RISK ANALYSIS – AQUATIC ANIMAL DISEASES ASSOCIATED …

RISK ANALYSIS – AQUATIC ANIMAL DISEASES ASSOCIATED WITH DOMESTIC BAIT TRANSLOCATION

FRDC Project Number 2009/072

RISK ANALYSIS – AQUATIC ANIMAL DISEASES ASSOCIATED WITH DOMESTIC BAIT TRANSLOCATION

Prepared by:

Ben Diggles PhD

DigsFish Services Pty Ltd, Banksia Beach, QLD 4507

Published by DigsFish Services Pty Ltd, (1 July 2011) for FRDC project no. 2009/072

ISBN 978-0-9806995-0-0

Copyright Fisheries Research and Development Corporation and DigsFish Services Pty Ltd 2011

This work is copyright. Except as permitted under the Copyright Act 1968 (Cth), no part of this

publication may be reproduced by any process, electronic or otherwise, without the specific written

permission of the copyright owners. Information may not be stored electronically in any form

whatsoever without such permission.

Disclaimer

The authors do not warrant that the information in this document is free from errors or omissions. The

authors do not accept any form of liability, be it contractual, tortious, or otherwise, for the contents of

this document or for any consequences arising from its use or any reliance placed upon it. The

information, opinions and advice contained in this document may not relate, or be relevant, to a readers

particular circumstances. Opinions expressed by the authors are the individual opinions expressed by

those persons and are not necessarily those of the publisher, research provider, the FRDC or the

Australian Federal Government.

The Fisheries Research and Development Corporation plans, invests in and manages fisheries research and development throughout Australia. It is a statutory authority within the portfolio of the federal Minister for Agriculture, Fisheries and Forestry, jointly funded by the Australian Government and the fishing industry.

_________________________________________________________________________________________________________________________________________________________________________________

FRDC Project No. 2009/072, Final Report July 2011

5

www.digsfish.com

Contents

List of Tables......................................................................................................................................................7

List of Figures ....................................................................................................................................................7

Abbreviations and Acronyms ............................................................................................................................8

Non – technical summary ..................................................................................................................................9

1.0 Introduction.....................................................................................................................12

1.1 Examples of spread of disease agents in bait ......................................................................13 1.1.1 Spread of disease agents in Finfish used as bait ..................................................................13 1.1.2 Spread of disease agents in Crustaceans used as bait...........................................................22 1.1.3 Spread of disease agents in Molluscs used as bait ...............................................................25 1.1.4 Spread of disease agents in Amphibians used as bait ..........................................................27 1.1.5 Spread of disease agents in Annelids used as bait ...............................................................29

2.0 Commodity Description...................................................................................................32

2.1 Finfish...............................................................................................................................34 2.1.1 Salmonids..........................................................................................................................34 2.1.2 Saltwater finfish ................................................................................................................34 2.1.3 Freshwater finfish..............................................................................................................35

2.2 Sharks and rays .................................................................................................................36 2.3 Crustaceans.......................................................................................................................36

2.3.1 Penaeid prawns and Palaemonid shrimp.............................................................................36 2.3.2 Crabs.................................................................................................................................37 2.3.3 Saltwater yabbies/Nippers (Ghost shrimp, Bass Yabbies, Family Callianassidae)................37 2.3.4 Crayfish (freshwater and saltwater)....................................................................................37

2.4 Molluscs............................................................................................................................38 2.4.1 Abalone (and other Gastropods).........................................................................................38 2.4.2 Bivalves (Cockles, mussels, pipis, scallops, oysters)...........................................................39 2.4.3 Cephalopods......................................................................................................................39

2.5 Amphibians .......................................................................................................................40 2.6 Annelids ............................................................................................................................40

2.6.1 Oligochaetes......................................................................................................................41 2.6.2 Polychaetes .......................................................................................................................41

2.7 Other.................................................................................................................................42 2.7.1 Echinoderms......................................................................................................................42 2.7.2 Cunjevoi (Ascidians) .........................................................................................................42

3.0 The methodology used for this Risk Analysis..................................................................47

3.1 Hazard Identification.........................................................................................................47 3.2 Risk assessment .................................................................................................................52

3.2.1 Release assessment............................................................................................................52 3.2.2 Exposure assessment .........................................................................................................53 3.2.3 Consequence assessment....................................................................................................54 3.2.4 Risk estimation..................................................................................................................56 3.2.5 Risk mitigation ..................................................................................................................57

4.0 The Risk Assessment .......................................................................................................58

4.1 Hazard Identification.........................................................................................................58 4.2 Elimination of insignificant diseases ..................................................................................65

4.2.1 Viruses..............................................................................................................................65 4.2.2 Bacteria.............................................................................................................................68 4.2.3 Fungi.................................................................................................................................69 4.2.4 Protozoa ............................................................................................................................69 4.2.5 Metazoa.............................................................................................................................71

4.3 The diseases of concern to be considered in the RA ............................................................71

_________________________________________________________________________________________________________________________________________________________________________________


6

www.digsfish.com

5.0 Detailed Risk Assessment ................................................................................................74

5.1 Infection of finfish and molluscs with Aquatic Birnavirus....................................................74 5.2 Infection of finfish with Epizootic Haematopoietic Necrosis Virus (EHNV) .........................79 5.3 Infection of finfish with Viral Encephalopathy and Retinopathy (Nodavirus).......................84 5.4 Emergence of a new previously unknown virus of finfish.....................................................88 5.5 Infection of finfish with Aeromonas salmonicida (Goldfish ulcer disease, GUD) .................92 5.6 Infection of salmonids with Lactococcus garvieae ..............................................................97 5.7 Infection of salmonids with Piscirickettsia-like bacteria (PLBs)........................................ 101 5.8 Infection of finfish with Yersinia ruckeri (Yersiniosis).......................................................105 5.9 Infection of finfish with Aphanomyces invadans (Epizootic Ulcerative Syndrome)............. 109 5.10 Infection of finfish with Microsporidians.......................................................................... 113 5.11 Systemic amoebic infections of goldfish............................................................................ 117 5.12 Infection of finfish with scuticociliates (including Uronema spp.)...................................... 119 5.13 Infection of finfish with monogeneans............................................................................... 122 5.14 Infection of finfish and molluscs with digeneans ............................................................... 125 5.15 Infection of finfish and crustaceans with nematodes and cestodes..................................... 129 5.16 Infection of finfish with copepods..................................................................................... 134 5.17 Infection of finfish and annelids with myxosporeans ......................................................... 138 5.18 Viral infections of freshwater crayfishes........................................................................... 142 5.19 Infection of prawns with Gill Associated Virus (GAV) ...................................................... 146 5.20 Infection of crustaceans with Hepatopancreatic Parvovirus (HPV)................................... 150 5.21 Infectious Hypodermal and Haematopoietic Necrosis of prawns (IHHNV)........................ 154 5.22 Infection of prawns with Monodon Baculovirus (MBV)..................................................... 158 5.23 Infection of prawns with Mourilyan virus (MoV) .............................................................. 163 5.24 Infection of prawns and crayfish with Spawner Isolated Mortality Virus (SMV) ................ 167 5.25 White Tail Disease of freshwater giant prawns................................................................. 171 5.26 Infection of crustaceans with rickettsia-like organisms (RLOs) ......................................... 175 5.27 Infection of crustaceans with Hematodinium spp. ............................................................. 179 5.28 Infection of crustaceans with Microsporidians ................................................................. 184 5.29 Infection of crabs with Sacculina spp. and other rhizocephalan barnacles ........................ 188 5.30 Viral Ganglioneuritis of Abalone (AVG)........................................................................... 192 5.31 Infection of oysters with Bonamia spp., and/or unidentified microcells.............................. 196 5.32 Infection of molluscs with Haplosporidians...................................................................... 201 5.33 Infection of oysters and annelids with Marteilia sydneyi (QX disease) .............................. 205 5.34 Infection of molluscs with Perkinsus olseni....................................................................... 209 5.35 Infection of molluscs with spionid mudworms................................................................... 213

6.0 Risk Mitigation .............................................................................................................. 218

6.1. Risk Evaluation ............................................................................................................... 218 6.2 Options for Risk Mitigation.............................................................................................. 223

7.0 References...................................................................................................................... 224

Appendix 1. The disease agents known to infect aquatic animals that are used as bait and berley. Data gaps are indicated where they occur. ............................................................................................. 289

Appendix 2. Example of educational posters employed by Michigan Department of Natural Resources as a form of risk mitigation to help control spread of VHS virus in the great lakes region of the USA. ..297

_________________________________________________________________________________________________________________________________________________________________________________


7

www.digsfish.com

List of Tables

Table 1. Recreational bait and berley use in Australia in 2002…………………………………………… 33

Table 2. Commodity List………………………………………………………………………………….. 43

Table 3a. National list of reportable diseases of aquatic animals………………………………………..… 49

Table 3b. State lists of reportable diseases of aquatic animals…………………………………………….. 50

Table 4. Nomenclature for the qualitative likelihood estimations used in this RA……………………….. 53

Table 5. Matrix of rules for combining descriptive likelihoods for the release and exposure assessments.. 54

Table 6. Definition of terms used to describe consequences of establishment of unwanted diseases...…... 55

Table 7. Risk estimation matrix showing the ALOP utilized for this RA………………………….…….... 56

Table 8. List disease agents of those aquatic animals in Australia that are used as bait and/or berley….… 59

Table 9. Disease agents not included in Table 8 because they are unlikely to cause serious disease and/or

are likely to be ubiquitous……………………………………………………………………….. 64

Table 10. The list of diseases of concern to be considered in the detailed risk assessment………………. 72

Table 11. The disease agents identified as requiring additional risk management……………………….. 219

Table 12. Commodities potentially harbouring disease agents that require additional risk management… 221

List of Figures

Figure 1. The potential sources of bait and berley that could spread aquatic animal pathogens in

Australia…………………………………………………………………………………………. 32

Figure 2. Flow chart showing the decision making process used to identify potential hazards in the

hazard identification step……………………………………………………………………….. 48

_________________________________________________________________________________________________________________________________________________________________________________


8

www.digsfish.com

Abbreviations and Acronyms AAHL Australian Animal Health Laboratory ALOP Appropriate level of protection AVG Abalone Viral Ganglioneuritis BMNV Baculoviral Midgut Gland Necrosis Virus BP Baculovirus penaei CdBV Cherax destructor bacilliform virus CdSPV Cherax destructor systemic parvo-like virus cfu colony forming units CqBV Cherax quadricarinatus bacilliform virus CqPV Cherax quadricarinatus parvo virus CGV Giardiavirus-like virus of Cherax quadricarinatus CLO Chlamydia-like organisms EHNV Epizootic Haematopoietic Necrosis Virus EUS Epizootic Ulcerative Syndrome GAV Gill Associated Virus GUD Goldfish Ulcer Disease HPV Hepatopancreatic Parvovirosis IHHNV Infectious Hypodermal and Haematopoietic Necrosis Virus IP intraperitoneal IPNV Infections Pancreatic Necrosis Virus LPV Lymphoid Parvo-like Virus MBV Monodon Baculovirus MCMS Mid Crop Mortality Syndrome MHD-SL Milky haemolymph disease of spiny lobsters MoV Mourilyan Virosis MrNV Macrobrachium rosenbergii nodavirus NACA Network of Aquaculture Centres in Asia-Pacific ng nanograms NSW New South Wales NT Northern Territory OIE Office International des Epizooties, the world organisation for animal health OMVD Oncorhynchus masou virus disease OVVD Oyster velar virus disease PCR Polymerase chain reaction PHV Pilchard herpesvirus PL Postlarvae PLB Piscirickettsia-like bacteria PPT Parts per thousand QLD Queensland qPCR Quantitative PCR QX Infection by Marteilia sydneyi RA Risk Analysis RLO Rickettsia-like organism RT-PCR Reverse Transcriptase PCR SA South Australia SBV Scylla baculovirus SMV Spawner-isolated Mortality Virus TAB Tasmanian aquabirnavirus TCID50 Tissue culture infectious dose 50% endpoint USA United States of America VER Viral Encephalopathy and Retinopathy (Nodavirus infection) WA Western Australia WTD White tail disease of freshwater prawns YHLV Yellow Head Like Viruses YHV Yellow Head Virus

_________________________________________________________________________________________________________________________________________________________________________________


9

www.digsfish.com

Non – technical summary

Empirical evidence both from within Australia and overseas demonstrates a direct pathway exists for the

introduction and establishment of aquatic animal pathogens through the translocation of bait and/or berley

during the process of fishing. Viruses, bacteria, and some protozoans are hazards because they often remain

viable after freezing, while most metazoan parasites generally present a lower risk, though a risk remains for

all types of disease agents if live bait is used. Once viable pathogens have been introduced directly into the

environment as bait or berley, the probability that they will infect aquatic organisms and become established

is determined by a number of factors including the presence of suitable hosts, the number of viable infective

stages, and environmental conditions at the fishing site.

