear inclusionbodies. Mayer-Bennett H&E. 1700 x magnification.(DV Lightner)cFig.C.5.4.2.1a. High magnification of hepatopancreasfrom a PL of P. monodon with a severeinfection by a BMN-type baculovirus. Mostof the hepatopancreas cells display infectednuclei. Mayer-Bennett H&E. 1700x magnification.(DV Lightner)Fig. C.5.4.2.1b, c. Sections of the hepatopancreasof a PL of P. japonicus with severe BMN.Hepatopancreas tubules are mostly destroyedand the remaining tubule epithelial cells containmarkedly hypertrophied nuclei that containa single eosinophilic to pale basophilic, irregularlyshaped inclusion body that fills the nucleus.BMNV infected nuclei also display diminishednuclear chromatin, marginated chromatin andabsence of occlusion bodies that characterizeinfections by the occluded baculoviruses.Mayer-Bennett H&E. Magnifications: (a) 1300x;(b) 1700x.>Fig.C.5.4.2.1d. MBV occlusion bodies whichappear as esosinophilic, generally multiple,spherical inclusion bodies in enormously hypertrophiednuclei (arrows). Mayer-BennettH&E. 1700x magnification.187

C.5 Baculoviral Midgut GlandNecrosis (BMN)Monodon Baculovirus (MBV) infections(Fig.C.5.4.2.1d).C.5.4.2.2 Transmission Electron Microscopy(TEM) (Level III)Tranmission electron microscopy can be usedconfirm diagnosis of BMN through demonstrationof the rod-shaped enveloped virions asdescribed in C.5.1.1.C.5.4 Modes of TransmissionThe oral route has been demonstrated to bethe main infection pathway for BMNV infection.Viruses released with faeces into the environmentalwater of intensive culture systems ofP. japonicus play an important role in diseasespread.C.5.5 Control MeasuresThe concentrations of various disinfectants requiredto kill BMNV are toxic to shrimp larvae.Complete or partial eradication of viral infectionmay be accomplished by thorough washingof fertile eggs or nauplii using clean seawater to remove the adhering excreta. Disinfectionof the culture facility and the avoidanceof re-introduction of the virus are critical factorsto control BMN disease.The suggested procedure for eradication ofBMN infection involves collection of fertile eggsfrom broodstock and passing them through asoft gauze with pore size of 800 mm to removedigested excrement or faeces of the shrimp.The eggs are then washed with running seawater at salinity level of 28-30% for 3-5 min tomake sure all the faecal debris has been removed.The eggs are then collected by passingthe suspension through a soft gauze withpore size of 100 mm. The eggs are then furtherwashed with running sea water at salinity levelof 28-30% for 3-5 min to remove the adhesiveviral particles.Dis. 8:585-589.Natividad, J.M. and D.V. Lightner. 1992. Prevalenceand geographic distribution of MBVand other diseases in cultured giant tigerprawns (Penaeus monodon) in the Philippines,pp.139-160. In: Diseases of CulturedPenaeid Shrimp in Asia and the UnitedStates, Fulks, W. and Main, K.L (eds.). TheOceanic Institute, Honolulu, Hawaii, USA.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.Park, M.A. 1992. The status of culture and diseasesof penaeid shrimp in Korea, pp. 161-167. In: Diseases of Cultured Penaeid Shrimpin Asia and the United States, Fulks, W. andMain, K.L (eds.). The Oceanic Institute, Honolulu,Hawaii, USA.Sano, T. and K. Momoyama. 1992. Baculovirusinfection of penaeid shrimp in Japan, pp. 169-174. In: Diseases of Cultured Penaeid Shrimpin Asia and the United States, Fulks, W. andMain, K.L (eds.). The Oceanic Institute, Honolulu,Hawaii, USA.Sano, T. T. Nishimura, K. Oguma, K. Momoyamaand N. Takeno. 1981. Baculovirus infectionof cultured Kuruma shrimp Penaeusjaponicus in Japan. Fish Pathol. 15:185-191.C.5.6 Selected ReferencesLightner, D.V. 1996. A Handbook of ShrimpPathology and Diagnostic Procedures forDisease of Cultured Penaeid Shrimp. WorldAquaculture Society, Baton Rouge, LA. 304p.Momoyama, K. and T. Sano. 1989. Developmentalstages of kuruma shrimp larvae,Penaeus japonicus Bate, with baculoviralmid-gut gland necrosis (BMN) virus. J. Fish188

C.6 GILL-ASSOCIATED VIRUS (GAV)C.6.1 BackgroundC.6.1.1 Causative AgentGill-associated virus (GAV) is a single-strandedRNA virus related to viruses of the familyCoronaviridae. It is closely related to yellow headvirus and is regarded as a member of the yellowhead complex. GAV can occur in healthy ordiseased shrimp and was previously called lymphoidorgan virus (LOV) when observed inhealthy shrimp.C.6.1.2 Host RangeNatural infection with GAV has only been reportedin Penaeus monodon but experimental infectionhas caused mortalities in P. esculentus, P.merguiensis and P. japonicus. An age or sizerelated resistance to disease was observed in P.japonicus.C.6.1.3 Geographic DistributionGAV has only been recorded from Queenslandon the north-east coast of Australia and is endemicto P. monodon in this region.C.6.1.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System(1999-2000)Australia reported widespread occurrence of LOVamong healthy farmed and wild P. monodon inQueensland. Other countries reported “no informationavailable” for GAV for the reporting periodfor 1999 and 2000 (OIE 1999, OIE 2000).C.6.2 Clinical AspectsGAV is endemic in healthy P. monodon in northernQueensland. It is unclear whether the onsetof disease results from environmental stress leadingto clinical expression of the pre-existing virusas can occur with YHD and WSD or whetherthe disease arises from a new infection with apathogenic strain of GAV. GAV is predominantlyfound in the gill and lymphoid organ but has alsobeen observed in haemocytes. During acute infections,there is a rapid loss of haemocytes, thelymphoid organs appear disorganised and devoidof normal tubule structure, and the virus is detectedin the connective tissues of all major organs.C.6.3C.6.3.1Screening MethodsConfirmatoryC.6.3.1.1 Reverse Transcriptase-PolymeraseChain Reaction (RT-PCR) (Level III)The PCR primers below are designed to amplifya 618 bp region of GAV:GAV-5 5’-AAC TTT GCC ATC CTC GTCAC-3’GAV-6 5’-TGG ATG TTG TGT GTT CTCAAC-3’The PCR primers below are designed to amplifya 317 bp region internal to the region amplifiedby GAV5 and GAV6:GAV-1 5’-ATC CAT ACT ACT CTA AAC TTCC-3’GAV-2 5’-GAA TTT CTC GAA CAA CAGACG-3’Total RNA (100 ng) is denatured in the presenceof 35 pmol of each primer (GAV-5 andGAV-6) by heating at 98°C for 8 min in 6 mlDEPC-water containing 0.5 ml deionisedformamide and quenched on dry ice. cDNA issynthesised by the addition of 2 ml SuperscriptII buffer x 5, 1 ml 100 mM DTT, 0.5 ml 10 mMdNTPs, 20 U rRNasinTM (Promega) and 100 USuperscript II Reverse Transcriptase (Life Technologies)and DEPC-water to 10 ml and the reactionis incubated at 42 o C for 1 hr followed byheating at 99 o C for 5 min before quenching onice. One tenth of the cDNA reaction (1 ml = 10ng RNA) is amplified in 50 ml using Taq buffer(10 mM Tris-HCl pH 9.0, 50 mM KCl, 0.1% TritonX-100), 1.5 mM MgCl 2, 35 pmol each primerGAV-5 and GAV-6 and 200 mM dNTPs overlaidwith 50 ml liquid paraffin. PCRs are initiatedusing a “hot-start” protocol in which the reactionwas heated at 85oC for 5 min prior to theaddition of 2.5 U Taq polymerase (Promega).DNA is amplified by 30 cycles of 95oC/1 min,58 o C/1 min, 72 o C/40 sec followed by 72 o C/10min final extension and 20oC hold using eithera Corbett Research or Omnigene (Hybaid) thermalcycler. PCR products (10 ml) are resolvedin 2% agarose-TAE gels containing 0.5 mg/mlethidium bromide.When the result of the primary RT-PCR is negativeor inconclusive, 0.5 ml of the primary PCRis amplified by nested PCR as above in a 50 mlreaction volume using primers GAV-1 and GAV-2. In some cases, 5 ml of the RT-PCR is used.Nested PCR conditions are as for the primaryPCR except that the extension time is reducedto 30 sec and number of cycles is reduced to20. Nested PCR aliquots (10 ml) are analysedin 2% agarose-TAE gels.189

C.6 Gill-Associated Virus (GAV)C.6.4 Diagnostic Methods(P Walker)C.6.4.1 PresumptiveC.6.4.1.1 Gross Observations (Level I)Shrimp with an acute GAV infection demonstratelethargy, lack of appetite and swim onthe surface or around the edge of ponds. Thebody may develop a dark red colour particularlyon the appendages, tail fan and mouthparts; gills tend to be yellow to pink in colour.Barnacle and tube worm attachment togetherwith gill fouling have also been observed. Thegross signs of acute GAV infection are variableand not always seen and thus, they are not reliable,even for preliminary diagnosis.C.6.4.1.2 Cytology/Histopathology(Level II)The cephalothorax of infected prawns is separatedfrom the abdomen and split longtudinally.The sample is then fixed in Davidson’s fixativeand processed for histology. Sections arestained with H&E. Lymphoid organs from diseasedshrimp display loss of the normal tubulestructure. Where tubule structure is disrupted,there is no obvious cellular or nuclear hypertrophy,pyknotic nuclei or vacuolization. Foci ofabnormal cells are observed within the lymphoidorgan and these may be darkly eosinophilic.The gills of diseased shrimp displaystructural damage including fusion of gill filamenttips, general necrosis and loss of cuticlefrom primary and secondary lamellae. The cytologyof the gills appears normal apart fromsmall basophilic foci of necrotic cells.C.6.4.2ConfirmatoryC.6.4.2.1 Transmission Electron Microscopy(TEM) (Level III)Tissue samples are fixed in 2.5% glutaraldehyde/2%paraformaldehyde in cacodylatebuffer and post-fixed in 1% osmium tetroxide.Fixed samples are then dehydrated through agraded series of ethanol concentrations andmounted in Spurr’s resin. 50 nm sections aremounted on Cu-200 grids, stained with uranylacetate/70% methanol and Reynold’s lead citrate.The cytoplasm of lymphoid organ cellsfrom diseased shrimps contains both rodshapedenveloped virus particles and viralnucleocapsids. The nucleocapsids are from166-435 nm in length 16-18 nm in width.Fig. C.6.4.2.1. Transmission electron microscopyof GAV.Nucleocapsids have striations with a periodicityof 7 nm and are often found associated withthe endoplasmic reticulum. Enveloped virionsare less common, occurring in about 20% ofcells within the disrupted areas of the lymphoidorgan. The enveloped virions (Fig. C.6.4.2.1) are183-200 nm long and 34-42 nm wide againassociated with the endoplasmic reticulum.Both enveloped virions and nucleocapsids arepresent in gill tissue but the nucleocpsids aremore commonly occurring in 40-70% of cellswhereas enveloped virions are present in lessthan 10% of cells.C.6.4.2.2 Reverse Transcriptase-PolymeraseChain Reaction (RT-PCR) (Level III)As described for C.6.3.1.1.C.6.5 Modes of TransmissionThe most effective form of horizontal transmissionis direct cannibalism but transmission canalso be water-borne. GAV is also transmittedvertically from healthy broodstock. The virusmay be transmitted from either or both parentsbut it is not clear if infection is within the egg.C.6.6 Control MeasuresThere are no known control measures for GAV.Prevention of the movement of GAV infectedstock into historically uninfected areas is recommended.Drying out of infected ponds appearseffective in preventing persistence of thevirus.190

C.6 Gill-Associated Virus (GAV)C.6.7Selected ReferencesCowley, J.A., C.M. Dimmock, C.Wongteerasupaya, V. Boonsaeng, S. Panyamand P.J. Walker. 1999.Yellow head virus fromThailand and gill-associated virus from Australianare closely related but distinct viruses.Dis. Aquat. Org. 36:153-157.Cowley, J.A., C.M. Dimmock, K.M. Spann andP.J. Walker. 2000b. Gill-associated virus ofPenaeus monodon prawns: an invertebratevirus with ORF1a and ORF 1b genes relatedto arteri- and coronaviruses. J. Gen. Virol. 81:1473 – 1484.Spann, K.M., J.E. Vickers and R.J.G.Lester.1995. Lymphoid organ virus ofPenaeus monodon from Australia. Dis.Aquat. Org. 23: 127-134Spann, K.M., J.A. Cowley, P.J. Walker andR.J.G. Lester.1997. A yellow-head-like virusfrom Penaeus monodon cultured in Australia.Dis. Aquat. Org. 31: 169-179.Spann, K.M., A.R. Donaldson, I.J. East, J.A.Cowley and P.J. Walker. 2000. Differencesin the susceptibility of four penaeid prawnspecies to gill-associated virus (GAV). Dis.Aquat. Org. 42: 221-225.Walker, P.J., J.A. Cowley, K.M. Spann, R.A.J.Hodgson, M.A. Hall and B.Withyachumnernkul. 2001. Yellow head complexviruses: transmission cycles and topographicaldistribution in the Asia-Pacific region,pp. 227-237. In: C.L. Browdy and D.E.Jory (eds).The New Wave: Proceedings of theSpecial Session on Sustainable Shrimp Culture,Aquaculture 2001. The World AquacultureSociety, Baton Rouge, LA.191

C.7 SPAWNER-ISOLATEDMORTALITY VIRUS DISEASE(SMVD) 4C.7.1 Background InformationC.7.1.1 Causative AgentSpawner-isolated Mortality Virus Disease(SMVD) is caused by a single-stranded icosahedralDNA virus measuring 20-25 nm. Thesecharacteristics are most closely associated withthose of the Family Parvoviridae. The virus hasbeen named Spawner-isolated Mortality Virus(SMV) and other disease names include SpawnerMortality Syndrome (SMS) and Midcrop MortalitySyndrome (MCMS). More detailed informationabout the disease can be found in OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000a).C.7.1.2 Host RangeSMVD affects Penaeus monodon. Experimentalinfections have also resulted in mortalities of P.esculentus, P. japonicus, P. merguiensis andMetapenaeus ensis. Moribund, farmed freshwatercrayfish (Cherax quadricarinatus) have alsobeen associated with putative SMV infection usingDNA-probe analyses.C.7.1.3 Geographic DistributionSMVD has been reported from Queensland, aswell as the Philippines and Sri Lanka.C.7.1.3.4 Asia-Pacific Quarterly AquaticAnimal Disease Reporting System (1999-2000)Most countries reported “no information available”or “never reported” for the 2 year reporting period(1999 and 2000) except for Sri Lanka whichsuspected the disease in August 1999 and reportedpositive occurrence in September 1999(OIE 1999, OIE 2000b). Philippines reported positiveoccurrence of SMV in October to December1998 where samples of P. monodon sent to Australiafor insitu hybridization using SMV probeproduced positive results (NACA/FAO 1999).from the Philippines were also infected with luminousvibriosis (Vibrio harveyi).C.7.3 Screening MethodsMore detailed information on methods forscreening SMVD can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000a), at http://www.oie.int, or at selected references.There are no standard screening methods availablefor asymptomatic animals.C.7.4 Diagnostic MethodsMore detailed information on methods for diagnosisof SMVD can be found in the OIE DiagnosticManual for Aquatic Animal Diseases(OIE 2000a), at http://www.oie.int, or at selectedreferences.C.7.4.1PresumptiveC.7.4.1.1 Gross Observations (Level 1)There are no specific clinical signs for SMVD.Juvenile P. monodon in grow-out ponds mayshow discolouration, lethargy, fouling and anorexia.Since this may be caused by severalviral or bacterial infections, however, other diagnosticmethods are required.C.7.4.1.2 Cytology/Histopathology(Level II)The histopathology associated with SMVD isnot disease specific. In naturally infected juvenileP. monodon, haemocyte infiltration andcytolysis is focussed around the enteric epithelialsurfaces. Experimental infections, using tissueextracts from shrimp with SMVD developsystemic infections manifest by systemichaemocytic infiltration, necrosis and sloughingof epithelial cells of the midgut and hepatopancreas.C.7.2 Clinical AspectsC.7.4.2ConfirmatoryThere are no specific clinical signs known forSMV. It is one of several viruses associated withmid-crop mortality syndrome (MCMS) which resultedin significant mortalities of juvenile andsub-adult P. monodon cultured in Australia from1994 to 1996. Similarly affected P. monodonC.7.4.2.1 Transmission Electron Microscopy(TEM) (Level III)SMV virions are found in the gut epithelial tissues.The viral particles measure approximately20-25 nm in diameter and have hexagonal4This disease is listed in the current FAO/NACA/OIE Quarterly Aquatic Animal Disease Reporting System as Spawner mortalitysyndrome (‘Midcrop mortality syndrome’).192