This Risk Analysis (RA) examined the risks of disease introduction associated with domestic translocation of

bait and berley products within Australia. Information on the types of commodities used in commercial and

recreational fisheries is generally available, however details of the quantity of each commodity used is scarce

as this information is not captured by State fisheries departments. The list of species most widely used as

bait or berley included representatives from 17 families of saltwater fishes, 16 families of freshwater fishes,

16 species of prawns, 13 species of crabs or nippers, 13 species of freshwater and saltwater crayfish/lobsters,

at least 32 species of molluscs (gastropods, bivalves, and cephalopods), 23 families or species of annelids, 4

species of echinoderms and 1 ascidean. Hazard identification for the disease agents reported from these

commodities identified at least 80 diseases of potential concern, including 30 viruses, 8 bacterial diseases, 20

protozoan diseases and 21 metazoan diseases from finfish, crustaceans and molluscs, as well as one fungal

disease from finfish. From the preliminary list of 80 potential hazards, 44 disease agents were classified as

diseases of concern that required detailed risk assessment. The 44 diseases of concern were placed into 35

different categories and detailed risk assessments were undertaken. The outcomes of the risk assessments

indicated 21 diseases for which the unmitigated risk exceeded the ALOP (see summary table over page).

Two diseases were classified as high risk, namely EHNV of finfish and AVG of abalone. Three diseases

were classified as moderate risk, including EUS of finfish, and infection of molluscs with Bonamia and

Perkinsus. Sixteen diseases were classified as low risk, including VER of finfish, goldfish ulcer disease,

microsporidian infections of finfish and crustaceans, infections of live finfish, molluscs and crustaceans with

introduced digeneans, nematodes and cestodes, infections of live finfish with introduced copepods and

Caligus epidemicus, infection of finfish and annelids with myxosporeans, viral infections of freshwater

crayfish, GAV, SMV and WTD of prawns, infections of crustaceans with Hematodinium spp. and Sacculina

spp., infections of molluscs with Haplosporidians and infections of molluscs and annelids with Marteilia

sydneyi. Options for mitigation of these risks to within the ALOP are presented.

Data gaps were identified for disease agents of pipis, cockles, callianassids, bait crabs, cephalopod molluscs,

annelids, echinoderms, and ascideans, all of which are commonly used as bait. Active disease surveillance

should be implemented in a structured manner to fill in the data gaps identified in this RA. The importance

of active surveillance was highlighted when a detailed risk analysis was undertaken for a hypothetical

unknown virus from finfish. Indeed, there remains a risk of transfer of unknown disease agents, even in the

absence of their identification, and disease surveillance is the only way to minimize these risks whenever

significant quantities of bait are being translocated to new geographical regions.

_________________________________________________________________________________________________________________________________________________________________________________


10

www.digsfish.com

RA Summary table. Commodities potentially harbouring disease agents that require additional risk

management. �� = high risk, �� = moderate risk, ��= low risk, X = within ALOP.

Commodity type Disease agent requiring risk management

FINFISH EHNV EUS VER GUD Microsporidians Myxosporeans Digeneans cestodes

nematodes copepods+

Live finfish ��

Whole fresh dead finfish

�� X

Frozen whole finfish

�� X �� X X X X

Frozen fish fillets �� X X X X X X

Frozen fish heads �� X �� X X X X

Frozen fish guts/offal

�� X X X X X X

CRUSTACEANS Viruses of FW

crayfish

GAV SMV WTD Microsporidians Hematodinium spp.

Sacculina spp.

Live prawns X �� * �� X X

Live crayfish/ lobsters

�� X �� X �� ? X

Live crabs X X X X ��

Whole fresh dead prawns

X �� * �� X X

Whole fresh dead crayfish / lobsters

�� X X X �� ? X

Whole fresh dead crabs

X X X X �� X

Frozen whole prawns

X X X X X X X

Frozen whole crayfish / lobsters

X X X X X X X

Frozen whole crabs

X X X X X X X

Frozen prawn tails X X X X X X X

Frozen crayfish / lobster tails

X X X X X X X

Frozen prawn heads

X X X X X X X

Frozen crayfish / lobster heads

X X X X X X X

_________________________________________________________________________________________________________________________________________________________________________________


11

www.digsfish.com


MOLLUSCS AVG Perkinsus olseni

Bonamia spp.

Marteilia sydneyi

Haplosporidians Digeneans

Live molluscs X �� X X ��

Live oysters X ��

Live abalone �� X X ��

Whole fresh dead molluscs

X �� X X X X

Whole fresh dead oysters

X �� X X

Whole fresh dead abalone

�� X X X X

Frozen whole molluscs

X �� X X X X

Frozen whole oysters

X �� X X X X

Frozen whole abalone

�� X X X X

Frozen mollusc meat

X �� X X X X

Frozen abalone meat and viscera

�� X X X X

ANNELIDS Marteilia sydneyi

Myxo-sporeans

Live annelids ��

Fresh dead annelids

��

Frozen annelids X X

Freeze dried annelids

X X

+ = introduced species, as well as Caligus epidemicus, * = freshwater prawns (Macrobrachium spp.) only,

? = unknown, as marine crayfish/lobsters in Australia have not been actively surveyed for Hematodinium

spp. at this time.

_________________________________________________________________________________________________________________________________________________________________________________


12

www.digsfish.com

1.0 Introduction

The committees responsible for aquatic animal health within Australia have long recognised the need to

assess the risks associated with translocation of bait and berley within Australia. In 2003, the Primary

Industries Health Committee (PIHC) and Aquatic Animal Health Committee (AAHC) determined that:

1. A risk assessment would provide a basis for determining where resources need to be allocated and where

regulation may be needed. In this regard jurisdictions would need to assess the feasibility of regulation

versus codes of practice.

2. A national approach is needed. Risk assessments should be used as a basis for developing the approach.

Since then the issue has remained as a high priority issue with PIHC and AAHC (and now the Animal Health

Committee (AHC)).

The International Union for Conservation of Nature (IUCN) defines translocation as “the movement of living

organisms from one area with free release in another” (ICUN 1987). Translocation of aquatic organisms

can provide economic benefits, but can also cause many ecological problems (Burrows 2004, Hannon 2008).

There is increasing evidence to suggest that translocation of aquatic animals and their products as bait and

berley represents a significant threat for facilitating introduction and/or range extensions of their disease

agents (Bauer 1991, Stewart 1991, Westman 1991, Ludwig and Leitch 1996, Englund and Eldredge 2001,

Corfield et al. 2007, Pernet 2008). In Australia, the magnitude of that threat is well evidenced by the fact

that the spread of significant disease agents such as the ciliates Ichthyophthirius multifiliis, Chilodonella

hexasticha, Trichodina spp. and C. cyprini, and helminths Gyrodactylus spp. and Bothriocephalus

acheilognathi into native fish populations has followed releases and translocations of both exotic and native

fish (Ashburner 1976, Langdon 1988, 1989a, 1990, Rowland and Ingram 1991, Humphrey 1995a, 1995b,

Dove et al. 1997, Dove 1998, 2000, Dove and Ernst 1998, Dove and Fletcher 2000, Dove and O’Donoghue

2005).

The national recreational and indigenous fishing survey (Henry and Lyle 2003) found that bait was used in

over 80% of the 19.7 million line fishing events estimated to have occurred in Australia in 2000-2001, with

recreational fishers using around 11.5 million baitfish (excluding prawns, cephalopods and other molluscs) in

this time. With release or disposal of unused baitfish at the fishing destination being a common behaviour

(Litvak and Mandrak 1993, Ludwig and Leitch 1996, Lindgren 2006), this suggests that the volume of bait

being translocated during recreational fishing in Australia is significant. Furthermore, the quantity of bait

used in some commercial fisheries (e.g. longline fisheries, lobster trap fisheries) is even higher again. For

example, until recently over 10,000 tonnes and up to 20,000 tonnes of bait were used each year in the

Western Rock Lobster Fishery (Jones and Gibson 1997, Diggles 2007). While a proportion of the bait used

in large commercial fisheries such as the Western Rock Lobster Fishery originates from overseas sources,

this Risk Analysis (RA) will not assess risks posed by international movements of aquatic animals and their

products, as these Import Risk Analyses (IRAs) are the responsibility of Biosecurity Australia (e.g.

Biosecurity Australia 2006, 2009). Instead, the current project will only assess risks associated with

domestic movements of bait and berley used in both commercial and recreational fisheries. Risks associated

_________________________________________________________________________________________________________________________________________________________________________________


13

www.digsfish.com

with movements of ornamental species in the aquarium industry and aquatic animal products used as feed in

the aquaculture industry are also outside the scope of this document.

Movement of live animals has been the main focus for previous studies that have examined the disease risks

associated with their translocation as bait (Bauer 1991, Stewart 1991, Westman 1991). More recently,

however, the potential for spread of disease through use of fresh and frozen aquatic animal products as bait

has also been recognized by several Government authorities1. For example, the emergence and spread of the

abalone ganglioneuritis disease in Victoria (Hooper et al. 2007) has lead to bans on the use of abalone

viscera and other products as bait in adjacent states 2,3,4 due to the likely (though as yet unquantified) risks of

translocating the pathogen via this route.

Once introduced, the spread of diseases in wild fisheries is usually irreversible and can have significant

ongoing economic and ecological implications for not only commercial and recreational fisheries and the

aquaculture industry, but also for biodiversity and conservation of threatened native aquatic species (Dove et

al. 1997, Dove 1998, Dove and Fletcher 2000). The consequences of introduction of diseases to new

geographic areas can be adverse, severe and far reaching, both spatially and temporally (Durand et al. 2000).

1.1 Examples of spread of disease agents in bait

The first step in this Risk Assessment (RA) will be to review previous examples of spread of disease agents

which have originated from, or been facilitated by, movements of aquatic animal commodities used as bait or

berley. Although there are many reports linking movements of bait or berley to the emergence of diseases in

aquatic animals, it is likely that the incidence of bait-related disease translocation is significantly under-

reported. This is because of the difficulty of proving the connection after the event, particularly in instances

where samples of bait for testing have long since disappeared.

1.1.1 Spread of disease agents in Finfish used as bait

Finfish are very popular bait species for both recreational and commercial fishing. They are used to bait

hooks and traps and are used live and dead in fisheries targeting both finfish and crustaceans. There are

many instances where diseases of finfish have been translocated, or suspected to have been translocated, with

movements of live fish, dead fish or frozen fish products for bait. Some of these are reviewed below.

Viruses

Goodwin et al (2004) reviewed potential hazards associated with viruses that may be present in cultured and

wild caught baitfish in the United States. They found that the freshwater baitfish industry in the United States

shipped more than 10 billion fish per year, with more than 80% of all the baitfish sold being farm raised

(Goodwin et al. 2004). They considered that translocations of live baitfish were likely to pose significant

risks for spread of the notifiable pathogens Spring Viraemia of Carp (SVCV) and Viral Haemorrhagic

1 http://www.dpi.nsw.gov.au/fisheries/recreational/info/abalone-disease-closure 2 http://www.dpi.nsw.gov.au/__data/assets/pdf_file/0015/200850/Fishing-closure-using-berley-as-bait.pdf 3 http://www.dpiw.tas.gov.au/inter.nsf/Attachments/SCAN-6ZW2B3?open 4 http://www.pir.sa.gov.au/__data/assets/pdf_file/0018/61452/abalone_virus_brochure_jan08.pdf

_________________________________________________________________________________________________________________________________________________________________________________


14

www.digsfish.com

Septicaemia (VHSV), as well as other less pathogenic agents such as aquareoviruses like Golder Shiner

Virus (GSV) (Goodwin et al. 2004). They also concluded that fish viruses may routinely survive in frozen

baitfish and thus may be disseminated to different watersheds by anglers (Goodwin et al. 2004). While

Goodwin et al. (2004) did not provide any specific examples of where viruses were proven to be spread with

movements of baitfish, they highlighted risks that manifested very soon after their paper was published in the

form of spread of VHSV throughout the Great Lakes region of the United States. The situation with VHSV,

as well as a number of other viruses that have been spread via movements of baitfish, are summarised below.

Viral Haemorrhagic Septicaemia Virus (VHSV)

In 2003, a novel sublineage of VHSV emerged in the Laurentian Great Lakes basin of the United States

causing fish kills in a wide range of freshwater fishes (Elsayed et al. 2006, USDA 2006, Gagne´ et al. 2007,

Groocock et al. 2007, Lumsden et al. 2007, Gustafson 2009, Kim and Faisal 2010). The main species

affected included freshwater drum (Aplodinotus grunniens), muskellunge (Esox masquinongy), Gizzard shad

(Dorosoma cepedianum) and Yellow perch (Perca flavescens) (Elsayed et al. 2006, Lumsden et al. 2007,

Gagne´ et al. 2007). The disease may have originally been introduced into the lakes via natural fish

movements (Groocock et al. 2007), or release of ballast water from shipping (Wisconsin Aquaculture

Association 2007, Bain et al. 2010), but once introduced the disease continued to spread via natural

movements of water and fishes. However, movement of the virus into some inland lakes and reservoirs

isolated from the Great Lakes was also observed (Wisconsin Aquaculture Association 2007, Gustafson

2009), and translocations of live baitfish were considered to be one of the highest risk activities which

contributed to such introductions (VHSV Expert Panel and Working Group 2010). Subsequent efforts at

limiting the spread of the disease in the region have implemented controls on movements of not only live

baitfish, but all finfish commodities that could become infected, as well as disinfection of boats and other

equipment (Michigan Department of Natural Resources 2009, Gustafson 2009, Appendix 2).

Spring Viraemia of Carp Virus (SVCV)

Goodwin et al. (2004) raised questions regarding the safety of translocating live freshwater baitfish after the

discovery of spring Viraemia of carp virus (SVCV) in the United States. In the spring of 2002, SVCV was

discovered in koi (Cyprinus carpio) grown by a major koi dealer in North Carolina who had over 200

commercial koi ponds in both North Carolina and Wisconsin (Goodwin 2002, Shivappa et al. 2008). At

about the same time, SVCV was reported causing mortalities in common carp (Cyprinus carpio) in several

lakes and rivers in Wisconsin including the Mississippi River (Marcquenski et al. 2003). SVCV infects a

broad range of fish species (primarily cyprinids), hence the discovery of the virus in the United States caused

state wildlife agencies to become concerned about the potential of baitfish to move the disease (Goodwin et

al. 2004). This was because the virus was found in several water bodies that were not naturally

interconnected (Marcquenski et al. 2003), suggesting introductions of the virus followed translocations of

fish. Furthermore, wild cyprinid baitfish, including fathead minnows (Pimephales promelas), common carp,

and native cyprinids were at the time being captured and sold interstate from regions in Wisconsin that had

been definitively identified as SVC virus positive (Goodwin et al. 2004). While there appear to be no

scientifically confirmed cases of translocation of SVCV in the USA via baitfish, the virus causes a highly

contagious disease in cyprinids including yearling common carp and fathead minnows, which are both

popular live bait species in the USA. Restrictions on movements of various live bait species and prohibition

of use of carp as bait have been implemented in several areas of the USA in recognition of the risk that

_________________________________________________________________________________________________________________________________________________________________________________


15

www.digsfish.com

baitfish pose for spread of disease following the outbreaks of VHSV and SVCV in that country (New York

State Department of Environmental Conservation 2009).

Epizootic Haematopoietic Necrosis Virus (EHNV)

In November and December 1984 a new iridovirus was discovered during investigations of epizootics in

juvenile redfin perch (Perca fluviatilis) populations in an artificial impoundment (Lake Nillahcootie) in

North East Victoria (Langdon et al. 1986). More outbreaks of the disease were subsequently observed in

Lakes Mokoan, Hume and Eildon (north east Victoria) and Blowering Reservoir (NSW) in the spring and

summer of 1986/87 (Langdon and Humpheries 1987), followed by further reports in other areas of Victoria,

NSW, the ACT and South Australia over the next decade (Whittington et al. 1996, 1999). The virus was

named epizootic haematopoietic necrosis virus (EHNV) due to the epizootic nature of the disease in redfin

and the hepatic and renal haematopoietic necrosis that was found in diseased fish (Langdon and Humphrey

1987, Langdon 1989b). It was considered that the spread of the disease was highly likely to have been

associated with movements of redfin perch (Perca fluviatilis) and possibly rainbow trout (Oncorhynchus

mykiss) around southern Australia through stocking for angling and the use of juvenile redfin perch as bait

(Langdon and Humphrey 1987, Langdon 1989a, 1990, Lintermans et al. 1990, Whittington et al. 1996,

1999). Instances where EHNV has been detected in introduced populations of redfin perch in isolated water

bodies such as fish farm ponds, farm dams (Langdon and Humphrey 1987), the Mount Bold Reservoir in

South Australia (Pierce et al. 1991), and in Lake Burley Griffin in Canberra (Lintermans 2009), together with

its discontinuous distribution (Whittington et al. 2010), provide further evidence to suggest the virus has

been spread to new areas with movements of live or frozen redfin (Whittington et al. 2010).

EHNV has been shown experimentally to cause disease in several native Australian fish species including

Macquarie perch (Macquaria australasica), mountain galaxias (Galaxias olidus) and silver perch (Bidyanus

bidyanus), and Langdon (1989b) postulated that the declines in these three species were due in part to the

introduction of the virus in previously free water catchments with redfin used as bait and/or for illegal

restocking (Langdon 1989a, 1990).

Largemouth Bass virus (LMBV)

Largemouth Bass virus (LMBV), also known as Santee-Cooper ranavirus, is the only virus known to cause

lethal disease in wild largemouth bass (Micropterus salmoides) in the USA (Grizzle and Brunner 2003).

LMBV is an iridovirus that was first isolated in 1991 in clinically normal largemouth bass in Florida (Grizzle

et al. 2002), however the first fish kill attributed to LMBV occurred in the summer and autumn of 1995 when

at least 1,000 largemouth bass died in a reservoir in South Carolina (Plumb et al. 1996, Grizzel et al. 2002).

Since then the virus has been detected in kills of largemouth bass from several states in the south eastern

USA, and has spread as far north as New York State (Groocock et al. 2008) and as far west as Texas

(Southard et al. 2009).

There is epidemiological evidence that suggests that LMBV was a recent introduction into the USA (Grizzle

and Brunner 2003). Viruses very similar to largemouth bass virus were isolated from aquarium fish,

including cleaner fish Labroides dimidiatus and guppy, Poecilia reticulata, that were imported to the USA

from south east Asia (Hedrick & McDowell 1995; Mao et al. 1997, 1999a). Given the lack of host specificity

of most iridoviruses it is possible that largemouth bass were first infected with LMBV by contact with

_________________________________________________________________________________________________________________________________________________________________________________


16

www.digsfish.com

ornamental fish (Mao et al. 1999a,b), which are often reared outdoors in Florida in the region where LMBV

first emerged. Transmission also occurs via the oral route (Woodland et al. 2002a), suggesting LMBV can

be spread via natural predation and by using infected fish as bait. Because LMBV can be carried by

clinically healthy fish of several species of the sunfish family (e.g. bluegill Lepomis macrochirus) that are

recognized as an effective live bait for largemouth bass, this virus was probably spread geographically by the

transport of water or fish in live wells of fishing boats and/or by private or government stocking of fish

(Woodland et al. 2002b, Grizzle and Brunner 2003, Southard et al. 2009).

Pilchard herpesvirus (PHV)

In 1995 and 1998-99 two epizootics occurred throughout Australian populations of pilchards (Sardinops

sagax neopilchardus) due to infection by a novel herpesvirus (Whittington et al. 1997, Hyatt et al. 1997,

Gaughan et al. 2000). In both events, a mortality front commenced near Port Lincoln in South Australia and

spread bi-directionally both eastwards and westwards over 6000 km of coastline throughout the range of the

Australian pilchard population (Fletcher et al. 1997, Gaughan 2002, Murray et al. 2003, Whittington et al.

2008). Exposure of a naïve population to herpesvirus carried by frozen pilchards used as bait or aquaculture

feed is considered by many scientists and epidemiologists to be the cause of both these events (Hine and

MacDiarmid 1997, Gaughan 2002, Murray et al. 2003, Whittington et al. 2008). In 1995 the same

herpesvirus was associated with mass mortalities of pilchards in New Zealand (Hine 1995, Smith et al.

1996b, Whittington et al. 1997, Fletcher et al. 1997). The herpesvirus was introduced into New Zealand in

1995 via infected frozen pilchards used by recreational and commercial fishers as bait after a shipment of

infected pilchards was received from Bremer Bay, Western Australia (Hine 1995, Fletcher et al. 1997,

Crockford 2007, P.M Hine, personal communication). Indeed, it is also notable that during the 1995

mortality event in New Zealand, recreational and commercial fishers were observed collecting large numbers

of clinically diseased, moribund and dead pilchards for later use as bait (Smith et al. 1996b). The fact that

mortalities due to herpesvirus were not recorded in NZ pilchards during the 1998-99 event is likely to be due

to the immediate implementation of a temporary ban on movements of frozen pilchards from Australia to

New Zealand during the entire course of the second event (P.M. Hine and B.K Diggles, personal

observation).