C.7 Spawner-IsolatedMortality Virus Disease (SMVD)(icosahedral) symmetry.C.7.5 Modes of TransmissionMoribund and dead individuals are cannibalisedby surviving animals, which is assumed to facilitatehorizontal transmission.C.7.6 Control MeasuresPrevention of introduction of shrimp from SMVinfected stock into historically uninfected areasis recommended. Daily removal of moribundanimals from ponds, particularly early in production,has also been recommended. Stockingof ponds with progeny of spawners withSMV-negative faecal testing using PCR-probeshas been shown to reduce mortality by 23%.Owens, L. and C. McElnea. 2000. Natural infectionof the redclaw crayfish Cheraxquadricarinatus with presumptive spawnerisolatedmortality virus. Dis. Aquat. Org. 40:219-233.Owens, L., G. Haqshenas, C. McElnea and R.Coelen. 1998. Putative spawner-isolatedmortality virus associated with mid-crop mortalitysyndrome in farmed Penaeus monodonfrom northern Australia. Dis. Aquat. Org. 34:177-185.C.7.7 Selected ReferencesAlbaladejo, J.D., L.M. Tapay, V.P. Migo, C.G.Alfafara, J.R. Somga, S.L. Mayo, R.C.Miranda, K. Natividad, F.O. Magbanua, T.Itami, M. Matsumura, E.C.B. Nadala, Jr. andP.C. Loh. 1998. Screening for shrimp virusesin the Philippines, pp. 251-254. In: Advancesin shrimp Biotechnology, Flegel, T.W. (ed).National Center for Genetic Engineering andBiotechnology. Bangkok, Thailand.Fraser, C.A. and L. Owens. 1996. Spawner-isolatedmortality virus from Australian Penaeusmonodon. Dis. Aquat. Org. 27: 141-148.NACA/FAO. 1999. Quarterly Aquatic AnimalDisease Report (Asia-Pacific Region), 98/2,October to December 1998. FAO ProjectTCP/RAS/6714. Bangkok, Thailand. 41p.OIE. 1999. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 35p.OIE. 2000a. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.OIE. 2000b. Regional Aquatic Animal DiseaseYearbook 1999 (Asian and Pacific Region).OIE Representation for Asia and the Pacific.Tokyo, Japan. 40p.193

C.8 TAURA SYNDROME (TS) 5C.8.1 Background InformationC.8.1.1 Causative AgentTaura Syndrome (TS) is caused by a virus, TauraSyndrome Virus (TSV) tentatively classified asa member of the Picornaviridae based on itsmorphology (31- 32 nm non-enveloped icosahedron),cytoplasmic replication, buoyant densityof 1.338 g/ml, genome consisting of a linear,positive-sense ssRNA of approximately 10.2 kbin length, and a capsid comprised of three major(55, 40, and 24 kD) and one minor (58 kD)polypeptides. More detailed information aboutthe pathogen can be found in the OIE DiagnosticManual for Aquatic Animal Diseases(2000) and Lightner (1996).C.8.1.2 Host RangeTSV infects a number of American penaeid species.The most susceptible species is the Pacificwhite shrimp Penaeus vannamei, althoughP. stylirostris, and P. setiferus can also be infected.Post-larvae and juvenile P. schmittii, P. aztecus,P. duorarum, P. chinensis, P. monodon, andMarsupenaeus (Penaeus) japonicus have beeninfected experimentally.C.8.1.3Geographic DistributionTaura Syndrome was first detected in shrimpfarms near the Taura River, Ecuador (hence thename of the disease) in 1992. It then spreadthroughout most shrimp growing regions of LatinAmerica including Hawaii (infections successfullyeradicated ) and the Pacific coasts of Colombia,Costa Rica, Ecuador, El Salvador, Guatemala,Honduras, Mexico, Nicaragua, Panama, andPeru.TSV has also been reported from cultured shrimpalong the Atlantic coasts of Belize, Brazil, Columbia,Mexico, and Venezuela and the southeasternU.S. states of Florida, South Carolina andTexas. TSV has, however, been successfullyeradicated from cultured stocks in Florida andBelize. TSV is found in wild penaeids in Ecuador,El Salvador, Honduras, and Mexico. The onlyrecord of TSV in the eastern hemisphere is fromTaiwan, Province of China, where the diseasewas likely introduced with P. vannamei from CentralAmerica.C.8.2 Clinical AspectsTaura Syndrome is particularly devastating topost-larval P. vannamei within approximately 14to 40 days of stocking into grow-out ponds ortanks, however, larger stages may also be severelyaffected. Three distinct phases characterizeTS disease progression: i) the acute stage,during which most mortalities occur; ii) a brieftransition phase, and iii) a chronic ‘carrier’ stage.In the acute phase, the cuticular epithelium isthe most severely affected tissue. In the chronicphase, the lymphoid organ becomes the predominantsite of infection. In P. vannamei, theacute phase of infection may result in highmortalities (40-90%), while most strains of P.stylirostris appear resistant to fatal levels ofinfection. Survivors of acute TSV infection passthrough a brief transition phase and enter thechronic phase which may persist for the rest oftheir lives. This sub-clinical phase of infectionis believed to have contributed to the spreadof the disease via carriage of viable TSV.C.8.3 Screening MethodsDetailed information on methods for screeningTSV can be found in the OIE Diagnostic Manualfor Aquatic Animal Diseases (OIE 2000), at http://www.oie.int, or at selected references.C.8.3.1 PresumptiveC.8.3.1.1 Gross Observation (Level I)Any Penaeus vannamei, or other susceptiblepenaeid survivors of a TS outbreak, should beconsidered suspect carriers of TSV. However,there are no gross observation or Level I signsthat can be used to screen sub-clinical carriers.C.8.3.1.2 Histopathology (Level II)Post-larvae, juveniles and adults can bescreened using routine histological techniquesand stains. Chronic stages of infection arecharacterised by the presence of spherical accumulationsof cells in the lymphoid organ, referredto as ‘lymphoid organ spheroids’ (LOS).These masses are composed of presumed phagocytichemocytes, which have sequesteredTSV and aggregate within intertubular spacesof the lymphoid organs.5Taura Syndrome (TS) is now classified as an OIE Notifiable Disease (OIE 2000).194

C.8 Taura Syndrome (TS)C.8.3.1.3 Immunoassays (Level III)A commercial dot blot detection kit is availablefor TSV from DiagXotics (Wilton, CT, USA).ELISA kits using a TSV MAb have also beenproduced. These can be used to screen possibleTSV carriers, but any positive resultsshould be cross-checked with another confirmatorytechnique, or by bioassay, sincevisualisation of clinical signs or the virus is notpossible with molecular screening techniques(this also applies to screening with PCR probes- C.8.3.1.5)C.8.3.1.4 In situ Hybridization (Level III)A commercial in situ hybridization detection kitis available for TSV from DiagXotics (Wilton, CT,USA). This technique is usually reserved forconfirmation of observations made using routinehistology (C.8.3.1.2), rather than as a standalonetechnique for screening.C.8.3.1.5 PCR Probes (Level III)An RT-PCR based assay uses shrimphaemolymph for screening purposes, giving theadvantage of being able to screen livebroodstock and assist selection of TSV-negativeshrimp for spawning. Positive results fromsurvivors of previous TSV outbreaks can beconsidered confirmatory, however, first timepositive results from non-susceptible speciesor non-enzootic sources should be analyzedusing another, confirmatory, technique for thesame reasons given for dot-bot hybridization(C.8.3.1.3).C.8.4 Diagnostic MethodsDetailed information on methods for diagnosisof TSV can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE 2000),at http://www.oie.int, or at selected references.C.8.4.1 PresumptiveC.8.4.1.1 Gross Observations (Level I)Penaeus vannamei post-larvae or older shrimpmay show a pale reddish discolouration, especiallyof the tail fan (Fig.C.8.4.1.1a,b) andpleiopods (hence the name “red tail” disease,applied by farmers when the disease first appearedin Ecuador). This colour change is dueto expansion of the red chromatophores withinthe cuticular epithelium. Magnification of theedges of the pleiopods or uropods may revealevidence of focal necrosis. Shrimp showingthese signs typically have soft shells, an emptygut and often die during moulting. During severeepizootics, sea birds (gulls, terns, cormorants,etc.) may be attracted to ponds containingshrimp over 1 gm in size.Although the transition stage of TS only lasts afew days, some shrimp may show signs of random,multi-focal, irregularly shaped melanizedcuticular lesions (Fig.C.8.4.1.1c,d,e). Thesecorrespond to blood cell repair activity aroundthe necrotic lesions induced by TSV infectionof the cuticular epithelium. Such shrimp may,or may not, have soft cuticles and reddiscolouration, and may be behaving and feedingnormally.C.8.4.1.2 Histopathology (Level II)Diagnosis of TS in acute stages of the diseaserequires histological (H&E stain preparations)demonstration of multi-focal areas of necrosisin the cuticular epithelium of the general bodysurface, appendages, gills (Fig.C.8.4.1.2a),hind-gut, esophagus and stomach(Fig.C.8.4.1.2b). Sub-cuticular connective tissueand striated muscle fibers basal or adjacentto affected cuticular epithelium may alsoshow signs of necrosis. Rarely, the antennalgland tubule epithelium is affected. Cuticularlesions may contain foci of cells with abnormallyeosinophilic (pink-staining) cytoplasm and pyknotic(condensed nucleoplasm) or karyorrhectic(fragmented nucleoplasm) nuclei. Remnantsof necrotic cells are often abundant withinacute phase lesions and appear as roughlyspherical bodies (1-20 µm diameter) that rangein stain uptake from eosinophilic to lightly basophilic(blue-staining). Another feature of acuteTS is the absence of haemocyte infiltration, orother signs of a host defense response. Thesefeatures combine to give acute phase TS lesionsa “peppered” appearance(Fig.C.8.4.1.2c), that is considered to be diagnosticfor the disease, and can be consideredconfirmatory (C.8.4.2.2) in susceptible speciesin enzootic waters. Confirmation by anothertechnique is recommended for first time observationsof these histopathological features, ortheir appearance in abnormal penaeid speciesor locations.In the transitional phase of TS, the number andseverity of the cuticular lesions that characterizeacute phase infections decrease andaffected tissues become infiltrated byhaemocytes. These may become melanized195

C.8 Taura Syndrome (TS)(C.8.4.1.1). If the acute cuticular lesions perforatethe epicuticle, the affected surfaces mayshow evidence of colonization and invasion byVibrio spp, or other secondary infections.(DV Lightner/F Jimenez)In the chronic phase of TS, the only sign of infectionis the presence of prominent lymphoidorgan spheres (LOS) (Fig.C.8.4.1.2d), whichcorrespond to aggregations of presumedhemocytes within the intertubular spaces of thelymphoid organ.c(DV Lightner)adbFig. C.8.4.1.1a,b. a. Moribund, juvenile, pondrearedPenaeus vannamei from Ecuador in theperacute phase of Taura Syndrome (TS). Shrimpare lethargic, have soft shells and a distinct redtail fan; b. Higher magnification of tail fan showingreddish discoloration and rough edges ofthe cuticular epithelium in the uropods suggestiveof focal necrosis at the epithelium of thosesites (arrows).eFig. C.8.4.1.1c,d,e. Juvenile, pond-reared P.vannamei (c – from Ecuador; d – from Texas; e– from Mexico) showing melanized foci marksites of resolving cuticular epithelium necrosisdue to TSV infection.196

C.8 Taura Syndrome (TS)(DV Lightner)(DV Lightner)Fig. C.8.4.1.2a. Focal TSV lesions in the gills(arrow). Nuclear pykinosis and karyorrhexis, increasedcytoplasmic eosinophilia, and an abundanceof variably staining generally sphericalcytoplasmic inclusions are distinguishing characteristicsof the lesions. 900x magnification.(DV Lightner)Fig. C.8.4.1.2c. Higher magnification of Fig.C.8.4.1.2b showing the cytoplasmic inclusionswith pyknotic and karyorrhectic nuclei giving a‘peppered’ appearance. Mayer-Bennett H&E.900x magnification.(DV Lightner)Fig. C.8.4.1.2b. Histological section throughstomach of juvenile P. vannamei showing prominentareas of necrosis in the cuticular epithelium(large arrow). Adjacent to focal lesions arenormal appearing epithelial cells (small arrows).Mayer-Bennett H&E. 300x magnification.Fig. C.8.4.1.2d. Mid-sagittal section of the lymphoidorgan (LO) of an experimentally infectedjuvenile P. vannamei. Interspersed among normalappearing lymphoid organ (LO) cords or tissue,which is characterized by multiple layersof sheath cells around a central hemolymphvessel (small arrow), are accumulations of disorganizedLO cells that form LO ‘spheroids”.Lymphoid organs spheres (LOS) lack a centralvessel and consists of cells which showkaryomegaly and large prominent cytoplasmicvacuoles and other cytoplasmic inclusions(large arrow). Mayer-Bennett H&E. 300x magnification.197

C.8 Taura Syndrome (TS)C.8.4.2ConfirmatoryC.8.4.2.4 Dot Blot (Level III)C.8.4.2.1 Bioassay (Levels I/II)Specific Pathogen Free (SPF) juvenile Penaeusvannamei can be used to test suspect TSV-infectedshrimp. Three exposure methods can beused:i) Suspect shrimp can be chopped up and fedto SPF juvenile P. vannamei held in smalltanks. Another tank should hold SPF shrimpfrom the same source, but fed regular feedonly (controls). If the suspect shrimp werepositive for TSV, gross signs and histopathologicallesions should become evident within3-4 days of initial exposure. Significant mortalitiesusually occur by 3-8 days post-exposure.The control shrimp should stayhealthy and show no gross or histologicalsigns of TS.ii) Whole shrimp collected from a presumptiveTSV epizootic can be homogenized for inoculationchallenge. Alternatively, heads maybe used where presumptive TS signs appearto be at the transitional phase of development(melanized lesions) or where there areno clinical signs of infection (presumptivechronic phase) since this contains the lymphoidorgan.iii) Haemolymph samples may be taken frombroodstock and used to expose SPF indicatorshrimp, as for method ii) above.C.8.4.2.2 Histopathology (Level II)Observation of the lesions described underC.8.4.1.2 can be considered confirmatory forsusceptible species from sources known to beenzootic for TSV.C.8.4.2.3 Transmission Electron Microscopy(TEM) (Level III)Transmission electron microscopy of acutephase epithelial lesions or lymphoid organspheroids that demonstrate the presence ofnon-enveloped icosahedral viral particles, 31-32 nm in diameter, in the cytoplasm of affectedcells, can be considered confirmatory whereconsistent with gross and histological clinicalsigns in a susceptible penaeid species. Furtherconfirmation using molecular techniques(C.8.4.2.4-6) are recommended, however, forfirst-time diagnoses or detection in speciesother than those listed as being naturally orexperimentally susceptible.As described under C.8.3.1.3.C.8.4.2.5 In situ Hybridization (Level III)As described under C.8.3.1.4.C.8.4.2.6 PCR Probes (Level III)As described under C.8.3.1.5.C.8.5 Modes of TransmissionShrimps that have survived the acute and transitionalphases of TS can maintain chronic subclinicalinfections within the lymphoid organ, forthe remainder of their lives. These shrimp maytransmit the virus horizontally to other susceptibleshrimp. Vertical transmission is suspected,but this has yet to be conclusively demonstrated.In addition to movement of sub-clinical carriersof TSV, aquatic insects and sea birds havebeen implicated in transmission of the disease.The water boatman, Trichocorixa reticulata(Corixidae), feeds on dead shrimp and is believedto spread TSV by flying from pond topond. Laughing gull, Larus atricilla, faeces collectedfrom around TSV-infected ponds in Texasduring the 1995 epizootic, were also found tocontain viable TSV. Viable TSV has also beenfound in frozen shrimp products.C.8.6 Control MeasuresIn much of Central America where TS is enzootic,shrimp farm management has shiftedtowards increased use of wild caught P.vannamei PL, rather than hatchery-reared PL.This has improved survival to harvest. It is suspectedthat wild PL may have increased toleranceof TS due to natural exposure and selectionof survivors. Another management strategyhas been doubling post-larval stockingdensities in semi-intensive pond culture. Heavylosses due to TS early in the production cycleare compensated for by the survivors (5-40%of the original number stocked) being TS tolerant.Selective breeding is showing promise fordevelopment of TSV resistant stocks of P.vannamei and P. stylirostris (which are resistantto both IHHNV and TSV). Initial results show a20-40% improvement in survival.Eradication depends on total removal of infectedstocks, disinfection of the culture facility,avoidance of re-introduction of the virus198