Bacteria

All fish have a “normal” bacterial flora that changes seasonally (Bisset 1948) and which is moved with the

fish whenever the host is translocated. There are also facultative bacterial pathogens such as those in the

Flavobacterium/Cytophaga/Tenacibaculum and Vibrio sp. groups that are considered to be ubiquitous in

aquatic environments (Austin and Austin 2007), but certain strains of which can cause disease and

mortalities in aquatic animals that are stressed, injured and/or exposed to adverse environmental conditions.

However there are also specific bacterial pathogens that are not considered to be ubiquitous and which are

limited in their distribution (Toranzo et al. 2005). The latter can also be translocated with movements of

live, dead and frozen fish, potentially resulting in the spread of undesirable infections to different fish

populations in new geographic areas. Some examples of the translocations of specific bacterial pathogens

with baitfish are reviewed below.

Furunculosis (Aeromonas salmonicida subsp salmonicida)

Ostland et al. (1987) demonstrated the transmission of furunculosis (Aeromonas salmonicida subsp

salmonicida) to salmonids through the use of infected baitfish minnows. Muscle lesions (furuncles) caused

_________________________________________________________________________________________________________________________________________________________________________________


17

www.digsfish.com

by A. salmonicida ss. salmonicida were found in minnows (common shiners Notropis cornutus, white

suckers Catostomus commersoni, creek chub Semotilus atromaculatus and golden shiners Notemogonus

crysoleucas) purchased from a commercial baitfish supplier. The baitfish were then used to transmit the

infection horizontally to coho salmon (Oncorhynchus kisutch) and brook trout (Salvelinus fontinalis) by co-

habitation of the salmon with the clinically diseased minnows in aquaria (Ostland et al. 1987).

Atypical Aeromonas salmonicida

Atypical Aeromonas salmonicida, the cause of goldfish ulcer disease, is thought to have entered Australia

with imported goldfish from Japan and the disease was first detected at a goldfish farm in South Gippsland,

Victoria, in 1975 (Trust et al. 1980). Subsequently, infections were confirmed in wild populations of goldfish

and in goldfish on native fish farms, probably as a consequence of use of live goldfish as bait and of the

disposal of unwanted goldfish into waterways (Whittington et al. 1987, Humphrey and Ashburner 1993). By

1992 the infection was identified in roach with ulcerative lesions from a water impoundment in Victoria

(Humphrey and Ashburner 1993) and in farmed silver perch (Bidyanus bidyanus) with ulcerative dermatitis

(Whittington et al. 1995). Atlantic salmon (Salmo salar) were shown to be extremely vulnerable to the strain

of bacterium from goldfish (Whittington and Cullis 1988) and because of this, restrictions were implemented

on movements of goldfish to Tasmania to protect the developing salmonid industry (Humphrey and

Ashburner 1993). Genomic analysis revealed that isolates from goldfish and silver perch comprised a

homogenous group, suggesting transmission of A. salmonicida across the species barrier from goldfish

(Whittington et al. 1995). Goldfish ulcer disease is now considered endemic in south-eastern Australia,

causing morbidity and mortality in several species of wild, cultured and ornamental fish.

Enteric redmouth (ERM)

Enteric redmouth disease (ERM) is caused by the bacterium Yersinia ruckeri, which was first described in

rainbow trout from the Hagerman Valley, Idaho, USA in the 1950s (Rucker 1966). ERM was reported for

the first time in Europe in March 1981 in rainbow trout farms of the south west of France during outbreaks

of clinical disease in young trout with mortalities between 5 and 10% of the farm stock (Lésel et al., 1983).

A likely source for the introduction of Y. ruckeri was found several years later by Michel et al. (1986), who

described a clinical case of ERM in fathead minnows (Pimephales promelas) that had been imported for

live-bait into France from the United States since 1981. Serotype data show that the European and North

American populations of Y. ruckeri are interrelated (Davies 1990), thus supporting the evidence from Michel

et al. (1986) which suggests that the organism was introduced into Europe from North America by the

importation of infected baitfish.

Fungi

Several types of fungi are considered to be ubiquitous opportunistic saprobes which can overwhelm the

innate immune system and infect aquatic animals that are injured, stressed or immunocompromised by

exposure to suboptimal conditions, such as pollutants or rapid drops in water temperature (Roberts 2001).

Examples include oomycete water moulds such as Saprolegnia, Branchiomyces and Achyla, which are well

known opportunistic invaders of compromised fish and shellfish. However there are a handful of specific

fungal pathogens of fish that are not considered to be ubiquitous in their distribution, such as Aphanomyces

invadans and Ichthyophonus hoferi. It appears that at least one of these pathogens has been spread to new

geographic areas through translocations of live fish, including popular baitfish species, as reviewed below.

_________________________________________________________________________________________________________________________________________________________________________________


18

www.digsfish.com

Epizootic Ulcerative Syndrome (EUS)

Systemic mycoses consistent with epizootic ulcerative syndrome (EUS) caused by the fungus Aphanomyces

invadans were first reported in Australia in the early 1970’s at the earlier stages of a regional epizootic

(McKenzie and Hall 1976, Fraser et al. 1992, Arthur and Subasinghe 2002). Subsequent studies of this agent

throughout Australia and Asia suggest that the emergence of EUS represented a panzootic caused by the

spread of a single clone of A. invadans throughout the region (Lilley et al. 1997, Roberts 2001).

Aphanomyces invadans is not tolerant of salt and can only be transmitted effectively in brackish and

freshwaters, hence the spread of the disease agent throughout many parts of Australia and Asia has most

likely been through translocation of live fish (Callinan et al. 1995, Lilley et al. 1997). Mullet (Mugil

cephalus) are highly susceptible to EUS (McKenzie and Hall 1976, Fraser et al. 1992, Roberts 2001) and this

species is very popular for use as live and fresh dead bait for a variety of estuarine fish and crab species in

Australia (Ross 1995). Indeed, translocations of EUS affected mullet, whiting (Sillago ciliata) and bream

(Acanthopagrus australis) used for bait into different river systems have been observed in southern QLD and

northern NSW (Diggles, personal observation). In other parts of Australia where EUS has a restricted

distribution, such as in the Northern Territory, restrictions have been placed on translocation of live fish from

affected river basins and estuaries, to attempt to prevent further spread of the disease (Humphrey and Pearce

2004).

Ulcerative dermal lesions in menhaden (Brevoortia tyrannus) in estuaries of Virginia and South Carolina

along the east coast of the USA were originally thought to be due to algal blooms and other disease agents,

but were subsequently confirmed to be due to infection by A. invadans (see Blazer et al. 2002).

Aphanomyces invadans has also been detected in freshwater and low salinity habitats in Florida, where it has

been associated with disease outbreaks in baitfish such as menhaden, mullet (Mugil cephalus), bluegill

(Lepomis macrochirus), shad (Alosa sapidissima), pinfish (Lagodon rhomboides), sportfish such as

largemouth bass, red drum (Sciaenops ocellatus), black drum (Pogonias cromis) and many other fish species

(Sosa et al. 2007). Infections in exotic species introduced into Florida, such as snakehead (Channa sp.), have

also been reported (Saylor et al. 2010). Menhaden are a highly popular baitfish in the USA, and this species

appears to be particularly susceptible to EUS (Kiryu et al. 2003). While natural movements of wild fish are

undoubtedly responsible for spread of the disease to many areas, anglers are known to translocate live

menhaden and other species such as mullet in live wells (D. Olander, editor, Sportfishing Magazine, personal

communication), suggesting that movement of infected bait is another probable mechanism facilitating

spread of EUS in the USA. The discovery of A. invadans in lesions on bluegill from a farm pond in Georgia

and channel catfish Ictalurus punctatus from a farm pond in Louisiana (Blazer et al. 2002), shows that EUS

can be spread to new areas via fish translocation.

Oidtmann et al. (2002) showed how fish (rainbow trout Oncorhynchus mykiss, carp Cyprinus carpio, eel

Anguilla anguilla and perch Perca fluviatilis ) that had eaten cuticle of crayfish (Astacus astacus) infected

with crayfish plague (caused by the fungus Aphanomyces astaci) could act as mechanical vectors for

transmission of the fungus to other crayfish via their faeces. Mortalities of naïve crayfish cohabited with fish

began from 30 days after the fish were force fed A. astaci infected cuticle (Oidtmann et al. 2002). The study

by Oidtmann et al. (2002) therefore suggested that baitfish that have fed on crayfish which had died from

_________________________________________________________________________________________________________________________________________________________________________________


19

www.digsfish.com

crayfish plague can be sources of infection of other crayfish for a period of up to 3 days after their meal,

through excretion of the fungus via their faeces.

Protozoa

Wild fish harbour a wide range of protozoan parasites and these can be spread into new areas whenever live

fish are translocated. Some protozoa have resistant resting spore or multiplicative stages of their lifecycle

that can survive harsh environments such as freezing or desiccation. Many protozoa have direct lifecycles,

and thus can be readily transferred to aquatic animal populations in new geographic areas, particularly those

parasite species with low host specificity (Dove and O’Donoghue 2005).

Microsporidia

Microsporidia are obligate, intracellular parasites that infect arthropods, fish, and mammals (Lom and

Dykova 1992). In fish, microsporidian infections can be widespread in various tissues or concentrated into

cysts that are often grossly visible. The lifecycle is usually direct, but can include an intermediate host

(Vossbrinck et al. 1998). Host specificity varies, but can be low with some microsporidian species affecting

a wide range of hosts (Lom and Dykova 1992). The infective stages are spores which may be released from

skin, faeces and urine of live hosts (Lom et al. 2000), or released after death and decomposition of the host.

Free spores are resistant to external conditions and can remain infective for many months, during which time

they may be ingested by predators or scavengers, while infection can also occur via the per-os route through

predation on infected individuals (Lom and Dykova 1992). The microsporidian parasite Heterosporis sp. has

caused significant disease in populations of wild freshwater fish in North America, as well as reduced

marketability of affected fishes, and there is a strong chance this parasite has been spread by movements of

baitfish (Miller 2009). Heterosporis sp. was first recognized in wild fish in the year 2000 when anglers

fishing the Eagle River chain of lakes in north central Wisconsin found yellow perch (Perca flavescens) with

opaque, white fillets (Sutherland 2002). Since that time, the same parasite has also been found in several

other species of freshwater fishes from a variety of water bodies in Wisconsin, Minnesota, and Lake Ontario

(Sutherland 2002). These findings represented the first documented cases of Heterosporis sp. in freshwater

fish in the Western Hemisphere, and the world’s first report of the parasite occurring in wild fish. Until

2000, this genus of parasites had only been reported in Europe from aquarium species such as angel fish,

bettas and cichlids, and from Asia in the Japanese eel (Wisconsin Department of Natural Resources 2005).