C.8 Taura Syndrome (TS)(from nearby culture facilities, wild shrimp, orsub-clinical carriers etc.), and re-stocking withTSV-free PL produced from TSV-freebroodstock.C.8.7 Selected ReferencesAragon-Noriega, E.A., J.H. Cordova-Murueta,and H.L. Trias-Hernandez. 1998. Effect ofTaura-like viral disease on survival of thewestern white shrimp (Penaeus vannamei)cultured at two densities in NorthwesternMexico. World Aquac. 29(3):66-72.Bonami, J.R., K.W. Hasson, J. Mari, B.T. Poulos,and D.V. Lightner. 1997. Taura syndrome ofmarine penaeid shrimp: Characterisation ofthe viral agent. J. Gen. Virol. 78(2):313-319.Brock, J.A., R. Gose, D.V. Lightner, and K.W.Hasson . 1995. An overview of Taura Syndrome,an important disease of farmedPenaeus vannamei, pp. 84-94. In: Swimmingthrough troubled water. Proceedings of theSpecial Session on Shrimp Farming, WorldAquaculture Society, Baton Rouge, LA.Dixon, H. and J. Dorado. 1997. Managing Taurasyndrome virus in Belize: A case study.Aquac. Mag. 23(2): 30-42.Garza, J.R., K.W. Hasson, B.T. Poulos, R.M.Redman, B.L. White, and D.V. Lightner. 1997.Demonstration of infectious Taura syndromevirus in the feces of seagulls collected duringan epizootic in Texas. J. Aquat. Anim.Health 9(2):156-159.Hasson, K.W., D.V. Lightner, B.T. Poulos, R.M.Redman, B.L. White, J.A. Brock, and J.R.Bonami. 1995. Taura syndrome in Penaeusvannamei: Demonstration of a viral etiology.Dis. Aquat. Org. 23(2):115-126.Hasson, K.W., J. Hasson, H. Aubert, R.M.Redman, and D.V. Lightner. 1997. A new RNA-friendly fixative for the preservation ofpenaeid shrimp samples for virological detectionusing cDNA genomic probes. J. Virol.Meth. 66:227-236.Hasson, K.W., D.V. Lightner, J. Mari, J.R.Bonami, B.T. Poulos, L.L. Mohney, R.M.Redman, and J.A Brock. 1999a. The geographicdistribution of Taura Syndrome Virus(TSV) in the Americas: determination byhistopathology and in situ hybridisation usingTSV-specific cDNA probes. Aquac. 171(1-2):13-26.Hasson, K.W., Lightner, D.V., Mohney, L.L.,Redman, R.M., Poulos, B.T. and B.M. White.1999b. Taura syndrome virus (TSV) lesion developmentand the disease cycle in the Pacificwhite shrimp Penaeus vannamei. Dis.Aquat. Org. 36(2):81-93.Hasson, K.W., D.V. Lightner, L.L. Mohney, R.M.Redman, and B.M. White. 1999c. Role oflymphoid organ spheroids in chronic Taurasyndrome virus (TSV) infections in Penaeusvannamei. Dis. Aquat. Org. 38(2):93-105.Jimenez, R., R. Barniol, L. Barniol and M.Machuca. 2000. Periodic occurrence of epithelialviral necrosis outbreaks in Penaeusvannamei in Ecuador. Dis. Aquat.Org.42(2):91-99.Lightner, D.V. 1996. A Handbook of ShrimpPathology and Diagnostic Procedures forDiseases of Cultured Penaeid Shrimp. WorldAquaculture Society, Baton Rouge, LA. 304p.Lightner, D.V. 1999. The penaeid shrimp virusesTSV, IHHNV, WSSV and YHV: Current Statusin the Americas, available diagnostic methodsand management strategies. J. AppliedAquac. 9(2):27-52.Lightner, D.V. and R.M. Redman. 1998. Strategiesfor the control of viral diseases of shrimpin the Americas. Fish Pathol. 33:165-180.Lightner, D.V., R.M. Redman, K.W. Hasson, andC.R. Pantoja. 1995. Taura syndrome inPenaeus vannamei (Crustacea: Decapoda):Gross signs, histopathology and ultrastructure.Dis. Aquat. Org. 21(1):53-59.Lotz, J.M. 1997a. Effect of host size on virulenceof Taura virus to the marine shrimpPenaeus vannamei (Crustacea: Penaeidae).Dis. Aquat. Org. 30(1):45-51.Lotz, J.M. 1997b. Disease control and pathogenstatus assurance in an SPF-basedshrimp aquaculture industry, with particularreference to the United States, pp. 243-254.In: Diseases in Asian Aquaculture III. Flegel,T.W. and I.H. MacRae (eds.). Fish Health Section,Asian Fisheries Society, Manila, The Philippines.Morales-Covarrubias, M.S. and C. Chavez-Sanchez. 1999. Histopathological studies onwild broodstock of white shrimp Penaeusvannamei in the Platanitos Area, adjacent toSan Blas, Nayarit, Mexico. J. World Aquac..199

C.8 Taura Syndrome (TS)Soc. 30(2):192-200.Nunan, L.M., B.T. Poulos, and D.V. Lightner.1998. Reverse transcriptase polymerasechain reaction (RT-PCR) used for the detectionof Taura Syndrome virus (TSV) in experimentallyinfected shrimp. Dis. Aquat. Org.34(2):87-91.OIE. 2000. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.Overstreet, R.M., D.V. Lightner, K.W. Hasson,S. McIlwain, and J.M. Lotz. 1997. Susceptibilityto Taura syndrome virus of somepenaeid shrimp species native to the Gulf ofMexico and the southeastern United States.J. Invert. Pathol. 69(2):165-176.Poulos, B.T., R. Kibler, D. Bradley-Dunlop, L.L.Mohney, and D.V. Lightner. 1999. Productionand use of antibodies for the detection of theTaura syndrome virus in penaeid shrimp. Dis.Aquat. Org. 37(2):99-106.Tu, C., H.-T. Huang, S.-H. Chuang, J.-P. Hsu,S.-T. Kuo, N.-J. Li, T.-L. Hsu, M.-C. Li, andS.-Y. Lin. 1999. Taura syndrome in Pacificwhite shrimp Penaeus vannamei cultured inTaiwan. Dis. Aquat. Org. 38(2):159-161.Yu, C.-I. and Y.-L. Song. 2000. Outbreaks ofTaura syndrome in Pacific white shrimpPenaeus vannamei cultured in Taiwan. FishPathol. 35(1):21-24.Zarain-Herzberg, M. and F. Ascencio-Valle.2001. Taura syndrome in Mexico: follow-upstudy in shrimp farms of Sinaloa. Aquac.193(1-2):1-9.200

C.9 NUCLEAR POLYHEDROSISBACULOVIROSES(BACULOVIRUS PENAEI [BP] PvSNPV; MONODON BACULOVIRUS [MBV]PmSNPV)C.9.1 Background InformationC.9.1.1 Causative AgentNuclear Polyhedrosis Baculoviroses (NPB) infectionsare caused by the Baculoviridae,Baculovirus penaei (BP - PvSNPV) and Mondonbaculovirus (MBV – PmSNPV). The diseasesassociated with these viruses are Baculovirusdisease, Nuclear polyhedrosis disease, polyhedralinclusion body virus disease (PIB), polyhedralocclusion body virus disease (POB) andBaculovirus penaei (BP) virus disease. Moredetailed information about the disease can befound at OIE Diagnostic Manual for Aquatic AnimalDiseases (OIE 2000).C.9.1.2 Host RangeBP infects in a wide range of penaeid shrimp includingPenaeus duorarum, P. aztecus, P.setiferus, P. vannamei, P. stylirostris and P.marginatus. BP has also been reported from P.penicillatus, P. schmitti, P. paulensis and P. subtilis.MBV-type baculoviruses are, by definition, primarilyfound in cultured P. monodon. Other coculturedspecies may also acquire MBV-type virusinfections, but these have not been associatedwith severe pathology, or developed nonmonodonreservoirs.C.9.1.3Geographical DistributionBP is found throughout the Americas from theGulf of Mexico to Central Brazil on the East Coastand from Peru to Mexico on the Pacific Coast.BP has also been found in wild shrimp in Hawaii.Multiple strains of BP are recorded within thisgeographic range.MBV has been reported from Australia, East Africa,the Middle East, many Indo-Pacific countriesand from south and eastern Asia. MBV-typeviruses have also been found in sites associatedwith P. monodon culture in the Mediterranean andWest Africa, Tahiti and Hawaii, as well as severallocations in North and South America andthe Caribbean.C.9.2 Clinical AspectsThe impact of BP varies from species to species.Penaeus aztecus and P. vannamei arehighly susceptible. Penaeus stylirostris is moderatelysusceptible and P. monodon and P.setiferus appear to be resistant/tolerant. In susceptiblespecies, BP infection is characterisedby a sudden onset of high morbidity and mortalityin larval and post larval stages. Growthrates decrease, the shrimp stop feeding, appearlethargic and show signs of epibiont fouling(due to reduced grooming activity). The virusattacks the nuclei of hepatopancreas epitheliabut can also infect mid-gut epithelia. Althoughinfections may be chronic to acute, withhigh cumulative mortality, presence of the BPvirus is not always associated with disease andpost-larvae older than 63 days show no clinicalsigns of infection (see C.9.6).MBV causes similar clinical signs to BP, due tosimilar infection of the hepatopancreatic andmid-gut epithelial nuclei. Infections of MBV mayalso occur in the lymphoid organ. Larval stagesof P. monodon are particularly susceptible, however,prevalences of >45% may be present injuvenile and adult developmental stages withno overt clinical effects.C.9.3 Screening MethodsMore detailed information on methods forscreening NPB can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000), at http://www.oie.int, or at selected references.C.9.3.1 PresumptiveThere are no presumptive screening methodsfor asymptomatic carriers of BP and MBV, sincedirect microscopic methods (C.9.3.2) demonstratingthe characteristic occlusion bodies (tetrahedralfor BP and spherical-ovoid for MBV)are considered to be confirmatory.C.9.3.2ConfirmatoryC.9.3.2.1 Wet Mount of Fresh Tissue(Level I/II)BP infections can be confirmed by bright-fieldor phase contrast microscopic observation ofsingle or multiple tetrahedral (polyhedral) inclusion(occlusion) bodies (Fig. C.9.3.2.1a) withinenlarged nuclei of hepatopancreas or midgutepithelia. These bodies can range in size from0.1-20.0 µm (modal range = 8-10 µm) along theperpendicular axis from the base of thepyrimidal shape to the opposite point.MBV infections observed using the same microscopeapparatus appear as single or multiplespherical or sub-spherical inclusion bodieswithin enlarged nuclei of hepatopancreas201

C.9 Nuclear PolyhedrosisBaculoviroses(Baculovirus penaei [BP] PvSNPV; Monodon Baculovirus [MBV]PmSNPV)(DV Lightner)(DV Lightner)bacFig. C.9.3.2.1b,c. Mid and high magnificationviews of tissue squash preparations of thehepatopancreas (HP) from PL of P. monodonwith MBV infections. Most HP cells in both PLsusually display multiple, generally spherical, intranuclearocclusion bodies (arrow) that are diagnosticfor MBV. 0.1% malachite green. 700x(b) and 1 700x (c) magnifications.(DV Lightner)bFig. C.9.3.2.3a,b. a. Mid-magnification view ofmid-sagittal sections of PL of P. vannamei withsevere BP infections of the hepatopancreasshowing multiple eosinophilic BP tetrahedralocclusion bodies within markedly hypertrophiedhepatopnacreas (HP) cell nuclei (arrows).Mayer-Bennett H&E. 700x magnification; b.High magnification of an HP tubule showingseveral BP-infected cells that illustrate well theintranuclear, eosinophilic, tetrahedral occlusionbodies of BP (arrows). Mayer-Bennett H&E.1800x magnification.>Fig. C.9.3.2.1a. Wet mount of feces from a P.vannamei infected with BP showing tetrahedralocclusion bodies (arrows) which are diagnosticfor infection of shrimp’s hepatopancreas ormidgut epithelial cells. Phase contrast, no stain.700x magnification.202

C.9 Nuclear PolyhedrosisBaculoviroses(Baculovirus penaei [BP] PvSNPV; Monodon Baculovirus [MBV]PmSNPV)or midgut epithelia. MBV occlusion bodiesmeasure 0.1 –20.0 µm in diameter (Fig.C.9.3.2.1b,c). The occlusion bodies can bestained using a 0.05% aqueous solution ofmalachite green, which stains them moredensely than surrounding, similarly sized sphericalbodies (cell nuclei, secretory granules, lipiddroplets, etc.).C.9.3.2.2 Faecal Examination (Level I/II)Make wet mounts of faecal strands and examinefor occlusion bodies, as described for freshtissue mounts (C.9.3.2.1).C.9.3.2.3 Histopathology (Level II)Tissues from live or moribund (but not dead,due to rapid liquefaction of the target organ –the hepatopancreas) shrimp should be fixed inDavidson’s fixative to ensure optimum fixationof the hepatopancreas (10% buffered formalinprovides sub-optimal hepatopancreas preservation).The fixative should be administered bydirect injection into the hepatopancreas. Thecuticle should be cut along the dorsal line ofthe cephalothorax to enhance fixative penetrationof the underlying tissues and the tissuesshould be fixed for 24-48 hr before transfer to70% ethanol for storage. The tissues can thenbe processed for routine paraffin embedding,sectioning at 5-7 µm thickness and staining withHarris’ haematoxylin and eosin or other Giemsaor Gram tissue-staining methods. Brown andBrenn’s histological Gram stain provides intensered or purple colouration of both MBV (see alsoFig. C.5.4.2.1d – C.5) and BP occlusion bodies(Fig. C.9.3.2.3a,b), aiding in their differentiationfrom surrounding tissues.C.9.3.2.4 Polymerase Chain ReactionAssays (Level III)Two primers sequences are available for theMBV polyhedrin gene (Lu et al 1993) and anotherpair are available for a 1017bp fragmentof the viral genome (Mari et al 1993). Details onthe PCR procedures for screening tissue or faecalsamples are provided in the OIE DiagnosticManual (OIE 2000) or selected references(C.9.7).C.9.4 Diagnostic MethodsMore detailed information on methods for diagnosisof NPB can be found in the OIE DiagnosticManual for Aquatic Animal Diseases (OIE2000), at http://www.oie.int, or at selected references.C.9.4.1PresumptiveC.9.4.1.1 Gross Observations (Level I)Gross signs of BP vary between susceptiblespecies but include decreased growth, cessationof feeding and preening, lethargy and increasedepibiont fouling. Some shrimp mayexhibit a white mid-gut line through the ventralabdominal cuticle. None of these symptoms arespecific to BP, but can be considered suspectin susceptible species and at early developmental/post-larvalstages which have a history ofbeing affected by BP.MBV causes similar clinical signs to BP, butprincipally affects larval of P. monodon with aninverse correlation between larval age andpathogenic effects. Adults can be infected withno overt signs (see C.9.3). As with BP, thesesigns are not specific to MBV.C.9.4.2ConfirmatoryC.9.4.2.1 Wet Mount of Fresh Tissue(Level I/II)As described for C.9.3.2.1.C.9.4.2.2 Faecal Examination (Level I/II)As described for C.9.3.2.2.C.9.4.2.3 Histopathology (Level II)As described for C.9.3.2.3.C.9.4.2.4 Autofluorescence with phloxinestain (Level II)An aqueous solution of 0.001% phloxine usedon tissue squash preparations or faeces, willcause occlusion bodies of both BP and MBVto fluoresce yellow-green when examined usinga fluorescent microscope (barrier filter 0-515 nm and exciter filter of 490 nm) (Thurmanet al. 1990). The same effect is achieved using0.005% phloxine in routine haematoxylin andeosin stain of histological tissue preparations.C.9.4.2.5 Transmission Electron Microscopy(TEM) (Level III)BP virions are rod-shaped with an envelopednucleocapsid measuring 286-337 nm x 56-79nm. The virions are found either free or occludedwithin a crystalline protein matrix (the occlusionbody). In early infections, virions are found203

C.9 Nuclear PolyhedrosisBaculoviroses(Baculovirus penaei [BP] PvSNPV; Monodon Baculovirus [MBV]PmSNPV)in association with nuclear enlargements, aberrantstromatic patterns of the nucleoplasm,degenerate nucleoli, and nuclear membraneproliferation into labyrinths. Occlusion bodiesoccur during later stages of infection.MBV has been shown to have two types ofocclusion bodies using electron microscopicexaminations (Ramasamy et al. 2000). Type 1has a paracrystalline array of polyhedrin unitswithin a lattice work spacing of 5-7 nm, whichcontains occluded virions (along with a fewperipheral non-occluded virions) that have adouble envelope and measure 267 ± 2 x 78 ± 3nm. Type 2 occlusion bodies consist of noncrystalline,granulin-like sub-units 12 nm in diameter,containing mostly non-occluded virionsmeasuring 326 ± 4 x 73 ± 1 nm. In addition, anon-enveloped stage has recently been detected(Vickers et al. 2000) in the cytoplasm ofinfected cells and close association with thenuclear membrane.C.9.4.2.6 In situ Hybridization (Level III)Details of the preparation and analytical proceduresrequried for in situ hybridisation for confirmingBP and MBV infections are provided inthe OIE Diagnostic Manual (OIE 2000a) underboth the Nuclear Polyhedrosis Baculoviroseschapter (Chapter 4.2.2) as well as the InfectiousHypodermal and Haematopoietic Necrosischapter (Chapter 4.2.3).C.9.5 Modes of TransmissionBP and MBV are both transmitted orally viauptake of virus shed with the faeces of infectedshrimp (C.9.3.2.2), or cannibalism on dead anddying shrimp. Infected adults have also beenshown to infect their offspring via faecal contaminationof the spawned egg masses.C.9.6 Control MeasuresOvercrowding, chemical and environmentallyinduced stress, have all been shown to increasethe virulence of MBV and BP infections in susceptibleshrimp species under culture conditions.Exposure of stocks to infection can be avoidedby pre-screening the faeces of potentialbroodstock and selecting adults shown to befree of faecal contamination by occlusion bodiesof either baculovirus. Prevention of infectionsmay also be achieved by surface disinfectionof nauplii larvae or fertilised eggs withformalin, iodophore and filtered clean seawateras follows:• Collect nauplii and wash in gently running seawater for 1-2 minutes.• Immerse the nauplii in a 400 ppm solution offormalin for 1 minute followed by a solutionof 0.1 ppm iodine for an additional minute.The same procedure can be used on fertilisedeggs except the formalin concentration is reducedto 100ppm.• Rinse the treated nauplii in running sea waterfor 3-5 min and introduce to the hatchery.Eradication of clinical outbreaks of BP and MBVmay be possible in certain aquaculture situationsby removal and sterile disposal of infectedstocks, disinfection of the culture facility, theavoidance of re-introduction of the virus (fromother nearby culture facilities, wild shrimp, etc.).C.9.7 Selected ReferencesAlcivar-Warren,A., R.M. Overstreet, A.K. Dhar,K. Astrofsky, W.H. Carr, J. Sweeny and J.M.Lotz. 1997. Genetic susceptibility of culturedshrimp (Penaeus vannamei) to infectioushypodermal and hematopoietic necrosis virusand Baculovirus penaei: Possible relationshipwith growth status and metabolic geneexpression. J. Invertebr. Pathol. 70(3): 190-197.Belcher, C.R. and P.R. Young. 1998.Colourimetric PCR-based detection ofmonodon baculovirus in whole Penaeusmonodon postlarvae. J. Virol. Methods 74(1):21-29.Brock, J.A., D.V. Lightner and T.A. Bell. 1983. Areview of four virus (BP, MBV, BMN, andIHHNV) diseases of penaeid shrimp with particularreference to clinical significance, diagnosisand control in shrimp aquaculture.Proc. 71st Intl. Council for the Exploration ofthe Sea, C.M. 1983/Gen: 10/1-18.Brock, J.A., L.K. Nakagawa, H. Van Campen,T. Hayashi, S. Teruya. 1986. A record ofBaculovirus penaei from Penaeus marginatusRandall in Hawaii. J. Fish Dis. 9: 353-355.Bruce, L.D., B.B. Trumper, and D.V. Lightner.1991. Methods of viral isolation and DNAextraction for a penaeid shrimp baculovirus.J. Virol. Meth. 34:245-254.Bruce, L.D., R.M. Redman and D.V. Lightner.1994. Application of gene probes to determinetarget organs of a penaeid shrimp204