Popular baitfish such as white suckers, golden shiners and fathead minnows were readily infected with

Heterosporis sp. in laboratory trials, and it was well known that these hosts were commonly translocated by

anglers (Miller 2009). Based on these findings, several species of baitfish continue to be monitored for

Heterosporis sp. in Wisconsin as they are recognised as a potential source of Heterosporis sp. transmission

into uninfected waters through normal angling processes which may result in the accidental or intentional

release of baitfish (Sutherland 2002, Wisconsin Department of Natural Resources 2005).

Ciliophora

The ciliate Ichthyophthirius multifiliis is the causative agent responsible for white spot disease in freshwater

fish (Matthews 2005). Ichthyophthirius multifiliis was introduced into Australia and has subsequently

established as a serious pathogen in native and introduced fish (Whittington and Chong 2007). There is

_________________________________________________________________________________________________________________________________________________________________________________


20

www.digsfish.com

evidence that suggests the introduction of ichthyophthiriasis in Tasmania in 1933 was a result of use of

infected goldfish as bait (Parliament of Tasmania 1933).

The Lake Eacham rainbowfish (Melanotaenia eachamensis) was reported to be abundant in Lake Eacham on

the Atherton Tableland, North Queensland, up until the early 1980’s. However between 1984 and 1987

several species of fish were introduced into the lake, including mouth almightly (Glossamia aprion), the

archer fish (Toxotes chatareus), bony bream (Nematalosa erebi), and the banded grunter (Amninataba

percoides) (see Barlow et al. 1987). Several of these species are popular live bait species, and it is possible

the translocation of at least some was as a result of use of bait or a misguided attempt to establish a food

chain in the lake that included bait. After the introduction of the other species, the Lake Eacham rainbowfish

disappeared from the lake (Barlow et al. 1987). While the main reason for the extirpation of the rainbowfish

was probably predation by the mouth almighty, and to a lesser extent, archer fish and the banded grunter,

Barlow et al. (1987) also suggested that diseases and parasites may have also been introduced with the

translocated species. Lake Eacham is a volcanic crater lake with a catchment that is isolated from all other

water courses (Barlow et al. 1987), hence I. multifiliis is one parasite which, due to its almost ubiquitous

distribution, was likely to have been first introduced into the lake with the translocation of the other fish

species. Subsequent research identified that M. eachamensis was indeed highly susceptible to I. multifiliis

compared to other species of rainbowfish (Gleeson et al. 2000), probably due to the population of fish in

Lake Eacham being naïve to I. multifiliis. Surveys of the parasites and other disease agent present in the

current fish fauna of Lake Eacham would be required to confirm whether I. multifiliis and/or other significant

disease agents now occur there.

Metazoa

Wild fish harbour a wide range of metazoan parasites and these can be spread into new areas whenever live

fish are translocated. Some metazoan parasites (digeneans, cestodes, acanthocephalans, myxosporeans) have

complicated multi-host lifecycles (Rohde 1984), which tends to reduce, but not completely eliminate (Bauer

1991, Bartholomew and Reno 2002, Choudhry et al. 2006), the likelihood of successful establishment of the

parasites in new areas. Other metazoan parasites (particularly ectoparasitic monogeneans and crustaceans)

have direct lifecycles, and these can be readily transferred to fish populations in new geographic areas,

particularly those species with low host specificity (Bauer 1991, Kennedy 1993).

Myxosporea

The myxosporean parasite Myxobolus cerebralis infects cartilage of the skeletal system and is the causative

agent of whirling disease in salmonids. First reported in Germany in diseased rainbow trout (Oncorhynchus

mykiss) and brook trout (Salvelinus fontinalis) in the late 19th century, M. cerebralis has since been

documented in temperate freshwater ecosystems around most of the world (Bartholomew and Reno 2002).

The parasite is likely to have originated from European brown trout (Salmo trutta), which are resistant to

whirling disease and are known sub-clinical carriers of M. cerebralis. In contrast, both wild and cultured

salmonids in North America have suffered significant disease outbreaks since the parasite was first

documented in the United States in Pennsylvania in 1956 (Bartholomew and Reno 2002). Myxobolus

cerebralis is thought to have been transported to North America in the 1950s in either live brown trout

imported into hatcheries as broodstock, or in frozen trout products imported from Europe and introduced into

_________________________________________________________________________________________________________________________________________________________________________________


21

www.digsfish.com

local waterways as bait or fishfeed (Nickum 1999, Bartholomew and Reno 2002). Since the original

introduction, M. cerebralis has since spread through much of the United States through stocking of infected

fingerlings to uninfected waterways (Bartholomew and Reno 2002), and angler activities (Budy et al. 2003,

Gates et al. 2008, 2009).

The lifecycle of M. cerebralis requires tubificid oligochaetes as an intermediate host (Markiw and Wolf

1983, Wolf and Markiw 1984). Myxosporean infective stages can be disseminated via translocation of

oligochaete worms (Hallett et al. 2006), and in regions where M. cerebralis has been introduced, sites with

highest angler activity tend to have the highest prevalences of the parasite (Budy et al. 2003). However, the

majority of trout anglers in many affected areas are fly fishers (Gates et al. 2009) and do not use bait,

suggesting that use of oligochaetes and/or infected salmonid products as bait is unlikely to have contributed

to the range expansion of M. cerebralis in the USA in any significant manner. Indeed, transfer of spores or

other infective stages of the parasite via soil or other material lodged in fishing boots, waders or other

angling equipment is a more likely source of unexpected spread of the parasite through angler activity (Gates

et al. 2008, 2009).

Monogeneans

Monogenean ectoparasites parasitise the gills and external body surfaces of fish. They have direct lifecycles

and hence are readily transferred horizontally from fish to fish, however many species have high host

specificity, which tends to reduce the likelihood of their establishment in new areas (Bakke et al. 1992). In

Australia, the monogeneans Dactylogyrus anchoratus and D. extensus have been found in introduced

populations of goldfish and carp (Dove and Ernst 1998). Given that both D. anchoratus and D. extensus

have quite low host specificity, there is a very real possibility that these parasites could be spread to native

fishes (Dove and Ernst 1998), including via use of their original hosts as bait (Lintermans 2004).

The OIE listed monogenean Gyrodactylus salaris also has a broad host range. Peeler and Thrush (2004)

assessed the risks of introduction of Gyrodactylus salaris from Europe into the United Kingdom and

concluded that viable parasites could enter local water bodies via the use of infected salmonid carcasses as

bait. However they also concluded that the overall risk of introduction would be negligible due to the

comparatively small volume of salmonid products that are used in this potential exposure pathway (Peeler

and Thrush 2004).

Digenea

Digeneans are endoparasitic helminths that live in the gastrointestinal tract of fishes and other vertebrates.

Their lifecycle requires a molluscan first intermediate host with plankton eating fishes as final hosts, or

second intermediate hosts in some lifecycles where final hosts include larger fishes, birds and mammals

(including humans). Centrocestus formosanus is a digenetic trematode with a freshwater snail first

intermediate host, various species of fish as second intermediate hosts, and piscivorous birds and mammals

as the final host (Mitchell et al. 2005). The worm encysts as metacercaria in the gills of a wide range of

freshwater fishes, including goldfish, poeciliids and other species used as bait in Australia (Dove 1998, Dove

2000, Evans and Lester 2001, Mitchell et al. 2005). Centrocestus formosanus has been spread to many parts

of the world with the spread of snail intermediate hosts, bird definitive hosts and/or movements of

ornamental fishes (Evans and Lester 2001, Font 2003, Mitchell et al. 2005). Once introduced into a new

_________________________________________________________________________________________________________________________________________________________________________________


22

www.digsfish.com

region there is a very real possibility that these parasites could be spread to different areas, including via use

of their fish intermediate hosts as bait (Lintermans 2004).

Cestodes

Cestodes are endoparasitic helminths that live in the gastrointestinal tract of fishes and other vertebrates.

Their lifecycle generally requires crustaceans (e.g. copepods) as the first intermediate host with plankton

eating fishes as final hosts, or second intermediate hosts in some lifecycles where final hosts include larger

fishes, sharks, birds or mammals (Rohde 1984, Noga 1996). European carp are a common host for the Asian

fish tapeworm Bothriocephalus acheilognathi, a cestode with low host specificity that has been introduced to

many parts of the world from its original distribution in Taiwan through movements of grass carp

(Ctenopharyngodon idella), other cyprinids and poeciliids (Bauer 1991, Kennedy 1993, Font 2003).

Although B. acheilognathi has a complex life cycle, its intermediate hosts are cosmopolitan copepods, thus it

easily infects many plankton-eating fishes, including carp, young catfish and even Gambusia affinis (see

Bauer 1991). The Asian fish tapeworm has been reported in the introduced European carp (Cyprinus

carpio), the western carp gudgeon (Hypseleotris klunzingeri) and several other native fish species in eastern

Australia (Dove et al. 1997, Dove and Fletcher 2000). In the USA, the spread of B. acheilognathi has been

facilitated by the baitfish industry through movements of cyprinid baitfish such as fathead minnows and

golden shiners (Heckmann et al. 1993, Choudhury et al. 2006). Indeed, Heckmann et al. (1993) found

minnows in four bait shops near Las Vegas were infected with B. acheilognathi, while Pullen et al. (2009)

found the mean abundance of B. acheilognathi was 50% greater at public fishing sites compared to other

areas in Kansas. In Australia, it is also possible that use of carp as live or dead bait (Lintermans 2004) has

also helped contribute to the spread of B. acheilognathi that has been observed in Australia (Dove and

Fletcher 2000).

Crustaceans

Crustacean ectoparasites of fish have direct lifecycles and invade the fins, gills, skin and other body cavities

(Kabata 1984). Some species of ectoparasitic crustaceans have low host specificity and these species are

likely to be introduced to new fish populations through translocations of infected fish. For example, the

copepod Lernaea cyprinacea (syn. Lernaea elegans, see Nagasawa et al. 2007) is a common parasite

(“anchor worm”) of the external surfaces of goldfish, but also infects European carp and even amphibians

(Tidd and Shields 1963). These parasites have also been found to infect several species of Australian native

fishes (Rowland and Ingram 1991, Hassan 2008, Lymbery et al. 2010). It is likely that use of carp and

goldfish as live or dead bait in many areas of Australia (Lintermans 2004) has helped to contribute to the

spread of L. cyprinacea into populations of native fishes.

1.1.2 Spread of disease agents in Crustaceans used as bait

Crustaceans are very popular baits for recreational fishing (Kewagama Research 2002, 2007), and are also

used to a lesser extent in commercial fisheries. They are used mainly to bait hooks used in fisheries targeting

finfish. Crustaceans harbour a wide range of viral, bacterial, fungal, protozoan and metazoan disease agents

(Bower et al. 1994). There are several instances where diseases of crustaceans have been translocated, or

suspected to be translocated, with movements of crustaceans for bait. Some of these are reviewed below.

_________________________________________________________________________________________________________________________________________________________________________________


23

www.digsfish.com

Viruses

The Caribbean spiny lobster (Panulirus argus) supports important fisheries throughout the Caribbean, and is

host to the first known pathogenic virus in a lobster, namely Panulirus argus virus 1 (PaV1) (Shields and

Behringer 2004). The virus is particularly prevalent (and also causes the highest mortality rates) in juvenile

lobsters less than 20 mm carapace length (Butler et al. 2008). It appears likely that the commercial practice

of retention of juvenile lobsters inside lobster traps as “social attractants” (i.e. bait) to attract larger lobsters

into the trap results in increased PaV1 transmission rates, and movements of infected lobsters when these

traps are moved helps facilitate the spread of the disease throughout the lobster population (Shields and

Behringer 2004).

Use of penaeid shrimp as bait presents a risk of introduction of viral diseases (Lightner et al. 1997,

Biosecurity Australia 2006, 2009). There is strong circumstantial evidence that penaeid prawns used as bait

introduced white spot syndrome virus (WSSV), an OIE listed pathogen, to the Gulf of Mexico and Texas in

the United States (Hasson et al. 2006). Frozen, uncooked prawns (Metapenaeopsis sp. and Parapenaeopsis

sp.) from China that were being sold as bait in Texas were examined for WSSV after the virus was isolated

from wild prawns from the coasts of Texas. The Chinese bait prawns (Parapenaeopsis sp. only) were found

to contain viable WSSV that was used to successfully infect specific pathogen free white shrimp (Penaeus

vannamei) via intramuscular inoculation, with all of the experimentally infected prawns exhibiting white

spot disease (WSD) and subsequently dying within 72 hours (Hasson et al. 2006). However, P. vannamei

exposed to the WSSV infected bait prawns via per-os exposure did not become diseased, which the authors

attributed to reduced viability of the virus due to repeated freeze-thaw cycles (Hasson et al. 2006). Indeed,

freezing and thawing, as well as other normal processing procedures for commodity shrimp sold for human

consumption, may significantly reduce the risk of disease transfer of Yellowhead Virus (YHV)

(Sritunyalucksana et al. 2010) , which only infects penaeids. However McColl et al. (2004) found that both

WSSV and YHV remained stable and infective after several freeze/thaw cycles.

Flegel (2009) reviewed some of the risks of transfer of viral pathogens with penaeid prawns. His review

found “no published reports in the peer-reviewed, scientific literature of shrimp disease outbreaks in wild or

cultivated shrimp that originated from shrimp packaged and processed for retail sale”. However, Flegel

(2009) did not consider prawns packaged as bait, nor the potential effects of viruses such as WSSV on non-

penaeid crustaceans, such as crabs, crayfish and lobsters. WSSV has a broad host range that includes all

decapod crustaceans (OIE 2010a). It is likely that crabs, crayfish and lobsters will be more susceptible to

transfer of viral diseases such as WSSV from shrimp, because these three groups of scavengers tend to eat

entire shrimp bodies, including viscera, whereas shrimp tend to eat only the appendages (Soto et al. 2001,

Sritunyalucksana et al. 2010). It is important to note that Biosecurity Australia (2009) documented the

detection of WSSV in Darwin in 2000 in mud crabs (Scylla serrata) and prawns (Penaeus monodon) that had

been fed frozen WSSV infected prawns that originated from Indonesia. The Indonesian prawns were

originally part of a larger shipment that was imported and sold as seafood (the box of frozen prawns was

labelled ‘Cocktail Prawns’ and ‘Product of Indonesia’), but a significant amount of the shipment had been

diverted to bait use, and some were used to feed mudcrabs at an aquaculture facility and prawns at a

university in Darwin (Biosecurity Australia 2009). Both facilities were destocked and disinfected and a

subsequent national survey failed to detect WSSV in any of the 3051 crustacean samples examined for the

virus, confirming Australias WSSV free status (Biosecurity Australia 2009).

_________________________________________________________________________________________________________________________________________________________________________________


24

www.digsfish.com

Halder and Ahne (1988) demonstrated that freshwater crayfish (Astacus astacus) could act as a mechanical

vector for infectious pancreatic necrosis virus (IPNV). The virus could be isolated from crayfish which were

infected via injection, per-os or horizontally through the water (Halder and Ahne 1988), though the crayfish

did not become diseased. Rainbow trout (Oncorhynchus mykiss) exposed to effluent water from tanks

containing the IPNV infected crayfish became infected, as did their eggs (Halder and Ahne 1988). Infected

crayfish carried IPNV in their tissues and haemolymph up to one year after their initial exposure to the virus

(Halder and Ahne 1988). Movements of infected crayfish as bait could therefore expose fish in the receiving

waters to the IPN virus (Freeman et al. 2009).

Bacteria

All crustaceans have a “normal” bacterial flora which is moved whenever the host is translocated (Najiah et

al. 2010). There are also facultative bacterial pathogens such as Vibrio sp. that are considered to be

ubiquitous in aquatic environments (Austin and Austin 2007), but certain strains of which can cause disease

and mortalities in aquatic animals that are stressed, injured and/or exposed to adverse environmental

conditions. However there are also specific bacterial pathogens of crustaceans, such as Aerococcus viridans

(causative agent of gaffkemia disease in lobsters) that are not considered ubiquitous (Alderman 1996), and

which are limited in their distribution. The latter could also be translocated with movements of live, dead

and frozen crustaceans, resulting in the spread of undesirable infections to new geographic areas (Alderman

1996). However, this literature review found it difficult to locate previous verified examples of

translocations of specific bacterial pathogens with crustaceans used as bait.

Fungi

The oomycete fungus Aphanomyces astaci causes crayfish plague, a disease characterised by high (usually

100%) mortality in many species of freshwater crayfish native to Europe, Asia and Australia, but has little or

no effect on American crayfish species (Unestam 1969, 1972, 1975). Crayfish plague probably originated

from the United States, where A. astaci is naturally found on native North American crayfish, species such as

Pacifastacus leniusculus (signal crayfish) and Procambarus clarkii (Louisiana swamp crayfish, in which the

fungus generally does not cause disease except in exceptional circumstances (Alderman 1996). However,

once introduced into Europe with infected, sub-clinical signal crayfish, A. astaci was found to be highly

pathogenic to native European freshwater crayfish and the fungus proceeded to destroy populations of native

crayfish throughout Europe (Alderman 1996). In Europe, recreational fishermen are often blamed for

spreading the disease by moving infected crayfish used as bait from one water body to another (Westman

1991, Oidtmann et al. 2002). Large numbers of crayfish, including exotic species, are also spread by anglers

throughout the USA to areas outside their natural distribution via “bait bucket transfer” (Di Stefano et al.

2009). This represents a direct pathway for introduction and establishment into the receiving waters of not

only the crayfish hosts, but also their infectious agents (Di Stefano et al. 2009).

Protozoa

Wild crustaceans harbour a wide range of protozoan parasites (Sprague and Couch 1971, Bower et al. 1994,

Stentiford and Shields 2005) and these can be spread into new areas whenever live crustaceans are

translocated. Some protozoa have resistant resting spore or multiplicative stages of their lifecycle that can

survive harsh environments such as freezing or desiccation (Lom and Dykova 1992). Many protozoa have

direct lifecycles, and thus can be readily transferred to aquatic animal populations in new geographic areas,

particularly those parasite species with low host specificity.

_________________________________________________________________________________________________________________________________________________________________________________


25

www.digsfish.com

In the USA, the microsporidian parasite Nosema nelsoni was found in the muscles of penaeid prawns

(mainly Penaeus duorarum) that were being sold in the bait shrimp fishery in Florida (Woodburn et al.

1957). The prawns infected with N. nelsoni were known as “cotton shrimp” and were at times common in

retail outlets (e.g. infecting 18 of 138 juvenile prawns, giving a prevalence of 13%), where they were

observed not to survive as well as normal shrimp (Woodburn et al. 1957). Woodburn et al. (1957) noted that

trucks carrying live prawns to retailers for on selling to recreational fishers may travel upwards of 270 miles

(434 km) in a day, including translocations from the west to east coast of Florida and interstate. Stentiford

and Shields (2005) noted that some commercial fishing practices (such as “baiting” crab pots with infected

male crabs (Callinectes sapidus) to attract premoult females for the softshell industry) was likely to spread

parasites of the genus Hematodinium throughout the crab population. In Australia, Horwitz (1990) discussed

the need for controls over translocation of live freshwater crayfish, including their use as bait, and suggested

introduction of disease into new areas was one the main risks involved. In, Australia, freshwater crayfish

can carry infections by the microsporidian genera Thelohania (see Jones and Lawrence 2001, Moodie et al.

2003a) and Vairimorpha (see Moodie et al. 2003b). In Western Australia, it is possible that Thelohania sp.

has been spread through the use of yabbies as live bait, and a zoning system has been implemented in an

attempt to minimise the further spread of the parasite (Western Australian Department of Fisheries 2002).

Metazoa

Wild and cultured crustaceans harbour a wide range of metazoan parasites (Bower et al. 1994, Jones and

Lawrence 2001) and these can be spread into new areas whenever live crustaceans are translocated (Pernet et

al. 2008). A few metazoan parasites of crustaceans, such as Temnocephalan trematodes, have low host

specificity and direct lifecycles (Jones and Lester 1993), and thus can readily establish populations in new

geographic areas once their hosts are translocated (Coughran et al. (2009). Callianassids (saltwater yabbies

or ghost shrimp) are a popular bait in Australia, and Pernet et al. (2008) found that translocated live

callianassids may carry parasites such as bopyrid isopods. Other metazoan parasites found in crustaceans are

early developmental stages of helminths that have birds, fish or elasmobranchs as final hosts. One example

of the latter is the larval cestodes, nematodes and metacercaria found by Woodland et al. (1957) in prawns

translocated in the live bait fishery in Florida, USA.

1.1.3 Spread of disease agents in Molluscs used as bait

Molluscs are very popular baits for recreational fishing, particularly cephalopods and bivalves, but also to a

lesser extent other molluscan groups such as gastropods, including abalone (Kewagama Research 2002,

2007). They are used mainly to bait hooks used in fisheries targeting finfish. Molluscs can be infected by a

wide variety of viral, bacterial, protozoan and metazoan disease agents (Bower et al. 1994), and bivalve

molluscs in particular can also accumulate viral and bacterial disease agents of fish and even humans by

filtering these from the water during normal feeding (Elston 1997). While there are many instances of

mollusc diseases being translocated during movements of molluscs for aquaculture (Hine 1996b), there are

relatively few instances recorded in the published literature where diseases of molluscs have been

translocated, or suspected to have been translocated, with movements of molluscs used for bait. Some of

these are reviewed below.

_________________________________________________________________________________________________________________________________________________________________________________


26

www.digsfish.com

Viruses

Molluscs are known to harbour several different types of viruses that can cause disease. These include

oyster velar virus disease of Pacific oysters (Crassostrea gigas) (see Elston 1980), herpes-like viral diseases

of cupped and flat oysters (Arzul et al. 2001, Hine et al. 1998, Friedman et al. 2005), enigmatic viral particles

visualised by electron microscopy in scallops, mussels and clams (Bower et al. 1994, Jones et al. 1996), and

the herpesvirus-like agents that cause abalone viral mortality in China and Taiwan (Chang et al. 2005) and

ganglioneuritis in Australia (Hooper et al. 2007). Movement of infected molluscs as bait is likely to result in

the spread of undesirable viral infections to new geographic areas5, though a literature review found it

difficult to find verified examples of the translocation of viral pathogens with molluscs used as bait.

Bacteria

All molluscs have a “normal” bacterial flora which is moved whenever the host is translocated. There are

also facultative bacterial pathogens such as Vibrio sp. that are considered to be ubiquitous in aquatic

environments (Austin and Austin 2007), but certain strains of which can cause disease and mortalities in

aquatic animals that are stressed, injured and/or exposed to adverse environmental conditions (Lester 1989).