C.9 Nuclear PolyhedrosisBaculoviroses(Baculovirus penaei [BP] PvSNPV; Monodon Baculovirus [MBV]PmSNPV)baculovirus using in situ hybridisation.Aquaculture 120(1-2): 45-51.Bruce, L.D., D.V. Lightner, R.M. Redman andK.C. Stuck. 1994. Comparison of traditionaland molecular tools for Baculovirus penaeiinfections in larval Penaeus vannamei. J.Aquatic Anim. Health 6(4): 355-359.Bueno, S.L., R.M. Nascimento and I.Nascimento. 1990. Baculovirus penaei infectionin Penaeus subtilis: A new host and anew geographic range of the disease. J.World Aquacult. Soc. 21(3): 235-237.Chen, S.N., P.S. Chang and G.S. Kou. 1993.Diseases and treatment strategies onPenaeus monodon in Taiwan. pp. 43-57 In:Proceedings of the Symposium on Aquacultureheld in Beijing, 21-23 December 1992,Taiwan Fisheries Research Institute, Keelung,TRFI Conf. Proc. #3.Chen, S.N., P.S. Chang, C.C. Chen and G.H.Kou. 1993. Studies on infection pathway ofMonodon Baculovirus (MBV). COA Fish. Ser.40: 81-85.Chen, X., D. Wu, H. Huang, X. Chi and P. Chen.1995. Ultrastructure on Penaeus monodonbaculovirus. J. Fish. China (Shuichan Xuebao)19(3): 203-209.Fegan, D.F., T.W. Flegel, S. Sriurairatana andM. Waiyakruttha. 1991. The occurrence, developmentand histopathology of monodonbaculovirus in Penaeus monodon in Thailand.Aquac. 96(3-4): 205-217.Flegel, T.W., V. Thamavit, T. Pasharawipas andV. Alday-Sanz. 1999. Statistical correlationbetween severity of hepatopancreaticparvovirus infection and stunting of farmedblack tiger shrimp (Penaeus monodon).Aquac.174(3-4): 197-206.Hammer, H.S., K.C. Stuck and R.M. Overstreet.1998. Infectivity and pathogenicity ofBaculovirus penaei (BP) in cultured larval andpostlarval Pacific white shrimp, Penaeusvannamei, related to the stage of viral development.J. Invertebr. Pathol. 72(1): 38-43.Hao, N.V., D.T. Thuy, L.T. Loan, L.T.T. Phi, L.H.Phuoc, H.H.T. Corsin and P. Chanratchakool.Presence of the two viral pathogens WSSVand MBV in three wild shrimp species(Penaeus indicus, Metapenaeus ensis andMetapenaeus lysianassa). Asian Fish. Sci.12(4): 309-325.Hsu, Y.L., K.H. Wang, Y.H. Yang, M.C. Tung,C.H. Hu, C.F. Lo, C.H. Wang and T. Hsu. 2000.Diagnosis of Penaeus monodon-typebaculovirus by PCR and by ELISA. Dis.Aquatic Org. 40(2): 93-99.Karunasagar, I., S.K. Otta and I. Karunasagar.1998. Monodon baculovirus (MBV) and bacterialsepticaemia associated with massmortality of cultivated shrimp (Penaeusmonodon) from the east coast of India. IndianJ. Virol. 14(1): 27-30.LeBlanc, B.D. and R.M. Overstreet. 1990.Prevalence of Baculovirus penaei in experimentallyinfected white shrimp (Penaeusvannamei) relative to age. Aquac. 87(3-4):237-242.LeBlanc, B.D. and R.M. Overstreet. 1991. Effectof dessication, pH, heat and ultravioletirradiation on viability of Baculovirus penaei.J. Invertebr. Pathol. 57(2): 277-286.LeBlanc, B.D. and R.M. Overstreet. 1991. Efficacyof calcium hypochlorite as a disinfectantagainst the shrimp virus Baculoviruspenaei. J. Aquatic Anim. Health 3(2): 141-145.LeBlanc, B.D., R.M. Overstreet and J.M. Lotz.1991. Relative susceptibility of Penaeusaztecus to Baculovirus penaei. J. WorldAquacult. Soc. 22(3): 173-177.Lightner, D.V. 1996. A Handbook of ShrimpPathology and Diagnostic Procedures forDisease of Cultured Penaeid Shrimp. WorldAquaculture Society, Baton Rouge, LA. 304p.Lightner, D.V. and R.M. Redman. 1989.Baculovirus penaei in Penaeus stylirostris(Crustacea: Decapoda) cultured in Mexico:Unique cytopathology and a new geographicrecord. J. Invertebr. Pathol. 53(1): 137-139.Lightner, D.V., R.M. Redman, and E.A. AlmadaRuiz. 1989. Baculovirus penaei in Penaeusstylirostris (Crustacea: Decapoda) cultured inMexico: unique cytopathology and a newgeographic record. J. Inverteb. Pathol.53:137-139.Lu, C.C., K.F.J. Tang, G.H. Kou and S.N. Chen.1995. Detection of Penaeus monodon-typebaculovirus (MBV) infection in Penaeusmonodon Fabricius by in situ hybridisation.J. Fish Dis. 18(4): 337-345.205

C.9 Nuclear PolyhedrosisBaculoviroses(Baculovirus penaei [BP] PvSNPV; Monodon Baculovirus [MBV]PmSNPV)Lu, C.C., K.F.J. Tang and S.N. Chen. 1996.Morphogenesis of the membrane labyrinthin penaeid shrimp cells infected with Penaeusmonodon-baculovirus (MBV). J. Fish Dis.19(5): 357-364.OIE. 2000. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.Poulos, B.T., J. Mari, J-R. Bonami, R. Redmanand D.V. Lightner. 1994. Use of non -radioactivelylabeled DNA probes for the detectionof a baculovirus from Penaeus monodonby in situ hybridisation on fixed tissues. J.Virol. Methods 49(2): 187-194.Ramasamy, P., P.R. Rajan, V. Purushothamanand G.P. Brennan. 2000. Ultrastructure andpathogenesis of Monodon baculovirus (PmSNPV) in cultuerd larvae and natural broodersof Penaeus monodon. Aquac.184(1-2):45-66.baculoviruses and their use in diagnosis ofinfections. J. Aquat. Anim. Health 2(2): 128-131.Vickers, J.E., J.L. Paynter, P.B. Spradbrow andR.J.G. Lester. 1993. An impression smearmethod for rapid detection of Penaeusmonodon-type baculovirus (MBV) in Australianprawns. J. Fish Dis. 16(5): 507-511.Vickers, J.E., R. Webb and P.R. Young. 2000.Monodon baculovirus from Australia: ultrastructuralobservations. Dis. Aquat. Org.39(3): 169-176.Wang, S.Y., C. Hong and J.M. Lotz. 1996. Developmentof a PCR procedure for the detectionof Baculovirus penaei in shrimp. Dis.Aquat. Org. 25(1-2): 123-131.Shariff, M., R.P. Subasinghe and J.R. Arthur(eds) (1992) Diseases in Asian Aqaculture.Proceedings of the First Symposium on Diseasesin Asian Aquaculture, Bali 1990. FishHealth Section, Asian Fisheries Society, Manila,Philippines, 585pp.Spann, K.M., R.J.G. Lester and J.L. Paynter.1993. Efficiency of chlorine as a disinfectantagainst Monodon baculovirus. Asian Fish.Sci. 6(3): 295-301.Stuck, K.C. and R.M. Overstreet. 1994. Effectof Baculovirus penaei on growth and survivalof experimentally infected postlarvae of thePacific white shrimp, Penaeus vannamei. J.Invertebr. Pathol. 64(1): 18-25.Stuck, K.C. and S.Y. Wang. 1996. Establishmentand persistence of Baculovirus penaei infectionsin cultured Pacific white shrimpPenaeus vannamei. J. Invertebr. Pathol. 68(1):59-64.Stuck, K.C., L.M. Stuck, R.M. Overstreet andS.Y. Wang. 1996. Relationship between BP(Baculovirus penaei) and energy reserves inlarval and postlarval Pacific white shrimpPenaeus vannamei. Dis. Aquat. Org. 24(3):191-198.Thurman, R.B., T.A. Bell, D.V. Lightner and S.Hazanow. 1990. Unique physicochemicalproperties of the occluded penaeid shrimp206

BACTERIAL DISEASE OF SHRIMPC.10 NECROTISINGHEPATOPANCREATITIS (NHP)C.10.1 Background InformationC.10.3Screening MethodsC.10.1.1 Causative AgentNecrotising Hepatopancreatitis (NHP) is causedby a bacterium that is relatively small, highly pleomorphic,Gram negative, and an apparent obligateintracellular pathogen. The NHP bacteriumhas two morphologically different forms: one is asmall pleomorphic rod and lacks flagella; whilethe other is a longer helical rod possessing eightflagella on the basal apex of the bacterrium, andan additional flagellum (or possibly two) on thecrest of the helix.. The NHP bacterium occupiesa new genus in the alpha Proteobacteria, and isclosely related to other bacterial endosymbiontsof protozoans. NHP is also known as Texasnecrotizing hepatopancreatitis (TNHP), TexasPond Mortality Syndrome (TPMS) and Peru necrotizinghepatopancreatitis (PNHP). More informationabout the disease is found in Lightner(1996).C.10.1.2 Host RangeNHP can infect both Penaeus vannamei and P.stylirostris but causes higher mortalities in theformer species. NHP has also been reported inP. aztecus, P. californiensis and P. setiferus.C.10.1.3 Geographic DistributionNHP was first described in Texas in 1985. Otheroutbreaks have been reported in most LatinAmerican countries on both the Pacific and AtlanticOcean coasts, including Brazil, Costa Rica,Ecuador, Mexico, Panama, Peru and Venezuela.C.10.2 Clinical AspectsThe NHP bacterium apparently infects only theepithelial cells lining the hepatopancreatic tubules,and, to date, no other cell type has beenshown to become infected. The hepatopancreasin shrimp is a critical organ involved in food digestion,nutrient absorption and storage, and anyinfection has obvious and serious consequencesfor the affected animal, from reduced growth todeath. Various environmental factors appear tobe important for the onset of NHP clinical signs;the most prominent ones are water salinity over16 ppt (parts per thousand) and water temperatureof 26˚C or higher.C.10.3.1 ConfirmatoryC.10.3.1.1 Dot Blot for AsymptomaticAnimals (Level III)A commercial dot blot detection kit is availablefor NHP from DiagXotics (Wilton, CT, USA).C.10.3.1.2 In situ Hybridization (Level III)A commercial in situ hybridization detection kitis available for NHP from DiagXotics (Wilton,CT, USA).C.10.3.1.3 Polymerase Chain Reaction(PCR) (Level III)Samples of hepatopancreas are fixed in 70%ethanol and triturated prior to processing. DNAis isolated as follows: 25 mg of the trituratedhepatopancreas is suspended in 250 µl of digestionbuffer (50 mM Tris, 20 mM EDTA, 0.5%SDS, pH 8.5) in 0.5 ml eppendorf tubes. ProteinaseK (7.5 µl of a 20 mg ml -1 stock solution)is added and the tube incubated at 60°C for2 h with periodic vortexing. The tube is thenincubated at 95°C for 10 min to inactivate theproteinase K. The tube is then centrifuged for 3min at 13,000 rpm (16,000 x g) and 75 µl of thesupernatant applied to a CHROMA SPIN TE-100 (Clontech Labs) column and centrifuged ina horizontal rotor according to themanufacturer’s protocol. The solution collectedby centrifugation is diluted 1:100 and 1:1000 indistilled water prior to use in the PCR assay.Below is the sequence of oligonucleotide primersused for amplifying variable regions of the16S rRNA sequence:Forward: 5'-ACG TTG GAG GTT CGT CCTTCA G-3'Reverse1 5'-TCA CCC CCT TGC TTC TCATTG T-3'Reverse2 5'-CCA GTC ATC ACC TTT TCTGTG GTC-3'The forward primer and reverse primer 1 amplifya 441 bp fragment, the forward primer andreverse primer 2 amplify a 660 bp fragent. PCRis performed in 50 µl reactions containing 10mM Tris-HCl (pH 8.3), 50mM KCl, 1.5 mM MgCl,200 mM deoxynucleotides, 0.5 m?M of the forwardand the paired reverse primers and 0.03to 0.3 µg of template DNA. The reactants areheated to 94°C in a programmable thermocycler207

C.10 Necrotising Hepatopancreatitis(NHP)(DV Lightner)(DV Lightner)Fig. C.10.4.1.1. Juvenile P. vannamei with NHPshowing markedly atrophied hepatopancreas,reduced to about 50% of its normal volume.a(DV Lightner)bFig. C.10.4.1.2. Wet- mount of the HP of infectedshrimp with inflamed hemocyte, melanizedHP tubules and absence of lipid droplets.No stain. 150x magnification.(DV Lightner)Fig. C.10.4.1.3a,b. Low and mid-magnificationof photographs of the HP of a severely NHPinfected juvenile P. vannamei. Severehemocytic inflammation of the intratubularspaces (small arrow) in response to necrosis,cytolysis and sloughing of HP tubule epithelialcells (large arrow), are among the principal histopathologicalchanges due to NHP. Mayer-Bennett H&E. 150x (a) and 300x (b) magnifications.>Fig. C.10.4.1.3c. Low magnification view of theHP of a juvenile P.vannamei with severe, chronicNHP. The HP tubule epithelium is markedly atrophied,resulting in the formation of largeedematous (fluid filled or “watery areas in theHP. Mayer-Bannett H & E. 100x magnification.208

C.10 Necrotising Hepatopancreatitis(NHP)prior to adding 1.25 U of Amplitaq DNA polymerase.The final solution is then overlayed withmineral oil. The amplification profile consists of35 cycles of 30 s at 94°C, 30 s at 58°C and 1min at 72°C with an additional 5 min at 72°Cfollowing the final cycle. PCR products is examinedby electrophoresis in 1% agarose in TAEbuffer containing 0.5 m?g ml -1 ethidium bromide.C.10.4 Diagnostic MethodsMore detailed information on methods for diagnosisof NHP can be found in Lightner (1996)or in selected references.C.10.4.1 PresumptiveC.10.4.1.1 Gross Observations (Level 1)A wide range of gross signs can be used toindicate the possible presence of NHP. Theseinclude: lethargy, reduced food intake, higherfood conversion ratios, anorexia and emptyguts, noticeable reduced growth and poorlength weight ratios (“thin tails”); soft shells andflaccid bodies; black or darkened gills; heavysurface fouling by epicommensal organisms;bacterial shell disease, including ulcerative cuticlelesions or melanized appendage erosion;and expanded chromatophores resulting in theappearance of darkened edges in uropods andpleopods. The hepatopancreas may be atrophied(Fig.C.10.4.1.1) and have any of the followingcharacteristics: soft and watery; fluidfilled center; paled with black stripes (melanizedtubules); pale center instead of the normal tanto orange coloration. Elevated mortality ratesreaching over 90% can occur within 30 days ofonset of clinical signs if not treated.C.10.4.1.2 Wet Mounts (Level II)Wet mounts of the hepatopancreas of shrimpwith NHP may show reduced or absent lipiddroplets and/or melanized hepatopancreas tubules(Fig.C.10.4.1.2).C.10.4.1.3 Histopathology (Level II)NHP is characterised by an atrophied hepatopancreasshowing moderate to extreme atrophyof the tubule mucosa and the presence ofthe bacterial forms through histological preparations.Principal histopathological changes dueto NHP include hemocytic inflammation of theintertubular spaces in response to necrosis,cytolysis, and sloughing of hepatopancreastubule epithelial cells (Fig. C.10.4.1.3a,b). Thehepatopancreas tubule epithelium is markedlyatropied, resulting in the formation of largeedematous (fluid filled or “watery”) areas in thehepatopancreas (Fig.C.10.4.1.3c).Tubule epithelialcells within granulomatous lesions aretypically atrophied and reduced from simplecolumnar to cuboidal in morphology. They containlittle or no stored lipid vacuoles(Fig.C.10.4.1.3d) and markedly reduced or nosecretory vacuoles.C.10.4.2 ConfirmatoryC.10.4.2.1 Transmission Electron Microscopy(TEM) (Level III)Two distinct versions of the NHP bacteriumoccur in infected hepatopancreatic cells. Thefirst is a rod-shaped rickettsial-like form measuring0.3 µm x 9 µm which lacks flagella. Thesecond is a helical form (Fig.C.10.4.2.1) measuring0.2 µm x 2.6-2.9 µm which has eightperiplasmic flagella at the basal apex of thebacterium and an additional 1-2 flagella on thecrest of the helix.C.10.4.2.2 Dot Blot for AsymptomaticAnimals (Level III)A commercial dot blot detection kit is availablefor NHP from DiagXotics (Wilton, CT, USA).C.10.4.2.3 In situ Hybridization (Level III)A commercial in situ hybridization detection kitis available for NHP from DiagXotics (Wilton,CT, USA).C.10.4.2.4 Polymerase Chain Reaction(PCR) (Level III)As described for C.10.3.1.3C.10.5Modes of TransmissionEarly detection of clinical NHP is important forsuccessful treatment because of the potentialfor cannibalism to amplify and transmit the disease.Molecular testing of PL from infectedbroodstock indicates that vertical transmissiondoes not occur.C.10.6 Control MeasuresPeriodic population sampling and examination(through histopathology, TEM or commercialgene probe) are highly recommended in farms209