However there are also specific bacterial pathogens of molluscs that are not considered to be ubiquitous and

which are limited in their distribution. An example of a specific bacterial pathogen of molluscs with a limited

distribution is the rickettsiales- like Xenohaliotis californiensis, the causative agent responsible for withering

syndrome in various species of abalone in California (Gardener et al. 1995, Friedman et al. 2000, Moore et

al. 2000). This OIE listed disease agent (OIE 2010a, 2010b) has recently been reported in Thailand,

probably introduced there after translocations of live abalone for aquaculture (Wetchateng et al. 2010).

Movement of infected molluscs as bait is also likely to result in the spread of undesirable bacterial infections

to new geographic areas, though a literature review found it difficult to find verified examples of the

translocation of specific bacterial pathogens with molluscs used as bait.

Fungi

Molluscs are hosts to several species of fungi that can cause significant disease. For example, the fungus

Ostracoblabe implexa causes shell disease in a variety of species of bivalve molluscs in many parts of the

world (Bower et al. 1994). Fungi appear particularly problematic for abalone. Atkinsiella dubia was found

on the mantle of abalone Haliotis sieboldii in Japan (Nakamura and Hatai 1995), as was Atkinsiella awabi,

which caused disease in H. sieboldii stocked into coastal areas in Japan (Kitancharoen et al. 1994).

Haliphthoros milfordensis was also isolated from H. sieboldii (Hatai 1982), as was Halioticida

noduliformans, which was associated with white nodules on the mantle of several species of abalone in Japan

(Muraosa et al. 2009). In New Zealand, paua (Haliotis iris) suffer from shell disease due to infection of the

inner shell by a fungus which can cause reduced growth and increased mortality (Grindley et al. 1998,

Nollens et al. 2003). Movement of molluscs as bait is likely to result in the spread of fungi, such as those

described above, to new geographic areas, though a literature review found it difficult to find verified

examples of the translocation of fungi with molluscs used as bait.

Protozoa

Molluscs are hosts to a variety of protozoan parasites that can cause significant disease. Indeed, besides

withering syndrome of abalone, all other OIE listed disease agents of molluscs are protozoan parasites of the

genera Bonamia, Marteilia and Perkinsus (OIE 2010a, 2010b). Other protozoan parasites of molluscs 5 http://www.pir.sa.gov.au/__data/assets/pdf_file/0018/61452/abalone_virus_brochure_jan08.pdf

_________________________________________________________________________________________________________________________________________________________________________________


27

www.digsfish.com

include Marteilioides spp. (see Itoh 2002), apicomplexans (Hochberg 1983, Hine 2002) and haplosporidians

(Diggles et al. 2002, Hine et al. 2002a, Hine et al. 2009). Haplosporidium nelsoni, cause of MSX disease in

eastern oysters (Crassostrea virginica) along the east coast of the United States (Andrews 1968), was

probably translocated to that region from Japan through imports of live seed oysters (Friedman 1996,

Burreson et al. 2000, Kamaishi and Yoshinaga 2002). Reports of H. nelsoni from Crassostrea gigas in

France (Renault et al. 2000) provide further evidence this parasite has been moved with translocation of

infected oysters, despite the fact H. nelsoni is thought to have an indirect lifecycle that requires an

intermediate host (Ford et al. 2001). Parasites of the genus Marteilia also have an indirect lifecycle which

includes copepods (in the case of Marteilia refringens, see Audemard et al. 2002) or polychaete worms (in

the case of M. sydneyi, see Adlard and Nolan 2008, Cribb 2010). Unlike other haplosporidians, microcells of

the genus Bonamia have a direct lifecycle that mainly infects flat oysters (B. ostreae, B. exitiosa, B.

perspora, see Carnegie et al. 2006), though cupped oysters can also be infected (B. roughleyi, Bonamia sp.

see Cochennec-Laureau et al. 2003, Burreson et al. 2004). Microcell diseases of oysters are increasingly

being detected in new geographic areas (e.g. Abollo et al. 2008, Hill et al. 2010). Other protozoa such as

Perkinsus sp. cause epizootic disease in a wide variety of molluscs, with Perkinsus olseni having an

extremely wide host range (Goggin and Lester 1995, OIE 2010b). Evidence of translocation of protozoan

diseases of molluscs in the past with movements of oysters for aquaculture (Hine 1996b) suggest that

translocations of molluscs as bait is also likely to result in the spread of protozoan disease agents to new

geographic areas. However, a literature review found it difficult to find verified examples of the

translocation of protozoan disease agents with molluscs used as bait.

Metazoa

Molluscs can host a variety of metazoan symbionts, including polychaetes (mudworms), turbellarians, and

crustaceans (copepods, shrimp, crabs), and they can also act as intermediate hosts for helminth parasites

(digeneans, cestodes, nematodes) (Lester 1989, Bower et al. 1994, Nolan and Cribb 2004). Cephalopod

molluscs can also harbour monogeneans as well as being common intermediate hosts for a wide range of

helminth parasites that infect fish, birds and marine mammals (Hochberg 1983). Movement of molluscs as

bait is likely to result in the spread of symbionts and parasites to new geographic areas, though a literature

review found it difficult to find verified examples of the translocation of metazoan endosymbionts with

molluscs used as bait.

1.1.4 Spread of disease agents in Amphibians used as bait

Amphibians are regularly used for bait in several overseas countries (Picco and Collins 2008). Amphibians

can carry a range of disease agents, particularly viruses, bacteria and fungi, but also protozoan and metazoan

parasites (Daszak et al. 1999, 2000, 2003). There are several instances recorded in the published literature

where diseases of amphibians have been translocated, or suspected to have been translocated, with

movements of molluscs used for bait. Some of these are reviewed below.

Viruses

Amphibians are susceptible to infection by ranaviruses within the family Iridoviridae (Whittington et al.

2010), and ranaviral infection of amphibians has recently become listed by the OIE (OIE 2010a, 2010b).

Mass mortalities of amphibians from ranaviruses have been reported on 5 continents (Gray et al. 2009), with

trade in amphibians is likely to be an important route for spread of these disease agents (Picco and Collins

_________________________________________________________________________________________________________________________________________________________________________________


28

www.digsfish.com

2008, Schloegel et al. 2009). Genetic analysis of strains of iridoviruses from amphibians and fish in North

America strongly support the hypothesis that iridoviruses have been spread with the movement of

salamanders (Ambystoma tigrinumas) used as bait (Jancovich et al. 2005, Picco and Collins 2008). About

2.5 million salamander larvae (waterdogs) were sold for bait in the lower Colorado River region, USA alone

as far back as 1968, and during a 7 month long study of the industry, 85% of bait shops sold at least one

iridovirus infected salamander (Picco and Collins 2008). Their study found that all salamanders in the bait

trade were collected from the wild, and in general they were moved east to west and north to south, bringing

with them their multiple ranavirus strains (Picco and Collins 2008). Surveys found that 26-73% of anglers

used live salamanders as fishing bait, 26-67% of anglers released salamanders bought as bait into fishing

waters, and 4% of bait shops released salamanders back into the wild after they were housed in shops with

infected animals (Picco and Collins 2008). Given the scale of this problem, authorities in North America

have moved to reduce the risk of translocation of ranaviruses with amphibians in the bait trade. For

example, in March 2008, the Ontario Ministry of Natural Resources banned the commercial harvest and sale

of all bait frog species in that state6, and banning use of salamanders in Canada (CMNR 2009).

Bacteria

All amphibians have a “normal” bacterial flora which is moved whenever the host is translocated. There are

also facultative bacterial pathogens such as Aeromonas hydrophila that are considered to be ubiquitous in

aquatic environments (Austin and Austin 2007), but certain strains of which can cause disease and

mortalities in aquatic animals that are stressed, injured and/or exposed to adverse environmental conditions.

As for viruses, bacterial disease agents could also be translocated with movements of live, dead and frozen

amphibians, potentially resulting in the spread of undesirable infections to new geographic areas. However,

a literature review found it difficult to find previous examples of the translocations of specific bacterial

pathogens with amphibians used as bait.

Fungi

Infection by the amphibian chytrid fungus (Batrachochytrium dendrobatidis) has been clearly linked to

declines of frog populations in several countries (Daszak et al. 2003, Retallick et al. 2004, Kilpatrick 2010).

Because of this, infection by B. dendrobatidis is considered to be a significant emerging disease of

amphibians (Daszak et al. 2003), and the disease has recently become listed by the OIE (OIE 2010a, 2010b).

There is evidence that B. dendrobatidis has been spread thorough trade of amphibians (Fisher and Garner

2007, Schloegel et al. 2009). One of the likely avenues for its spread in the USA has been use of

salamanders (Ambystoma tigrinum and Necturus maculosus) for bait (Fisher and Garner 2007, Picco and

Collins 2008). Picco and Collins (2008) found a low number of salamanders (A. tigrinum) in 33% of bait

shops surveyed in Arizona were infected with B. dendrobatidis, and, as for ranaviruses, a significant

proportion of anglers used the salamanders for bait, released unused salamanders bought as bait into fishing

waters, and 4% of bait shops also released salamanders back into the wild after they were housed in shops

with infected animals (Picco and Collins 2008).

Protozoa

Wild amphibians harbour a wide range of protozoan parasites, including ciliates, microsporidians,

apicomplexans, trypanosomes and haemogregarines (Levine and Nye 1977, Barta and Desser 1984,

Delvinquier and Freeland 1988, Delvinquier 1989, Lom and Dykova 1992, Paperna and Lainson 1995) and

6 http://www.eco.on.ca/eng/index.php?page=275

_________________________________________________________________________________________________________________________________________________________________________________


29

www.digsfish.com

these can be spread into new areas whenever live amphibians are translocated. However, a literature review

found it difficult to find verified examples of the translocations of protozoan pathogens with amphibians

used as bait.

Metazoa

Wild amphibians harbour a range of metazoan parasites, particularly helminths (Johnston and Simpson 1942,

Barton 1994), but also leeches (Mann and Tyler 1963) and myxosporeans (Levine and Nye 1977). These

parasites can be spread into new areas whenever live amphibians are translocated. However, a literature

review found it difficult to find verified examples of translocations of metazoan pathogens with amphibians

used as bait.

1.1.5 Spread of disease agents in Annelids used as bait

Annelids (Phylum Annelida) are very popular baits for recreational fishing, particularly polychaetes (Class

Polychaeta) for marine species (Kewagama Research 2002, 2007), but oligochaetes (Class Clitella, subclass

Oligochaeta) are also popular baits in freshwater regions. In some parts of the world, leeches (Class Clitella,

subclass Hirudinea) are popular baits for freshwater fishes (Pennuto 1989). Annelids are used almost

entirely to bait hooks used in fisheries targeting finfish. Annelids have been widely translocated throughout

the world (Hendrix et al. 2008) and can be infected by a range of protozoan and metazoan parasites as well

as act as mechanical vectors for several bacterial, fungal and viral agents (Vijayan et al. 2005a, Laoaroon et

al. 2005, Idowu et al. 2006, Hallett et al. 2006, Faisal and Schultz 2010). However, there are relatively few

instances recorded in the published literature where annelids have been implicated with translocation of

disease agents when used as bait. Some of these are reviewed below.