C.10 Necrotising Hepatopancreatitis(NHP)(DV Lightner)Fig. C.10.4.1.3d. The HP tubule epithelial cellsshow no cytoplasmic lipid droplets, but insteadcontain masses of the tiny, non-membranebound, intracytoplasmic NHP bacteria (arrow).Mayer-Bennett H&E. 1700x magnification.(DV Lightner)with a history of NHP occurrence and whereenvironmental conditions favor outbreaks. Theuse of the antibiotic oxytetracycline (OTC) inmedicated feeds is probably the best NHP treatmentcurrently available, particularly if diseasepresence is detected early.There is also some evidence that deeper productionponds (2 m) and the use of hydratedlime (Ca(OH) 2) to treat pond bottoms duringpond preparation before stocking can help reduceNHP incidence. Preventive measures caninclude raking, tilling and removing pond bottomsediments, prolonged sun drying of pondsand water distribution canals for several weeks,disinfection of fishing gear and other farmequipment using calcium hypochlorite and dryingand extensive liming of ponds.C.10.7 Selected ReferencesBrock, J.A. and K. Main. 1994. A Guide to theCommon Problems and Diseases of CulturedPenaeus vannamei. Oceanic Institute,Makapuu Point, Honolulu, Hawaii. 241p.Frelier,P.F., R.F. Sis,T.A. Bell and D.H. Lewis.1992. Microscopic and ultrastructural studiesof necrotizing hepatopancreatitis in Texascultured shrimp (Penaeus vannamei). Vet.Pathol. 29:269-277.Lightner, D.V. 1996. A Handbook of ShrimpPathology and Diagnostic Procedures forDiseases of Cultured Penaeid Shrimp. WorldAquaculture Society, Baton Rouge, LA. 304p.Fig. C.10.4.2.1. Low magnification TEM of ahepatopancreatocyte from a juvenile P.vannamei with NHP. Profiles of intracellular rodshapedforms (large arrow) and helical forms(small arrow) of the NHP bacterium are abundantin the cytoplasm. 10 000x magnification.Lightner, D.V., R.M. Redman and J.R. Bonami.1992. Morphological evidence for a singlebacterial aetiology in Texas necrotizinghepatopancreatitis in Penaeus vannamei(Crustacea:Decapoda). Dis. Aquat. Org.13:235-239.210

FUNGAL DISEASE OF CRAYFISHC.11 CRAYFISH PLAGUEC.11.1Background InformationC.11.1.1 Causative AgentCrayfish Plague (also known as Krebspest,Kraftpest, ‘la peste’ or ‘crayfishaphanomyciasis’) is caused by the Oomycetefungus, Aphanomyces astaci. This is a closerelative of species associated with serious finfishdiseases, such as A. invadans, in EpizooticUlcerative Syndrome (EUS) of South-East Asia(see section F.11).C.11.1.2 Host RangeCrayfish plague affects the Noble crayfishAstacus astacus of north-west Europe, thestone crayfish Austropotamobius pallipes ofsouth-west and west Europe, the mountaincrayfish Austropotamobius torrentium of southwestEurope, and the slender clawed or Turkishcrayfish Astacus leptodactylus of easternEurope and Asia Minor. The Chinese mitten crab(Eriocheir sinensis) can be infected experimentally.North American crayfish (Pacifasticusleniusculus, the signal crayfish, andProcambarus clarkii, the Louisiana swamp crayfish)can also be infected by A. astaci, but arerelatively tolerant of the disease, only exhibitingclinical signs under intensive culture conditions.C.11.1.3 Geographical DistributionAphanomyces astaci is widespread in Europe,as well as in North America. The disease firstappeared in northern Italy in the mid 19th century,and then spread down to the Balkans andBlack Sea, as well as into Russia, Finland andSweden. In the 1960’s the disease appeared inSpain with further spread to the British Isles, Turkey,Greece and Norway in the 1980’s.survive initial infection can become sub-clinicalcarriers of the fungus. Under adverse holdingconditions, however, such infections maybecome pathogenic.C.11.3 Screening MethodsMore detailed information on methods forscreening crayfish plague can be found in theOIE Diagnostic Manual for Aquatic Animal Diseases(OIE 2000), at http://www.oie.int or selectedreferences.C.11.3.1 PresumptiveC.11.3.1.1 Gross Observations (Level I)Melanized spots in the cuticle of any crayfishspecies may be indicative of crayfish plaguesurvival. Such crayfish should be consideredto be potential carriers of the disease andscreened for Aphanomyces astaci using confirmatorydiagnostic techniques (C.11.3.2 andC.11.4.2).C.11.3.1.2 Microscopy (Level I/II)Foci of infection as described under C.12.3.1.1,may not be readily visible. Examination using adissecting microscope may reveal small whitenedpatches in the muscle tissues underlyingthin spots in the cuticle. There may also bebrownish discolouration of the cuticle. Finebrown lines through the cuticle should also beconsidered as suspect fungal hyphae. The areasthat should be examined closely are theintersternal soft-ventral cuticle of the abdomenand tail; the cuticle between the carapace andtail, the joints of the periopods (especially theproximal joints), the perianal cuticle and the gills.C.11.3.2 ConfirmatoryC.11.2Clinical AspectsC.11.3.2.1 Culture (Level II)The hyphae of A. astaci grow throughout the noncalcifiedparts of the cuticle and may extend alongthe nerve cord. The more disease tolerant speciesof crayfish (North American) encapsulate thefungal hyphae within melanised nodules, arrestingthe hyphal proliferation. Susceptible speciesappear incapable of producing such a defensereaction, and the fungus proliferates throughoutthe epicuticle and exocuticular layers of the exoskeleton.The cuticle and related soft-tissuedamage leads to death which, under warm waterconditions, can be rapid and result in 100%mortality. Resistant North American species thatThe fungus can be isolated from suspect cuticleand tissues using an agar medium thatcontains yeast extract, glucose and antibiotics(penicillin G and oxolinic acide) made up withnatural (not demineralised) river water. Identificationto species requires morphologicalcharacterisation of the sexual reproductiveparts of the fungus, however, these stages areabsent in A. astaci, thus, confirmation of infectionis usually based on isolation of fungalcolonies with the following characteristics (sinceno other closely-related Oomycetes are knownto infect crayfish):211

C.11 Crayfish Plague• growth within the agar medium (unless culturedat < 7°C, which promotes superficialgrowth);• colourless colonies;• aseptate, highly branching, vegetative hyphae,7-9 µm in diameter (min-max 5-10 µm);• young hyphae are densely packed withcoarse, granular cytoplasm and containhighly refractile globules;• older hyphae are highly vacuolated and theoldest hyphae appear to be emptyWhen thalli are transferred from the culturemedium to sterile distilled water, they developsporangia within 12-15 h (20°C) or 20-30 h(16°C). Elongate, irregularly amoeboid shapedspores are released and rapidly encyst as amass around the sporangial tip(Fig.C.11.3.2.1a). Encysted primary sporesmeasure 9-11 µm in diameter (min-max 8-15µm). Release of the secondary zoospores occursfrom papillae that develop on the surfaceof the primary spore cyst. This occurs at temperaturesas low as 4°C, peaking at 20°C andstopping at temperatures >24°C. Thezoospores have lateral flagella and measure 8x 12 µm. More details on culture media, techniquesand developmental stage morphologyare provided in the OIE Manual (OIE 2000).(EAFP/DJ Alderman)Fig. C.11.3.2.1a. Fresh microscopic mount ofa piece of infected exoskeleton showing fungalspores.(EAFP/DJ Alderman)aC.11.3.2.2 Bioassay (Level I/II)Confirmation of crayfish plague can be doneusing zoospores cultured from fungal isolatesfrom suspect crayfish tissues. Rapid mortalitiesin the susceptible crayfish, along with reisolationof the fungus as described above,should be considered conclusive for A. astaci.C.11.4 Diagnostic MethodsMore detailed information on methods for diagnosisof crayfish plague can be found in theOIE Diagnostic Manual for Aquatic Animal Diseases(OIE 2000), at http://www.oie.int or selectedreferences.There is no other disease, or pollution effect,that can cause total mortality of crayfish butleave all other animals in the same water unharmed.In such situations and with known susceptiblespecies, presumptive diagnosis can befairly conclusive. In first-time cases or in situationswith resistant species, however, confirmatoryisolation of the pathogen is recommended.bFig. C.11.4.1.1a,b. Clinical signs of infectedcrayfish showing whitened necrotic musculaturein the tail, and often accompanied inchronic infections by melanisation (blackening)of affected exoskeleton.C.11.4.1 PresumptiveC.11.4.1.1 Gross observation (Level I)Large numbers of crayfish showing activity duringdaylight should be considered suspect,since crayfish are normally nocturnal. Somemay show uncoordinated movement, easily tiponto their backs, and be unable to right themselves.Gross clinical signs of crayfish plague vary fromnone to a wide range of external lesions. White212

C.11 Crayfish Plaguepatches of muscle tissue underlying transparentareas of cuticle (especially the ventral abdomenand periopod joints), and focal brownmelanised spots (Fig.C.11.4.1.1a,b), are themost consistent signs.avoided or undertaken with disinfection precautions.Sodium hypochlorite and iodophores canbe used to disinfect equipment and thoroughdrying (>24 hours) is also effective, sinceoomycetes cannot withstand desiccation.C.11.4.1.2 Microscopy (Level I/II)C.11.7Selected ReferencesAs for C.11.3.1.2.C.11.4.2 ConfirmatoryC.11.4.2.1 Culture (Level II)As for C.12.3.2.1, diagnosis of crayfish plaguerequires the isolation and characterisation of thepathogen, A. astaci, using mycological mediafortified with antibiotics to control bacterial contamination.Isolation is only likely to be successfulbefore or within 12 hours of the death ofinfected crayfish.C.11.4.2.2 Bioassay (Level I/II)As for C.11.3.2.2.C.11.5Mode of TransmissionTransmission is horizontal and direct via themotile biflagellate zoospore stage of A. astaci,which posseses a positive chemotaxis towardscrayfish. The disease can spread downstreamat the speed of flow of the river, and has beendocumented to spread upstream at 2-4 km peryear. The upstream spread is suspected todriven by movements of crayfish between infectionand the terminal stages of the disease.Transmission has also been linked to the waterused to move fish between farms, as well as tocontaminated equipment (boots, fishing gear,crayfish traps, etc.). Introductions of NorthAmerican crayfish for crayfish farming are believedto have been the source of the Europeanoutbreaks of crayfish plague.C.11.6Control MeasuresThere is no treatment for crayfish plague, andthe high levels of mortality have precluded naturalselection for disease resistance in the mostsusceptible species (some populations are nowendangered). Control of the disease is bestachieved by preventing introductions or escapeof crayfish into unaffected waters. In addition,movement of water or any equipment betweenaffected to unaffected watersheds should beAlderman, D.J. 1996. Geographical spread ofbacterial and fungal diseases of crustaceans.OIE International Conference on the preventionof diseases of aquatic animals throughinternational trade. Office International desEpizooties, Paris, France, June 7-9 1995. Rev.Sci. Tech. Off. Int. Epiz. 15: 603-632.Alderman, D.J. and J.L. Polglase. 1986.Aphanomyces astaci: isolation and culture.J. Fish Dis. 9: 367-379.Alderman, D.J., J.L. Polglase, . Frayling and J.Hogger. 1984. Crayfish plague in Britain. J.Fish Dis. 7(5): 401-405.Alderman, D.J., J.L. Polglase and M. Frayling.1987. Aphanomyces astaci pathoogenicityunder laboratory and field conditions. J. FishDis. 10: 385-393.Alderman, D.J., D. Holdich and I. Reeve. 1990.Signal crayfish as vectors of crayfish plaguein Britain. Aquac. 86(1): 306.Dieguez-Uribeondo, J., C. Temino and J.L.Muzquiz. 1997. The crayfish plagueAphanomyces astaci in Spain. Bull. Fr. PechePiscic. 1(347): 753-763.Fuerst, M. 1995. On the recovery of Astacusastacus L. populations after an epizootic ofthe crayfish plague (Aphanomyces astaciShikora). Eighth Int. Symp. Astacol., LouisianaState Univ. Printing Office, Baton Rouge,LA, pp. 565-576.Holdich, D.M. and I.D. Reeve. 1991. Distributionof freshwater crayfish in the British Isles,with particular reference to crayfish plague,alien introductions and water quality. Aquat.Conserv. Mar. Freshwat. Ecosyst, 1(2): 139-158.Lilley, J.H. and V. Inglis. 1997. Comparative effectsof various antibiotics, fungicides anddisinfectants on Aphanomyces invaderis andother saprolegniaceous fungi. Aquac. Res.28(6): 461-469.213

C.11 Crayfish PlagueLilley, J.H., L. Cerenius and K. Soderhall. 1997.RAPD evidence for the origin of crayfishplague outbreaks in Britain. Aquac. 157(3-4): 181-185.Nylund, V. and K. Westman. 1995. Fequency ofvisible symptoms of the crayfish plague fungus(Aphanomyces astaci) on the signal crayfish(Pacifasticus leniusculus) in natural populationsin Finland in 1979-1988. Eighth Int.Symp. Astacol., Louisiana State Univ. PrintingOffice, Baton Rouge, LA.Oidtmann, B., M. El-Matbouli, H. Fischer, R.Hoffmann, K. Klaerding, I. Schmidt and R.Schmidt. 1997. Light microscopy of Astacusastacus L. under normal and selected pathologicalconditions, with special emphasis toporcelain disease and crayfish plague. FreshwaterCrayfish 11. A Journal of Astacology,Int. Assoc. Astacology, pp. 465-480.Oidtmann B., L. Cerenius, I. Schmid, R. Hoffmanand K. Soederhaell. 1999. Crayfish plagueepizootics in Germany – classification of twoGerman isolates of the crayfish plague fungusApahnomyces astaci by random amplificationof polymorphic DNA. Dis. Aquat. Org.35(3): 235-238.OIE. 2000. Diagnostic Manual for Aquatic AnimalDiseases, Third Edition, 2000. Office Internationaldes Epizooties, Paris, France.237p.Reynolds, J.D. 1988. Crayfish extinctions andcrayfish plague in central Ireland. Biol.Conserv. 45(4): 279-285.Vennerstroem, P., K. Soederhaell andL.Cerenius. 1998. The origin of two crayfishplague (Aphanomyces astaci) epizootics inFinland on noble crayfish, Astacus astacus.Ann. Zool. Fenn. 35(1): 43-46.214

ANNEX C.AI. OIE REFERENCE LABORATORYFOR CRUSTACEAN DISEASESDiseaseCrustacean pathogensExpert/LaboratoryProf. D. LightnerAquaculture Pathology SectionDepartment of Veterinary ScienceUniversity of ArizonaBuilding 90, Room 202Tucson AZ 85721USATel: (1.520) 621.84.14Fax: (1.520) 621.48.99E-mail: dvl@u.arizona.eduProf. S.N. ChenDepartment of ZoologyDirector, Institute of Fishery BiologyNational Taiwan UniversityNo. 1 Roosevelt RoadSection 4 , Taipei, Taiwan 10764TAIWAN PROVINCE of CHINATel: 886-2-368-71-01Fax: 886-2-368-71-22E-mail: snchen@cc.ntu.edu.tw215

ANNEX C.AII. LIST OF REGIONAL RESOURCEEXPERTS FOR CRUSTACEAN DISEASESIN ASIA-PACIFIC 1DiseaseShrimp diseasesExpertDr. Richard CallinanNSW Fisheries, Regional Veterinary LaboratoryWollongbar NSW 2477AUSTRALIATel (61) 2 6626 1294Mob 0427492027Fax (61) 2 6626 1276E-mail: richard.callinan@agric.nsw.gov.auDr. Indrani KarunasagarDepartment of Fishery MicrobiologyUniversity of Agricultural SciencesMangalore – 575 002INDIATel: 91-824 436384Fax: 91-824 436384E-mail: mircen@giasbg01.vsnl.net.inDr. C.V. MohanDepartment of AquacultureCollege of FisheriesUniversity of Agricultural SciencesMangalore-575002INDIATel: 91 824 439256 (College); 434356 (Dept), 439412 (Res)Fax: 91 824 438366E-mail: cv_mohan@yahoo.comProf. Mohammed ShariffFaculty of Veterinary MedicineUniversiti Putra Malaysia43400 Serdang, SelangorMALAYSIATel: 603-9431064; 9488246Fax: 603-9488246; 9430626E-mail: shariff@vet.upm.edu.myDr. Jie HuangYellow Sea Fisheries Research InstituteChinese Academy of Fishery Sciences106 Nanjing RoadQingdao, Shandong 266071PEOPLE’S REPUBLIC of CHINATel: 86 (532) 582 3062Fax: 86 (532) 581 1514E-mail: aqudis@public.qd.sd.cnDr. Jian-Guo HeSchool of Life SciencesZhongshan UniversityGuangzhou 510275PEOPLE’S REPUBLIC of CHINATel: +86-20-84110976Fax: +86-20-84036215E-mail: lsbrc05@zsu.edu.cnDr. Juan D. AlbaladejoFish Health SectionBureau of Fisheries and Aquatic ResourcesArcadia Building, 860 Quezon AvenueQuezon City, Metro ManilaPHILIPPINES216

Annex C.AII. List of Regional Resource Expertsfor Crustacean Diseases in Asia-PacificTel/Fax: 632-372-5055E-mail: jalbaladejo99@yahoo.comDr. Joselito R. SomgaFish Health SectionBureau of Fisheries and Aquatic ResourcesArcadia Building, 860 Quezon AvenueQuezon City, Metro ManilaPHILIPPINESTel/Fax: 632-372-5055E-mail: jrsomga@edsamail.com.phDr. Leobert de la PenaFish Health SectionAquaculture DepartmentSoutheast Asian Fisheries Development CenterTigbauan, Iloilo 5021PHILIPPINESTel: 63 33 335 1009Fax: 63 33 335 1008E-mail: leobert65@yahoo.com; leobertd@aqd.seafdec.org.phDr. P.P.G.S.N. SiriwardenaHead, Inland Aquatic Resources and AquacultureNational Aquatic Resources Research and Develoment AgencyColombo 15,SRI LANKATel: 941-522005Fax: 941-522932E-mail: sunil_siriwardena@hotmail.comDr. Yen-Ling SongDepartment of ZoologyCollege of ScienceNational Taiwan University1, Sec. 4, Roosevelt Rd.TAIWAN PROVINCE OF CHINAE-mail: yenlingsong@hotmail.comDr. Pornlerd ChanratchakoolAquatic Animal Health Research InstituteDepartment of FisheriesKasetsart University CampusJatujak, Ladyao, Bangkok 10900THAILANDTel: 662-5794122Fax: 662-5613993E-mail: pornlerc@fisheries.go.thMr. Daniel F. FeganNational Center for Genetic Engineering and Biotechnology (BIOTEC)Shrimp Biotechnology Programme18th Fl. Gypsum BuidlingSri Ayuthya Road, BangkokTHAILANDTel: 662-261-7225Fax:662-261-7225E-mail: dfegan@usa.netDr. Chalor LimsuanChalor LimsuwanFaculty of Fisheries, Kasetsart UnviersityJatujak, Bangkok 10900THAILANDTel: 66-2-940-5695217

Annex C.AII. List of Regional Resource Expertsfor Crustacean Diseases in Asia-PacificShrimp VirusesBacterial DiseasesDr. Gary NashCenter for Excellence for Shrimp Molecular Biology and BiotechnologyChalerm Prakiat BuildingFaculty of Science, Mahidol UniversityRama 6 RoadBangkok 10400THAILANDTel: 66-2-201-5870 to 5872Fax: 66-2-201-5873E-mail: gnash@asiaaccess.net.thDr Nguyen Thanh PhuongAquaculture and Fisheries Sciences Institute (AFSI)College of AgricultureCantho University, CanthoVIETNAMTel.: 84-71-830-931/830246Fax: 84-71-830-247.E-mail: ntphuong@ctu.edu.vnDr Peter WalkerAssociate Professor and Principal Research ScientistCSIRO Livestock IndustriesPMB 3 Indooroopilly Q 4068AUSTRALIATel: 61 7 3214 3758Fax: 61 7 3214 2718E-mail : peter.walker@tag.csiro.auMrs P.K.M. WijegoonawardenaNational Aquatic Resources Research and Development AgencyColombo 15,SRI LANKATel: 941-522005Fax: 941-522932E-mail: priyanjaliew@hotmail.comProf. Tim FlegelCentex Shrimp, Chalerm Prakiat BuildingFaculty of Science, Mahidol UniversityRama 6 Road, Bangkok 10400THAILANDPersonal Tel: (66-2) 201-5876Office Tel: (66-2) 201-5870 or 201-5871 or 201-5872Fax: (66-2) 201-5873Mobile Phone: (66-1) 403-5833E-mail: sctwf@mahidol.ac.thMrs. Celia Lavilla-TorresFish Health SectionAquaculture DepartmentSoutheast Asian Fisheries Development CenterTigbauan, Iloilo 5021PHILIPPINESTel: 63 33 335 1009Fax: 63 33 335 1008E-mail: celiap@aqd.seafdec.org.ph218

ANNEX C.AIII. LIST OF USEFUL DIAGNOSTICMANUALS/GUIDES TO CRUSTACEANDISEASES IN ASIA-PACIFICAsian Fish Health Bibliography III Japan by Wakabayashi H (editor). Fish Health SpecialPublication No. 3. Japanese Society of Fish Pathology, Japan and Fish Health Section of AsianFisheries Society, Manila, PhilippinesInformation: Japanese Society of Fish PathologyManual for Fish Diseases Diagnosis: Marine Fish and Crustacean Diseases in Indonesia(1998) by Zafran, Des Roza, Isti Koesharyani, Fris Johnny and Kei YuasaInformation: Gondol Research Station for Coastal FisheriesP.O. Box 140 Singaraja, Bali, IndonesiaTel: (62) 362 92278Fax: (62) 362 92272Health Management in Shrimp Ponds. Third Edition (1998) by P. Chanratchakool, J.F.Turnbull,S.J.Funge-Smith,I.H. MacRae and C. Limsuan.Information: Aquatic Animal Health Research InstituteDepartment of FisheriesKasetsart University CampusJatujak, Ladyao, Bangkok 10900THAILANDTel: (66.2) 579.41.22Fax: (66.2) 561.39.93E-mail: aahri@fisheries.go.thFish Health for Fishfarmers (1999) by Tina ThorneInformation: Fisheries Western Australia3rd Floor, SGIO Atrium186 St. Georges Terrace, Perth WA 6000Tel: (08) 9482 7333 Fax: (08) 9482 7389Web: http://www.gov.au.westfishAustralian Aquatic Animal Disease – Identification Field Guide (1999) by Alistair Herfort andGrant RawlinInformation: AFFA Shopfront – Agriculture, Fisheries and Forestry – AustraliaGPO Box 858, Canberra, ACT 2601Tel: (02) 6272 5550 or free call: 1800 020 157Fax: (02) 6272 5771E-mail: shopfront@affa.gov.auDiseases in Penaeid Shrimps in the Philippines. Second Edition (2000). By CR Lavilla-Pitogo,G.D. Lio-Po, E.R. Cruz-Lacierda, E.V. Alapide-Tendencia and L.D. de la PenaInformation: Fish Health SectionSEAFDEC Aquaculture DepartmentTigbauan, Iloilo 5021, PhilippinesFax: 63-33 335 1008E-mail: aqdchief@aqd.seafdec.org.phdevcom@aqd.seafdec.org.phManual for Fish Disease Diagnosis - II: Marine Fish and Crustacean Diseases in Indonesia(2001) by Isti Koesharyani, Des Roza, Ketut Mahardika, Fris Johnny, Zafran and Kei Yuasa,edited by K. Sugama, K. Hatai, and T NakaiInformation: Gondol Research Station for Coastal FisheriesP.O. Box 140 Singaraja, Bali, IndonesiaTel: (62) 362 92278Fax: (62) 362 92272219

Annex C.AIII.List of Useful Diagnostic Manuals/Guides to Crustacean Diseases in Asia-PacificReference PCR Protocol for Detection of White Spot Syndrome Virus (WSSV) in Shrimp.Shrimp Biotechnology Service Laboratory. Vol. 1, No. 1, March 2001Information: Shrimp Biotechnology Service Laboratory73/1 Rama 6 Rd., Rajdhewee, Bangkok 10400Tel. (662) 644-8150Fax: (662) 644-8107220

LIST OFNATIONAL COORDINATORS*CountryAustraliaBangladeshCambodiaChina P.R.DPR KoreaName and AddressDr. Eva -Maria BernothManager, Aquatic Animal Health Unit,Office of the Chief Veterinary OfficerDepartment of Agriculture, Fisheries and ForestryGPO Box 858, Canberra ACT 2601, AustraliaFax: 61-2-6272 3150; Tel: 61-2-6272 4328Email: Eva-Maria.Bernoth@affa.gov.auDr. Alistair Herfort (Focal point for disease reporting)Aquatic Animal Health Unit , Office of the Chief Veterinary OfficerDepartment of Agriculture, Fisheries and ForestryGPO Box 858, Canberra ACT 2601, AustraliaFax: +61 2 6272 3150; tel: +61 2 6272 4009E-mail: Alistair.Herfort@affa.gov.auDr. M. A. MazidDirector General, Bangladesh Fisheries Research Institute (BFRI)Mymensingh 2201, BangladeshFax: 880-2-55259, Tel: 880-2-54874E-mail: frifs@bdmail.netMr. Srun Lim SongHead, Laboratory Section, Department of Fisheries186 Norodom Blvd.,P.O. Box 835, Phnom Penh, CambodiaFax: (855) 23 210 565; Tel: (855) 23 210 565E-mail: smallfish@bigpond.com.khMr. Wei QiExtension Officer, Disease Prevention and Control DivisionNational Fisheries Technology Extension Centre, No. 18Ministry of AgricultureMai Zi dian Street, Chaoyang District, Beijing 100026, ChinaFax: 0086-1—65074250; Tel: 0086-10-65074250E-mail: weiqi_moa@hotmail.comProf. Yang Ningsheng (Focal point for AAPQIS)Director, Information Center, China Academy of Fisheries Science150 Qingta Cun, South Yongding Road, Beijing 100039, ChinaFax: 86-010-68676685; Tel: 86-010-68673942E-mail: ningsheng.yang@mh.bj.col.com.cnMr. Chong Yong HoDirector of Fish Farming Technical DepartmentBureau of Freshwater CultureSochangdong Central District, P.O.Box. 95 , Pyongyong, DPR KoreaFax- 850-2-814416; Tel- 3816001, 3816121*The matrix provides a list of National Coordinators nominated by Governments and focal points for the Asia-Pacific QuarterlyAquatic Animal Disease Reports.221

List of National CoordinatorsHong Kong ChinaIndiaIndonesiaIranJapanLao PDRDr. Roger S.M. ChongNational Coordinator and Fish Health OfficerAgriculture, Fisheries and Conservation DepartmentCastle Peak Veterinary LaboratorySan Fuk Road, Tuen MunNew Territories, Hong KongFax: +852 2461 8412Tel: + 852 2461 6412E-mail: vfhoafd@netvigator.comDr. AG PonniahDirectorNational Bureau of Fish Genetic ResourcesCanal Ring Road, P.O. DilkushaLucknow-226 002, U.O., IndiaFax: (911-522) 442403; Tel: (91-522) 442403/442441E-mail: nbfgr@1w1.vsnl.net.in; nbfgr@400.nicgw.nic.inShri M.K.R. NairFisheries Development CommissionerMr. Bambang Edy PriyonoNational Coordinator (from September 2000)Head, Division of Fish Health ManagementDirectorate General of FisheriesJl. Harsono RM No. 3Ragunan Pasar MingguTromol Pos No.: 1794/JKSJakarta – 12550 IndonesiaTel: 7804116-119Fax: 7803196 – 7812866E-mail: dfrmdgf@indosat.net.idDr. Reza PourgholamNational Coordinator (from November 2000)Veterinary OrganizationMinistry of Jihad – E – SazandegiVali-ASR AveS.J.Asad Abadi StPO Box 14155 – 6349Tehran, IranTel: 8857007-8857193Fax: 8857252Dr. Shunichi ShinkawaFisheries Promotion Division, Fishery Agency1-2-1, KasumigasekiChiyoda-ku, Tokyo 100-8907, JapanFax: 813-3591-1084; Tel: 813-350-28111(7365)E-mail: shunichi_shinkawa@nm.maff.go.jpMr. Bounma Luang AmathFisheries and Livestock DepartmentMinistry of Agriculture, Forestry and FisheriesP.O. Box 811, Vientianne, Lao PDRTeleFax: (856-21) 415674; Tel: (856-21) 4169321The experts included in this list have previously been consulted and agreed to provide valuable information and health adviseconcerning their particular expertise.222

List of National CoordinatorsMalaysia Mr. Ambigadevi Palanisamy (from September 2001)National CoordinatorFisheries Research InstituteDepartment of FisheriesPenang, MalaysiaE-mail: ambigadevip@yahoo.comDr. Ong Bee Lee (focal point for disease reporting)Head, Regional Veterinary Laboratory ServicesDepartment of Veterinary Services8th & 9th Floor, Wisma Chase PerdanaOff Jln Semantan 50630, Kuala Lumpur, MalaysiaFax: (60-3) 254 0092/253 5804; Tel: (60-3) 254 0077 ext.173E-mail: ong@jph.gov.myMyanmarNepalPakistanPhilippinesRepublic of KoreaMs. Daw May Thanda WintAssistant Staff Officer, Aquatic Animal Health SectionDepartment of FisheriesSinmin Road, Alone Township , Yangon, MyanmarFax: (95-01) 228-253; Tel: (95-01) 283-304/705-547Mr. M. B. PanthaChief, District Agri Devt. OfficerDist Agric. Devt OfficeJanakpur, DhanushaNepalFax: (977-1) 486895E-mail: image@bhawani.wlink.com.npRana Muhammad IqbalAssistant Fisheries Development Commissioner IIMinistry of Food, Agriculture and CooperativesR#310, B-Block, Islamabad,Government of Pakistan, Islamabad, PakistanFax: 92-051-9201246; Tel: 92-051-9208267Dr. Rukshana AnjumAssistant Fisheries Development CommissionerMinistry of Food, Agriculture and LivestockGovernment of PakistanFax No. 051 9221246Dr. Joselito R. SomgaAquaculturist II, Fish Health Section, BFAR860 Arcadia Building, Quezon Avenue, Quezon City 1003Fax: (632)3725055/4109987;Tel:(632) 3723878 loc206 or 4109988 to 89E-mail: sssomga@edsamail.co.phDr. Mi-Seon ParkDirector of Pathology DivisionNational Fisheries Research and Development Institute408-1 Sirang, KijangPusan 619-900 Korea ROTel: 82-51-720-2470; Fax: 82-51-720-2498E-mail: parkms@haema.nfrda.re.kr223

List of National CoordinatorsSingaporeSri LankaThailandVietnamMr. Chao Tien MeeSAVAO (Senior Agri-Food and Veterinary Authority Officer)OIC, Marine Aquaculture Centre (MAC)Agri-Food & Veterinary Authority of Singapore (AVA)300 Nicoll Drive, Changi Point, Singapore 498989Tel: (65) 5428455; Fax No.: (65) 5427696E-mail: CHAO_Tien_Mee@ava.gov.sgDr. Chang Siow Foong (focal person for disease reporting)Agri-Food and Veterinary Authority of SingaporeCentral Veterinary Laboratory60 Sengkang East WaySingapore 548596Tel: (65) 3863572; Fax No. (65) 3862181E-mail: CHANG_Siow_Foong@AVA.gov.sgMr. A. M. JayasekeraDirector-GeneralNational Aquaculture Development Authority of Sri LankaMinistry of Fisheries and Aquatic Resources Development,317 1/1 T.B. Jayah Mawatha, Colombo 10, Sri LankaTel: (94-1) 675316 to 8; Fax: (94-1) 675437E-mail: aqua1@eureka.lkDr. Geetha Ramani Rajapaksa (focal point for disease reporting)Veterinary SurgeonDepartment of Animal Production and HealthVeterinary Investigation Centre, Welisara, Ragama, Sri LankaTel: + 01-958213E-mail: sser@sri.lanka.netDr. Somkiat KanchanakhanFish Virologist, Aquatic Animal Health Research Institute (AAHRI)Department of Fisheries , Kasetsart University CampusJatujak, Bangkok 10900, ThailandFax: 662-561-3993; Tel: 662-579-4122, 6977E-mail: somkiatk@fisheries.go.thDr. Le Thanh LuuVice-DirectorResearch Institute for Aquaculture No. 1 (RIA No. 1)Dinh Bang, Tien Son, Bac Ninh, VietnamFax: 84-4-827-1368; Tel: 84-4-827-3070E-mail: ria1@hn.vnn.vnMs Dang Thi Lua (Focal point for disease reporting)Researcher, Research Institute for Aquaculture No.1 (RIA No.1 )Dinh Bang , Tien Son, Bac Ninh, VietnamFax: 84-4-827-1368; Tel : 84-4-827 - 3070E-mail: ria1@hn.vnn.vn; danglua@hotmail.com224

MEMBERS OF THE REGIONALWORKING GROUP(RWG, 1998 TO 2001)Dr. Eva-Maria BernothManager, Aquatic Animal Health UnitOffice of the Chief Veterinary OfficerDepartment of Agriculture, Fisheries and Forestry – AustraliaGPO Box 858, Canberra Act 2601AUSTRALIATel: 61-2-6272-4328Fax: 61-2-6272-3150E-mail: Eva-Maria.Bernoth@affa.gov.auMr. Daniel FeganApt. 1D Prestige Tower B168/25 Sukhumvit Soi 23Klongtoey, Bangkok 10110THAILANDTel: (662) 261-7225/(661) 825-8714Fax: (662) 261-7225E-mail: dfegan@usa.netProfessor Jiang YulinShenzhen Exit & Entry Inspection and Quarantine Bureau40 Heping Road, Shenzhen 518010PEOPLE’S REPUBLIC OF CHINATel: 86-755-5592980Fax: 86-755-5588630E-mail: szapqbxi@public.szptt.net.cnDr. Indrani KarunasagarUNESCO MIRCEN for Marine BiotechnologyDepartment of Fishery MicrobiologyUniversity of Agricultural SciencesCollege of FisheriesMangalore – 575 002KarnatakaINDIATel: 91-824 436384Fax: 91-824 436384/91-824 438366E-mail: mircen@giasbg01.vsnl.net.inMs. Celia Lavilla-Pitogo TorresSEAFDEC Aquaculture Department5021 TigbauanREPUBLIC OF THE PHILIPPINESTel: 63-33 336 2965Fax: 63-33 335 1008 E-mail: celiap@seafdec.org.phProfessor Mohammed ShariffFaculty of Veterinary MedicineUniversiti Putra Malaysia43400 Serdang, Selangor Darul EhsanMALAYSIATel: 60-3-89431064Fax: 60-3-89430626E-mail: shariff@vet.upm.edu.my225

Members of the Regional WorkingGroup (RWG, 1998 to 2001)Dr. Kamonporn TonguthaiDepartment of FisheriesKasetsart University CampusLadyao, Jatujak, Bangkok 10900THAILANDTel: (662) 940-6562Fax: (662) 562-0571E-mail: kamonpot@fisheries.go.thDr. Yugraj Singh YadavaNational Agriculture Technology ProjectMinistry of AgriculturePusa, New Delhi 110012INDIATel: (91-11)-6254812 (residence)(91-11) 5822380/5822381 (office)E-mail: y.yugraj@mailcity.com226

MEMBERS OF THE TECHNICALSUPPORT SERVICES:Dr. James Richard ArthurFAO ConsultantRR1, Box 13, Savarie Rd.Sparwood, B.C.Canada V0B 2G0Tel: 250-425-2287Fax: 250 425-0045 (indicate for delivery to R. Arthur, Tel. 425-2287)E-mail: rarthur@titanlink.comDr. Chris BaldockDirector, AusVet Animal Health ServicesPO Box 3180South Brisbane Qld 4101AUSTRALIATel: 61-7-3255 1712Fax: 61-7-3511 6032E-mail: ausvet@eis.net.auMr. Pedro BuenoCo-ordinatorNetwork of Aquaculture Centres inAsia-PacificDepartment of FisheriesKasetsart University CampusLadyao, Jatujak, Bangkok 10900THAILANDTel: (662) 561-1728 to 9Fax: (662) 561-1727E-mail: pedro.bueno@enaca.orgDr. Supranee ChinabutDirector, Aquatic Animal Health Research InstituteDepartment of FisheriesKasetsart University Campus,Ladyao, Jatujak, Bangkok 10900THAILANDTel: (662) 579-4122Fax: (662) 561-3993E-mail: supranee@fisheries.go.thProfessor Timothy FlegelDepartment of BiotechnologyFaculty of ScienceMahidol UniversityRama 6 Road, Bangkok 10400THAILANDTel: (662) 245-5650Fax: (662) 246-3026E-mail: sctwf@mahidol.ac.thProfessor Tore HasteinNational Veterinary InstituteUllevalsveien 68,P.O. Box 8156 Dep. 0033NORWAYTel: 47 22964710Fax: 47 22463877E-mail: Tore.Hastein@vetinst.no227

Members of the TechnicalSupport Services:Dr. Barry HillOIE Fish Disease CommissionCEFAS Weymouth LaboratoryThe Nothe, Weymouth, Dorset DT4 8UBUNITED KINGDOMTel: 44-1305 206 626Fax: 44-1305-206 627E-mail: B.J.HILL@cefas.co.ukMr. Hassanai KongkeoSpecial Advisor Network of Aquaculture Centres in Asia-PacificDepartment of FisheriesKasetsart University CampusLadyao, Jatujak, Bangkok 10900THAILANDTel: (662) 561-1728 to 9Fax: (662) 561-1727E-mail: hassanak@fisheries.go.thDr. Sharon E. McGladderyShellfish Health PathologistDepartment of Fisheries and Oceans – CanadaGulf Fisheries Centre, P.O. Box 5030Moncton, NB, CANADA E1C 9B6Tel: 506 851-2018Fax: 506 851-2079E-mail: McGladderyS@mar.dfo-mpo.gc.caDr. Kazuhiro NakajimaHead, Pathogen SectionFish Pathology DivisionNational Research Institute of Aquaculture422-1 Nansei-cho, Watarai-gunMie 516-0193JAPANTel: 81-599 66-1830Fax: 81-599 6 6-1962E-mail: kazuhiro@nria.affrc.go.jpDr. Yoshihiro OzawaOIE Representation for Asia and the PacificOIE Tokyo, East 311, Shin Aoyama Bldg1-1-1 Minami-aoyama, Minato-kuTokyo 107JAPANTel: 81-3-5411-0520Fax: 81-3-5411-0526E-mail: Oietokyo@tky.3web.ne.jpDr. Michael J. PhillipsEnvironment SpecialistNetwork of Aquaculture Centres inAsia-PacificDepartment of FisheriesKasetsart University CampusLadyao, Jatujak, Bangkok 10900THAILANDTel: (662) 561-1728 to 9Fax: (662) 561-1727228

Members of the TechnicalSupport Services:E-mail: Michael.Phillips@enaca.orgDr. Melba B. ReantasoAquatic Animal Health SpecialistNetwork of Aquaculture Centres inAsia-PacificDepartment of Fisheries, Kasetsart University CampusLadyao, Jatujak, Bangkok 10900THAILANDTel: (662) 561-1728 to 9Fax: (662) 561-1727E-mail: Melba.Reantaso@enaca.org; melbar@fisheries.go.thDr. Rohana P. SubasingheSenior Fishery Resources Officer(Aquaculture)Inland Water Resources andAquaculture ServiceFisheries Department,Food and Agriculture Organization ofthe United NationsViale delle Terme di CaracallaRome 00100ITALYTel: 39-06 570 56473Fax: 39-06 570 53020E-mail: Rohana.Subasinghe@fao.orgMr. Zhou Xiao WeiProgram Officer (Training)Network of Aquaculture Centres inAsia-PacificDepartment of FisheriesKasetsart University CampusLadyao, Jatujak, Bangkok 10900THAILANDTel: (662) 561-1728 to 9Fax: (662) 561-1727E-mail: zhoux@fisheries.go.th229

LIST OF FIGURESSECTION 2 – FINFISH DISEASESSECTION F.1 GENERAL TECHNIQUESFig. F.1.1.2.1a. Red spot disease of grass carp (MG Bondad-Reantaso)Fig. F.1.1.2.1b. Surface parasites, Lerneae cyprinacea infection of giant gouramy (JR Arthur)Fig. F.1.1.2.1c. Ayu, Plecoglossus altivelis, infected with Posthodiplostomum cuticola (?)metacercariae appearing as black spots on skin (K Ogawa)Fig. F.1.1.2.1d. Typical ulcerative, popeye, fin and tail rot caused by Vibrio spp. (R Chong)Fig. F.1.1.2.2a. Example of gill erosion on Atlantic salmon, Salmo salar, due to intenseinfestation by the copepod parasite Salmincola salmoneus (SE McGladdery)Fig. F.1.1.2.2b. Fish gills infected with monogenean parasites (MG Bondad-Reantaso)Fig. F.1.3.1a. Myxobolus artus infection in the skeletal muscle of 0+ carp (H Yokoyama)Fig. F.1.3.1b. Ligula sp. (Cestoda) larvae infection in the body cavity of Japanese yellow goby,Acanthogobius flavimanus (K Ogawa)Fig. F.1.3.2a. Distended abdomen of goldfish (H Yokoyama)Fig. F.1.3.2b. Japanese Yamame salmon (Onchorynchus masou) fingerlings showing swollenbelly due to yeast infection (MG Bondad-Reantaso)SECTION F.2 – EPIZOOTIC HAEMATOPOIETIC NECROSIS (EHN)Fig. F.2.2. Mass mortality of single species of redfin perch. Note the small size of fish affectedand swollen stomach of the individual to the center of the photograph. Note the characteristichaemorrhagic gills in the fish on the left in the inset (AAHL)SECTION F.3 – INFECTIOUS HAEMATOPOIETIC NECROSIS (IHN)Fig.F.3.2a. IHN infected fry showing yolk sac haemorrhages (EAFP)Fig.F.3.2b. Clinical signs of IHN infected fish include darkening of skin, haemorrhages on theabdomen and in the eye around the pupil (EAFP)SECTION F.4 ONCORHYNCHUS MASOU VIRUS (OMV)Fig. F.4.4.1.1a. OMV-infected chum salmon showing white spots on the liver (M Yoshimizu)Fig. F.4.4.1.1b. OMV-induced tumour developing around the mouth of chum salmon fingerling(M Yoshimizu)Fig. F.4.4.1.3. OMV particles isolated from masou salmon, size of nucleocapsid is 100 to 110nm (M Yoshimizu)SECTION F.5 INFECTIOUS PANCREATIC NECROSIS (IPN)Fig.F.5.2a. IPN infected fish showing dark colouration of the lower third of the body and smallswellings on the head (EAFP)Fig.F.5.2b. Rainbow trout fry showing distended abdomen characteristic of IPN infection.Eyed-eggs of this species were imported from Japan into China in 1987 (J Yulin)Fig.F.5.2c. Top: normal rainbow trout fry, below: diseased fry (EAFP)Fig. F.5.4.1.3. CPE of IHNV (J Yulin)Fig.F.5.4.1.4. IPN virus isolated from rainbow trout fry imported from Japan in 1987. Virusparticles are 55 nm in diameter (J Yulin)SECTION F.6 VIRAL ENCEPHALOPATHY AND RETINOPATHY (VER)Fig.F.6.2. Fish mortalities caused by VER (J Yulin)Figs. F.6.4.1.2a, b. Vacuolation in brain (Br) and retina (Re) of GNNV-infected grouper inChinese Taipei (bar = 100 mm) (S Chi Chi)230

List Of FiguresSECTION F.7 SPRING VIRAEMIA OF CARP (SVC)Figs.F.7.4.1.1a, b,c,d. Non-specific clinical signs of SVC infected fish, which may includeswollen abdomen, haemorrhages on the skin, abdominal fat tissue, swim bladder and otherinternal organs and in the muscle (EAFP)SECTION F.8 VIRAL HAEMORRHAGIC SEPTICAEMIA (VHS)Fig.F.8.4.1.1. Non-specific internal sign (petechial haemorrhage on muscle) of VHS infectedfish (EAFP)SECTION F.9 LYMPHOCYSTISFig.F.9.2a. Wild snakehead infected with lymphocystis showing irregularly elevated masses ofpebbled structure (MG Bondad-Reantaso)Fig.F.9.4.1.1a. Flounder with severe lymphocystis (J Yulin)Fig.F.9.4.1.1b. Lymphocystis lesions showing granular particle inclusions (J Yulin)Fig.F.9.4.1.1c. Carp Pox Disease caused by Herpesvirus (J Yulin)Fig.F.9.4.1.1d. Goldfish with fungal (mycotic) skin lesions (J Yulin)Fig.F.9.4.2.1a. Giant (hypertrophied) lymphoma cells with reticulate or branching inclusionbodies around the nuclei (J Yulin)Fig.F.9.4.2.1b. Impression smear of lymphocystis showing some giant cells, and hyalinecapsules (membrane) (J Yulin)Fig.F.9.4.2.2a. Electron micrograph showing numerous viral particles in cytoplasm (J Yulin)Fig. F.9.4.2.2b. Enlarged viral particles showing typical morphology of iridovirus (100 mm = bar)(J Yulin)Fig.F.9.4.2.2c. Herpervirus in Carp Pox showing enveloped and smaller virions compared tolymphocystis virus (J Yulin)SECTION F.10 BACTERIAL KIDNEY DISEASE (BKD)Fig.F.10.3.1.2a. Pinpoint colonies up to 2 mm in diameter of Renibacteriium salmonimarum,white-creamy, shiny, smooth, raised and entire; three weeks after incubation at 15°C on KDM-2 medium (M Yoshimizu)Fig.F.10.3.1.2b. Renibacterium salmoninarum rods, isolated from masou salmon(M Yoshimizu)Fig.F.10.4.1.1a. Kidney of masou salmon showing swelling with irregular grayish patches(M Yoshimizu)Fig.F.10.4.1.1b. Enlargement of spleen is also observed from BKD infected fish (EAFP)SECTION F.11 EPIZOOTIC ULCERATIVE SYNDROME (EUS)Fig.F.11.1.2a. Ayu, Plecoglatus altivelis, infected with mycotic granulomatosis (K Hatai)Fig.F.11.1.2b. EUS affected farmed silver perch Bidyanus bidyanus from Eastern Australia(RB Callinan)Fig.F.11.2a. Catfish showing initial EUS red spots (MG Bondad-Reantaso)Fig.F.11.2b. Snakehead in Philippines (1985) showing typical EUS lesions(MG Bondad-Reantaso)Fig.F.11.4.1.1a. Wild mullet in Philippines (1989) with EUS (MG Bondad-Reantaso)Fig.F.11.4.1.1b. Red spot disease of grass carp in Vietnam showing ulcerative lesions (MGBondad-Reantaso)Fig.F.11.4.1.2. Granuloma from squash preparation of muscle of EUS fish (MG Bondad-Reantaso)Fig.F.11.4.2.1a. Typical severe mycotic granulomas from muscle section of EUS fish (H & E)(MG Bondad-Reantaso)Fig.F.11.4.2.1b. Mycotic granulomas showing fungal hyphae (stained black) using Grocottsstain (MG Bondad-Reantaso)Fig.F.11.4.2.2a. Typical characteristic of Aphanomyces sporangium (K Hatai)Fig.F.11.4.2.2b. Growth of Aphanomyces invadans on GP agar (MG Bondad-Reantaso)231

List Of FiguresSECTION 3 MOLLUSCAN DISEASESSECTION M.1 GENERAL TECHNIQUESFig.M.1.1.1. Gaping hard shell clam, Mercenaria mercenaria, despite air exposure andmechnical tapping (SE McGladdery)Fig.M.1.1.2a. Mollusc encrustment (arrows) of winged oyster, Pteria penguin, Guian PearlFarm, Eastern Samar, Philippines (1996) (MG Bondad-Reantaso)Fig.M.1.1.2b. Pteria penguin cultured at Guian Pearl Farm, Eastern Samar, Philippines withextensive shell damage due to clionid (boring) sponge (1992) (D Ladra)Fig.M.1.1.2c,d. Pteria penguin shell with dense multi-taxa fouling, Guian Pearl Farm, EasternPhilippines (1996) (MG Bondad-Reantaso)Fig.M.1.1.2e. Polydora sp. tunnels and shell damage at hinge of American oyster, Crassostreavirginica, plus barnacle encrusting of other shell surfaces (SE McGladdery)Fig.M.1.1.2f. Winged oyster, Pteria penguin, shell with clionid sponge damage. Guian PearlFarm, Eastern, Philippines (1996) (MG Bondad-Reantaso)Fig.M.1.1.2g,h. Pinctada maxima, shell with clionid sponge damage due to excavation oftunnels exhalent-inhalent openings (holes) to the surface (arrows). Other holes (small arrows)are also present that may have been caused by polychaetes, gastropod molluscs or otherfouling organisms. Guian Pearl Farm, Eastern Philippines (1996) (MG Bondad-Reantaso)Fig.M.1.1.3a. Winged oyster, Pteria penguin, shell showing clionid sponge damage through tothe inner shell surfaces, Guian Pearl Farm, Eastern Philippines (1996) (MG Bondad-Reantaso)Fig.M.1.1.3a1. Abalone (Haliotis roei) from a batch killed by polydoriid worms (B Jones)Fig.M.1.1.3b,c. b. Shells of Pinctada maxima showing a erosion of the nacreous inner surfaces(arrows), probably related to chronic mantle retraction; c. Inner surface of shell showingcomplete penetration by boring sponges (thin arrows) (D Ladra)Fig.M.1.1.3d,e,f. Pinctada maxima (d), Pteria penguin (e) and edible oyster (Crassostrea sp.) (f)shells showing Polydora-related tunnel damage that has led to the formation of mud-filledblisters (MG Bondad-Reantaso)Fig.M.1.1.3g. Inner shell of winged pearl oyster showing: tunnels at edge of the shell (straightthick arrow); light sponge tunnel excavation (transparent arrow); and blisters (small thick arrow)at the adductor muscle attachment site. Guian Pearl Farm, Eastern Philippines (1996)(MG Bondad-Reantaso)Fig.M.1.1.3.h. Extensive shell penetration by polychaetes and sponges causing weakening andretraction of soft-tissues away from the shell margin of an American oyster Crassostreavirginica(SE McGladdery)Fig.M.1.1.4a. Normal oyster (Crassostrea virginica) tissues (SE McGladdery)Fig.M.1.1.4b. Watery oyster (Crassostrea virginica) tissues – compare with M.1.1.4a(SE McGladdery)Fig.M.1.1.4c. Abscess lesions (creamy-yellow spots) in the mantle tissue of a Pacific oyster(Crassostrea gigas) (SE McGladdery)Fig.M.1.1.4d. Gross surface lesions in Pacific oyster (Crassostrea gigas) due to Marteiliodeschungmuensis (MS Park and DL Choi)Fig.M.1.1.4e. Water blister (oedema/edema) in the soft-tissues of the mantle margin of anAmerican oyster (Crassostrea virginica) (SE McGladdery)Fig.M.1.1.4f. Calcareous deposits (“pearls”) in the mantle tissues of mussels in response toirritants such as mud or digenean flatworm cysts (SE McGladdery)Fig.M.1.1.4g. Polydoriid tunnels underlying the nacre at the inner edge of an American oyster(Crassostrea virginica) shell, plus another free-living polychaete, Nereis diversicolor on the innershell surface (SE McGladdery and M Stephenson)SECTION M.2 BONAMIOSISFig.M.2.2a. Haemocyte infiltration and diapedesis across intestinal wall of a European oyster(Ostrea edulis) infected by Bonamia ostreae (SE McGladdery)Fig.M.2.2b. Oil immersion of Bonamia ostreae inside European oyster (Ostrea edulis)haemocytes (arrows). Scale bar 20µm (SE McGladdery)Fig.M.2.2c. Systemic blood cell infiltration in Australian flat oyster (Ostrea angasi) infected by232

List Of FiguresBonamia sp. Note vacuolised appearance of base of intestinal loop and duct walls (H&E)(PM Hine)Fig.M.2.2d. Oil immersion of Bonamia sp. infecting blood cells and lying free (arrows) in thehaemolymph of an infected Australian flat oyster, Ostrea angasi. Scale bar 20µm (H&E)(PM Hine)Fig.M.2.2e. Focal infiltration of haemocytes around gut wall (star) of Tiostrea lutaria (NewZealand flat oyster) typical of infection by Bonamia sp. (H&E) (PM Hine)Fig.M.2.2f. Oil immersion of haemocytes packed with Bonamia sp. (arrows) in an infectedTiostrea lutaria (H&E) (PM Hine)SECTION M.3. MARTEILIOSISFig.M.3.2a. Digestive duct of a European oyster, Ostrea edulis, showing infection of distalportion of the epithelial cells by plasmodia (arrows) of Marteilia refringens. Scale bar 15µm(H&E) (SE McGladdery)Fig.M.3.2b. Digestive tubule of a European oyster, Ostrea edulis, showing refringent sporestage of Marteilia refringens (star). Scale bar 50µm (H&E) (SE McGladdery)Fig.M.3.4.1.1a. Tissue imprint from Saccostrea commercialis (Sydney rock oyster) heavilyinfected by Marteilia sydneyi (arrows) (QX disease). Scale bar 250µm (H&E) (RD Adlard)Fig.M.3.4.1.1b. Oil immersion of tissue squash preparation of spore stages of Marteilia sydneyifrom Sydney rock oyster (Saccostrea commercialis) with magnified inset showing two sporeswithin the sporangium. Scale bar 50µm (H&E) (RD Adlard)SECTION M.4 MIKROCYTOSISFig.M.4.2a. Gross abscess lesions (arrows) in the mantle tissues of a Pacific oyster(Crassostrea virginica) severely infected by Mikrocytos mackini (Denman Island Disease)(SM Bower)Fig.M.4.3.2.1a. Histological section through mantle tissue abscess corresponding to the grosslesions pictured in Fig.M.4.2a, in a Pacific oyster (Crassostrea gigas) infected by Mikrocytosmackini (H&E) (SM Bower)Fig.M.4.3.2.1b. Oil immersion of Mikrocytos mackini (arrows) in the connective tissue surroundingthe abscess lesion pictured in Fig.M.4.3.2.1a. Scale bar 20µm (H&E) (SM Bower)SECTION M.5. PERKINSOSISFig.M.5.1.2a. Arca clam showing a Perkinsus-like parasite within the connective tissue.Magnified insert shows details of an advanced ‘schizont’ like stage with trophozoites showingvacuole-like inclusions. Scale bar 100µm. (H&E) (PM Hine)Fig.M.5.1.2b. Pinctada albicans pearl oyster showing a Perkinsus-like parasite. Magnifiedinsert shows details of a ‘schizont’-like stage containing ‘trophozoites’ with vacuole-likeinclusions. Scale bar 250µm (H&E) (PM Hine)Fig.M.5.3.2.1a. Trophozoite (‘signet-ring’) stages of Perkinsus marinus (arrows), the cause of‘Dermo’ disease in American oyster (Crassostrea virginica) connective tissue. Scale bar 20µm(H&E) (SM Bower)Fig.M.5.3.2.1b. Schizont (‘rosette’) stages of Perkinsus marinus (arrows), the cause of ‘Dermo’disease in American oyster (Crassostrea virginica) digestive gland connective tissue. Scale bar30µm (H&E) (SE McGladdery)Fig.5.3.2.2. Enlarged hypnospores of Perkinsus marinus stained blue-black with Lugol’s iodinefollowing incubation in fluid thioglycollate medium. Scale bar 200µm (SE McGladdery)SECTION M.6 HAPLOSPORIDIOSISFig.M.6.1.3a. Massive connective tissue and digestive tubule infection by an unidentifiedHaplosporidium-like parasite in the gold-lipped pearl oyster Pinctada maxima from northWestern Australia. Scale bar 0.5 mm (H&E) (PM Hine)Fig.M.6.1.3b. Oil immersion magnification of the operculated spore stage of theHaplosporidium-like parasite in the gold-lipped pearl oyster Pinctada maxima from northWestern Australia. Scale bar 10 µm. (H&E) (PM Hine)233

List Of FiguresFig.M.6.1.3c. Haemocyte infiltration activity in the connective tissue of a Sydney rock oyster(Saccostrea cucullata) containing spores of a Haplosporidium-like parasite (arrow). Scale bar0.5 mm. (H&E) (PM Hine)Fig.M.6.1.3d. Oil immersion magnification of Haplosporidium-like spores (arrow) associatedwith heavy haemocyte infiltration in a Sydney rock oyster (Saccostrea cucullata). Scale bar10µm. (H&E) (PM Hine)Fig.M.6.3.1.2a. Plasmodia (black arrows) and spores (white arrows) of Haplosporidium costale,the cause of SSO disease, throughout the connective tissue of an American oyster(Crassostrea virginica). Scale bar 50µm (SE McGladdery)Fig.M.6.3.1.2b. Plasmodia (black arrows) and spores (white arrows) of Haplosporidium nelsoni,the cause of MSX disease, throughout the connective tissue and digestive tubules of anAmerican oyster (Crassostrea virginica). Scale bar 100µm (SE McGladdery)Fig.M.6.4.2.2a. Oil immersion magnification of SSO spores in the connective tissue of anAmerican oyster Crassostrea virginica. Scale bar 15µm (SE McGladdery)Fig.M.6.4.2.2b. Oil immersion magnification of MSX spores in the digestive tubule epitheliumof an American oyster Crassostrea virginica. Scale bar 25µm. (H&E) (SE McGladdery)SECTION M.7 MARTEILIODOSISFig.M.7.2a,b. a. Gross deformation of mantle tissues of Pacific oyster (Crassostrea gigas) fromKorea, due to infection by the protistan parasite Marteiloides chungmuensis causing retentionof the infected ova within the ovary and gonoducts; b. (insert) normal mantle tissues of aPacific oyster (MS Park and DL Choi)Fig.M.7.4.2.1. Histological section through the ovary of a Pacific oyster (Crassostrea gigas)with normal ova (white arrows) and ova severely infected by the protistan parasite Marteiliodeschungmuensis (black arrows). Scale bar 100µm (MS Park)SECTION 4 CRUSTACEAN DISEASESSECTION 4.1 GENERAL TECHNIQUESFig.C.1.1.1.3a. Behaviour observation of shrimp PL in a bowl (P Chanratchakool)Fig.C.1.1.1.3b. Light coloured shrimp with full guts from a pond with healthy phytoplankton(P Chanratchakool)Fig.C.1.1.2.1a. Black discoloration of damaged appendages (P Chanratchakool)Fig.C.1.1.2.1b. Swollen tail due to localized bacterial infection (P Chanratchakool)Fig.C.1.1.2.2a,b. Shrimp with persistent soft shell (P Chanratchakool/MG Bondad-Reantaso)Fig.C.1.1.2.3a. Abnormal blue and red discoloration (P Chanratchakool)Fig.C.1.1.2.3b. Red discoloration of swollen appendage (P Chanratchakool)Fig.C.1.1.3a. Severe fouling on the gills (P Chanratchakool)Fig.C.1.1.3b. Brown discolouration of the gills (P Chanratchakool)Fig.C.1.1.3c. Shrimp on left side with small hepatopancreas (P Chanratchakool)Fig.C.1.2a, b, c. Examples of different kinds of plankton blooms (a- yellow/green colouredbloom; b- brown coloured bloom; c- blue-green coloured bloom (P Chanratchakool)Fig.C.1.2d. Dead phytoplankton (P Chanratchakool)Fig. C.1.3.6. Points of injection of fixative (V Alday de Graindorge and TW Flegel)SECTION C.2 YELLOWHEAD DISEASE (YHD)Fig.C.2.2. Gross sign of yellow head disease (YHD) are displayed by the three Penaeusmonodon on the left (TW Flegel)Fig.C.2.3.1.4a,b. Histological section of the lymphoid organ of a juvenile P. monodon withsevere acute YHD at low and high magnification. A generalized, diffuse necrosis of LO cells isshown. Affected cells display pyknotic and karyorrhectic nuclei. Single or multiple perinuclearinclusion bodies, that range from pale to darkly basophilic, are apparent in some affected cells(arrows). This marked necrosis in acute YHD distinguishes YHD from infections due to Taurasyndrome virus,which produces similar cytopathology in other target tissues but not in the LO.Mayer-Bennett H&E. 525x and 1700x magnifications, respectively (DV Lightner)234

List Of FiguresFig.C.2.3.1.4c. Histological section of the gills from a juvenile P. monodon with YHD. Ageneralized diffuse necrosis of cells in the gill lamellae is shown, and affected cells displaypyknotic and karyorrhectic nuclei (arrows). A few large conspicuous, generally spherical cellswith basophilic cytoplasm are present in the section.These cells may be immature hemocytes,released prematurely in response to a YHV-induced hemocytopenia. Mayer-Bennett H&E.1000x magnification (DV Lightner)SECTION C.3 INFECTIOUS HYPODERMAL AND HAEMATOPOIETIC NECROSIS (IHHN)Fig.C.3.2a. A small juvenile Penaeus stylirostris showing gross signs of acute IHHN disease.Visible through the cuticle, especially on the abdomen, are multifocal white to buff coloredlesions in the cuticular epithelium or subcutis (arrows). While such lesions are common in P.stylirostris with acute terminal IHHN disease, they are not pathognomonic for IHHN disease(DV Lightner)Fig.C.3.2b. Dorsal view of juvenile P. vannamei (preserved in Davidson’s AFA) showing grosssigns of IHHNV-caused RDS. Cuticular abnormalities of the sixth abdominal segment and tailfan are illustrated (DV Lightner)Fig.C.3.2c. Lateral view of juvenile P. vannamei (preserved in Davidson’s AFA) showing grosssigns of IHHNV-caused RDS. Cuticular abnormalities of the sixth abdominal segment and tailfan are illustrated (DV Lightner)Fig.C.3.4.1.2a. A low magnification photomicrograph (LM) of an H&E stained section of ajuvenile P. stylirostris with severe acute IHHN disease. This section is through the cuticularepithelium and subcuticular connective tissues just dorsal and posterior to the heart. Numerousnecrotic cells with pyknotic nuclei or with pathognomonic eosinophilic intranuclearinclusion bodies (Cowdry type A) are present (arrows). Mayer-Bennett H&E. 830x magnification(DV Lightner)Fig.C.3.4.1.2b. A high magnification of gills showing eosinophilic intranuclear inclusions(Cowdry type A inclusions or CAIs) that are pathognomonic for IHHNV infections. Mayer-Bennett H&E. 1800x magnification (DV Lightner)SECTION C.4 WHITE SPOT DISEASE (WSD)Fig.C.4.2a. A juvenile P. monodon with distinctive white spots of WSD (DV Lightner)Fig.C.4.2b. Carapace from a juvenile P. monodon with WSD. Calcareous deposits on theunderside of the shell account for the white spots (DV Lightner/P. Saibaba)Fig.C.4.3.3.1.2a. Histological section from the stomach of a juvenile P.chinensis infected withWSD. Prominent intranuclear inclusion bodies are abundant in the cuticular epithelium andsubcuticular connective tissue of the organ (arrows) (DV Lightner)Fig.C.4.3.3.1.2b. Section of the gills from a juvenile P. chinensis with WSBV. Infected cellsshow developing and fully developed intranuclear inclusion bodies of WSBV (arrows). Mayer-Bennett H&E. 900x magnification (DV Lightner)SECTION C.4a BACTERIAL WHITE SPOT SYNDROME (BWSS)Fig. C.4a.2. Penaeus monodon dense white spots on the carapace induced by WSD (M. Shariff)Fig. C.4a.4.2.2a, b. Bacterial white spots (BWS), which are less dense than virus-inducedwhite spots. Note some BWS have a distinct whitish marginal ring and maybe with or without apinpoint whitish dot in the center (M. Shariff/ Wang et al. 2000 (DAO 41:9-18))Fig. C.4a.4.2.2c. Presence of large number of bacteria attached to exposed fibrillar laminae ofthe endocuticle (M. Shariff/ Wang et al. 2000 (DAO 41:9-18))SECTION C.5 BACULOVIRAL MIDGUT GLAND NECROSIS (BMGN)Fig.C.5.1.2a. Section of the hepatopancreas of P. plebejus displaying several hepatopancreascells containing BMN-type intranuclear inclusion bodies. Mayer-Bennett H&E. 1700 x magnification(DV Lightner)Fig.C.5.4.2.1a. High magnification of hepatopancreas from a PL of P. monodon with a severeinfection by a BMN-type baculovirus. Most of the hepatopancreas cells display infected nuclei.Mayer-Bennett H&E. 1700x magnification (DV Lightner)235

List Of FiguresFig. C.5.4.2.1b, c. Sections of the hepatopancreas of a PL of P. japonicus with severe BMN.Hepatopancreas tubules are mostly destroyed and the remaining tubule epithelial cells containmarkedly hypertrophied nuclei that contain a single eosinophilic to pale basophilic, irregularlyshaped inclusion body that fills the nucleus. BMNV infected nuclei also display diminishednuclear chromatin, marginated chromatin and absence of occlusion bodies that characterizeinfections by the occluded baculoviruses. Mayer-Bennett H&E. Magnifications: (a) 1300x; (b)1700x (DV Lightner)Fig.C.5.4.2.1d. MBV occlusion bodies which appear as esosinophilic, generally multiple,spherical inclusion bodies in enormously hypertrophied nuclei (arrows). Mayer-Bennett H&E.1700x magnification (DV Lightner)SECTION C.6 GILL-ASSOCIATED VIRUS (GAV)Fig. C.6.4.2.1. Transmission electron microscopy of GAV (P Walker)SECTION C.8 TAURA SYNDROME (TS)Fig. C.8.4.1.1a,b. a. Moribund, juvenile, pond-reared Penaeus vannamei from Ecuador in theperacute phase of Taura Syndrome (TS). Shrimp are lethargic, have soft shells and a distinctred tail fan; b. Higher magnification of tail fan showing reddish discoloration and rough edgesof the cuticular epithelium in the uropods suggestive of focal necrosis at the epithelium ofthose sites (arrows) (DV Lightner)Fig. C.8.4.1.1c,d,e. Juvenile, pond-reared P. vannamei (c – from Ecuador; d – from Texas; e –from Mexico) showing melanized foci mark sites of resolving cuticular epithelium necrosis dueto TSV infection (DV Lightner/F Jimenez)Fig. C.8.4.1.2a. Focal TSV lesions in the gills (arrow). Nuclear pykinosis and karyorrhexis,increased cytoplasmic eosinophilia, and an abundance of variably staining generally sphericalcytoplasmic inclusions are distinguishing characteristics of the lesions. 900x magnification(DV Lightner)Fig. C.8.4.1.2b. Histological section through stomach of juvenile P. vannamei showingprominent areas of necrosis in the cuticular epithelium (large arrow). Adjacent to focal lesionsare normal appearing epithelial cells (small arrows). Mayer-Bennett H&E. 300x magnification(DV Lightner)Fig. C.8.4.1.2c. Higher magnification of Fig. C.8.4.1.2b showing the cytoplasmic inclusionswith pyknotic and karyorrhectic nuclei giving a ‘peppered’ appearance. Mayer-Bennett H&E.900x magnification (DV Lightner)Fig. C.8.4.1.2d. Mid-sagittal section of the lymphoid organ (LO) of an experimentally infectedjuvenile P. vannamei. Interspersed among normal appearing lymphoid organ (LO) cords ortissue, which is characterized by multiple layers of sheath cells around a central hemolymphvessel (small arrow), are accumulations of disorganized LO cells that form LO ‘spheroids”.Lymphoid organs spheres (LOS) lack a central vessel and consists of cells which showkaryomegaly and large prominent cytoplasmic vacuoles and other cytoplasmic inclusions (largearrow). Mayer-Bennett H&E. 300x magnification (DV Lightner)SECTION C.9 NUCLEAR POLYHEDROSIS BACULOVIROSES (NPB)Fig. C.9.3.2.1a. Wet mount of feces from a P. vannamei infected with BP showing tetrahedralocclusion bodies (arrows) which are diagnostic for infection of shrimp’s hepatopancreas ormidgut epithelial cells. Phase contrast, no stain. 700x magnification (DV Lightner)Fig. C.9.3.2.1b,c. Mid and high magnification views of tissue squash preparations of thehepatopancreas (HP) from PL of P. monodon with MBV infections. Most HP cells in both PLsusually display multiple, generally spherical, intranuclear occlusion bodies (arrow) that arediagnostic for MBV. 0.1% malachite green. 700x (b) and 1 700x (c) magnifications(DV Lightner)236

List Of FiguresFig. C.9.3.2.3a,b. a. Mid-magnification view of mid-sagittal sections of PL of P. vannamei withsevere BP infections of the hepatopancreas showing multiple eosinophilic BP tetrahedralocclusion bodies within markedly hypertrophied hepatopnacreas (HP) cell nuclei (arrows).Mayer-Bennett H&E. 700x magnification; b. High magnification of an HP tubule showingseveral BP-infected cells that illustrate well the intranuclear, eosinophilic, tetrahedral occlusionbodies of BP (arrows). Mayer-Bennett H&E. 1800x magnification (DV Lightner)SECTION C.10 NECROTISING HEPATOPANCREATITIS (NH)Fig. C.10.4.1.1. Juvenile P. vannamei with NHP showing markedly atrophied hepatopancreas,reduced to about 50% of its normal volume (DV Lightner)Fig. C.10.4.1.2. Wet- mount of the HP of infected shrimp with inflamed hemocyte, melanizedHP tubules and absence of lipid droplets. No stain. 150x magnification (DV Lightner)Fig. C.10.4.1.3a,b. Low and mid-magnification of photographs of the HP of a severely NHPinfected juvenile P. vannamei. Severe hemocytic inflammation of the intratubular spaces (smallarrow) in response to necrosis, cytolysis and sloughing of HP tubule epithelial cells (largearrow), are among the principal histopathological changes due to NHP. Mayer-Bennett H&E.150x (a) and 300x (b) magnifications (DV Lightner)Fig. C.10.4.1.3c. Low magnification view of the HP of a juvenile P. vannamei with severe,chronic NHP. The HP tubule epithelium is markedly atrophied, resulting in the formation of largeedematous (fluid filled or “watery” areas in the HP. Mayer-Bennett H & E. 100x magnification(DV Lightner)Fig. C.10.4.1.3d. The HP tubule epithelial cells show no cytoplasmic lipid droplets, but insteadcontain masses of the tiny, non-membrane bound, intracytoplasmic NHP bacteria (arrow).Mayer-Bennett H&E. 1700x magnification (DV Lightner)Fig. C.10.4.2.1. Low magnification TEM of a hepatopancreatocyte from a juvenile P. vannameiwith NHP. Profiles of intracellular rod-shaped forms (large arrow) and helical forms (small arrow)of the NHP bacterium are abundant in the cytoplasm. 10 000x magnification (DV Lightner)SECTION C.11 CRAYFISH PLAGUEFig. C.11.3.2.1a. Fresh microscopic mount of a piece of infected exoskeleton showing fungalspores (EAFP/DJ Alderman)Fig. C.11.4.1.1a,b. Clinical signs of infected crayfish showing whitened necrotic musculature inthe tail, and often accompanied in chronic infections by melanisation (blackening) of affectedexoskeleton (EAFP/DJ Alderman)237

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