Viruses

Polychaete worms have been shown to be mechanical vectors for White Spot Syndrome Virus (WSSV)

(Vijayan et al. 2005a), which is an OIE listed disease of decapod crustaceans (OIE 2009a, 2010b). Live

polychaete worms of the genus Marphysa, obtained from worm suppliers in India (who collected them from

areas near prawn farms) were found to carry WSSV at high prevalences (up to 75%), and tiger prawn

(Penaeus monodon) broodstock were able to be experimentally infected with WSSV by feeding them the

infected polychaetes (Vijayan et al. 2005a). The widespread use of infected polychaetes as a conditioning

feed for broodstock prawns was probably an important avenue by which WSSV was spread throughout the

aquaculture industry in India (Vijayan et al. 2005a).

Leeches have been shown to act as mechanical vectors for Viral Haemorrhagic Septicaemia virus (VHSV)

(Faisal and Schultz 2009), which is an OIE listed disease of finfish (OIE 2010a, 2010b). VHSV was isolated

from parasitic leeches by both PCR (prevalence 72.5%) and cell culture (prevalence 62.6%), demonstrating

that the virus remained infective, and suggesting that leeches (and perhaps other ectoparasites) could play a

significant role in transmission of VHSV as vectors and/or reservoirs of infection (Faisal and Schulz 2009).

While leeches are popular baits for freshwater fishes in the USA (Pennuto 1989), this literature review found

it difficult to find verified examples of the translocations of viral pathogens of aquatic animals with leeches,

or other annelids, used as bait.

_________________________________________________________________________________________________________________________________________________________________________________


30

www.digsfish.com

Bacteria

All annelids have a “normal” bacterial flora (Idowu et al. 2006) which is moved whenever the host is

translocated. There are also facultative bacterial pathogens such as Vibrio sp. that are considered to be

ubiquitous in aquatic environments (Austin and Austin 2007), but certain strains of which can cause disease

and mortalities in aquatic animals that are stressed, injured and/or exposed to adverse environmental

conditions. The bacterial flora of annelids could be translocated with movements of live, dead and frozen

annelids, possibly resulting in the spread of undesirable infections to new geographic areas. However, this

literature review found it difficult to find previous verified examples of translocations of specific bacterial

pathogens with annelids used as bait.

Fungi

Annelids can host to several species of fungi (Idowu et al. 2006). Movement of annelids as bait is likely to

result in the spread of fungi to new geographic areas, though this literature review did not find verified

examples of the translocation of fungi with annelids used as bait.

Protozoa

The detection of DNA of the protozoan parasites Marteilia refringens and Marteilia sydneyi in sediment

dwelling annelids (polychaetes) suggests that these worms can act as either potential intermediate hosts

(Cribb 2010) or mechanical vectors (Audemard et al. 2002) for parasites of the genus Marteilia. In

Australia, the common baitworm Nephtys australiensis, may act as a true intermediate host for M. sydneyi

(Adlard and Nolan 2008). Spread of these parasites to new geographic areas through movement of annelids

used as bait is therefore theoretically possible, but this literature review did not find verified examples of the

translocation of Marteilia, or other protozoans with annelids used as bait.

Metazoa

Hallett et al. (2006) surveyed several species of live freshwater oligochaetes being sold as “tubifex” worms

in Munich, Germany for infections by myxosporeans. From 7 samples taken over a 1 year period, the water

associated with 5 samples contained actinospores at the time of purchase and spores were subsequently

released in the laboratory in 6 of 7 samples (85.7% prevalence). In all, 12 types of myxosporeans were

isolated from the infected oligochaetes (Hallett et al. 2006). Their results lead Hallett et al. (2006) to

conclude that the sale of worms hundreds of kilometers away from their point of origin was an effective

means for dissemination of myxozoan parasites.

This literature review did not find specific verified examples of movements of metazoan disease agents

through use of annelids as bait, however there is a large amount of evidence that metazoan pests and invasive

species that could act as vectors or intermediate hosts for disease agents have been translocated through use

of annelids as bait. For example, seaweed is often used as packing material to cover live marine polychaete

worms used as bait in the United States. Studies have found that the seaweed can be discarded into the water

at several points along the distribution chain or during the end use by an angler, resulting in the release of

organisms that may be present on the seaweed (Cohen et al. 2001). Crawford (2001) found 35 species of

invertebrates present in wormweed (Ascophyllum nodosum scorpioides) used as shipping material for the

transport of live marine baitworms in the Gulf of Maine. Species present included crabs (Carcinus maenas),

_________________________________________________________________________________________________________________________________________________________________________________


31

www.digsfish.com

9 species of gastropods, 10 species of amphipods, 2 species of bivalves, 2 species of isopods, three types of

insects, and nematodes, trematodes, nemerteans, polychaetes, oligochaetes, copepods and mysids (Crawford

2001). In addition to the baitworms and seaweed used for packing material, Cohen et al. (2001) found 38

species of organisms in baitworm shipments arriving in California. Yarish et al. (2009) examined the

metazoan assemblages that occurred with marine polychaete sand and blood worms sold as bait from retail

shops along the east coast of the United States. They identified translocation of 14 species of macro algae

and two species of harmful microalgae during their survey. In addition, 23 different taxa of invertebrate

animals were discovered in the samples, including five amphipod species, four gastropod species, four

species of bivalve molluscs, three species of annelid, two arachnid species, the larvae of two insect species,

and one species each of isopoda, copepoda and ostracoda (Yarish et al. 2009). Higher summer temperatures

led to a higher diversity and abundance of the phytoplankton species identified, as well as greater abundance

of the invertebrate species. They noted that the bait worms were commonly shipped to other parts of the US,

including the west coast, as well as to Europe (Yarish et al. 2009). While these studies did not focus on the

potential for the translocated invasive species to introduce disease, because the introduced species can act as

vectors or intermediate hosts for many types of disease agents, this demonstrates another pathway through

which disease can theoretically be translocated through use of annelids as bait.

_________________________________________________________________________________________________________________________________________________________________________________


32

www.digsfish.com

2.0 Commodity Description

The next step in the RA is to identify the relevant commodities used for bait and berley in Australia. Bait

and berley are defined for the purposes of this paper as uncooked, live, chilled or frozen animals or parts of

animals used by recreational anglers and commercial fishers to attract fish or crustaceans. Bait and berley in

Australia is derived from a number of sources (Ross 1995, Kewagama Research 2002, 2007), ranging from

invertebrates such as annelids, molluscs, crustaceans and echinoderms, to aquatic vertebrates such as

goldfish, carp and pilchards, to insects and even terrestrial vertebrates such as chicken gut, cattle hide and

hocks, the latter which were used in the past as bait in commercial lobster fisheries (Caputi et al. 2001,

Ghisalberti et al. 2004, Diggles 2007). Use of offal in the Western Australian rock lobster fishery was

banned in the 2001/2002 season (Ghisalberti et al. 2004), and because use of offal from terrestrial animals is

not common (Kewagama Research 2002, 2007), offal from terrestrial vertebrates will not be considered here,

nor will insects or non-animal products that are used as bait or in berley, such as seaweed, bread and dough.

The scope of this RA will be limited to aquatic vertebrate and invertebrate animals used as bait and berley.

Aquatic animals used for bait and berley can be obtained live at the fishing site during the process of fishing.

Alternatively, aquatic animal products can be purchased from bait suppliers or over the internet as live (e.g.

live polychaete worms (Davies et al. 2008, 2010a)), preserved/freeze dried7, chilled, or frozen products

which may be translocated from interstate. Occasionally, live animals from the ornamental fish trade are

used as bait (Lintermans 2004). Figure 1 shows the various sources of bait and berley used in Australia.

Figure 1. The potential sources of bait and berley that could spread aquatic animal pathogens in Australia.

7 www.aquabait.com.au

Aquaculture farm Processors Wild capture

Live animals

Frozen carcases or parts

Chilled carcases or parts Live

animals

Chilled by-products

Frozen by-products

Human food

Diverted to bait

Aquatic environment- marine, freshwater, estuarine

Aquaculture farm: Foodfish, baitfish or ornamental animals

Bait and seafood

processors

Wild capture for food, or

bait

Live animals

Frozen / preserved carcases or parts

Chilled carcases or parts Live or

fresh dead animals

Chilled by-products

Frozen by-products

Human food

Diverted to bait

Aquatic environment- freshwater, estuarine, marine

Use as bait Use as bait Use as bait Use as bait

_________________________________________________________________________________________________________________________________________________________________________________


33

www.digsfish.com

Kewagama Research (2002) undertook a national telephone survey on recreational bait use during May-

August 2002 which targeted a stratified random sample of 8,000 private dwelling households across

Australia. The survey estimated that 1,602,618 households (21.7% of the population) did some kind of

recreational fishing in the previous 12 months. The vast majority (2,479,043, or 86% of all fishers), used

some bait and/or berley during the previous 12 month period (Kewagama Research 2002). Some details of

the bait used by these fishers are included in Table 1 (from Kewagama Research 2002).

Table 1. Recreational bait and berley use in Australia in 2002. Number of fishers using various baits.

Bait type NSW/ACT VIC QLD SA WA TAS NT Total / (%)

Prawns/shrimp 575,540 211,753 442,321 62,625 220,306 18,285 20,889 1,551,721 (62.6%)

Cephalopods 237,719 214,513 288,177 96,601 179,079 32,174 39,593 1,087,856 (43.9%)

Crabs 43,170 23,660 21,701 9,817 10,909 5,320 901 115,478 (4.7%)

S.W. crayfish

/lobsters

0 4,166 0 1,756 2,057 0 0 7,979 (0.3%)

F.W. crayfish 45,982 112,299 46,694 1,558 0 703 0 207,236 (8.4%)

Abalone 5,646 1,334 3,174 0 1,166 2,592 0 13,912 (0.6%)

Other shellfish 115,212 307,173 121,019 153,768 16,591 16,335 901 730,999 (29.5%)

Salmonids 5,124 4,000 0 0 0 0 0 9,125 (0.4%)

Saltwater fish 359,563 294,051 367,351 87,700 247,081 64,341 32,481 1,452,569 (58.6%)

Freshwater fish 7,128 12,108 11,720 0 1,300 0 0 32,257 (1.3%)

Sharks and rays 0 0 9,945 663 4,080 506 0 15,192 (0.6%)

Worms 270,184 107,140 234,931 48,900 12,304 2,752 618 676,828 (27.3%)

Yabbies/nippers 90,394 42,593 230,312 0 0 1,223 1,616 366,139 (14.8%)

Cunjevoi,

urchins, other

34,186 0 6,136 818 1,206 8,388 0 50,734 (2.1%)

_________________________________________________________________________________________________________________________________________________________________________________


34

www.digsfish.com

A follow-up survey conducted in 2006 suggested some temporal changes in bait use patterns had occurred,

most notably increased use of cephalopods and decreased usage of crabs, however the actual extent of the

changes could not be accurately determined from the survey design (Kewagama Research 2007). Indeed, in

recent times, the emergence and wider use of highly effective soft plastic lures may have slightly reduced the

proportion of recreational fishers using bait (B. K. Diggles, personal observation), though no consistent or

significant increases (or decreases) in bait use between 2002 and 2006 were detected by Kewagama Research

(2007).

The following sections will examine the main species groups used for bait in Australia and group them under

the major headings used in the national survey of recreational bait use, in order to remain consistent with

existing data and available statistics as much as possible.

2.1 Finfish

Finfish are widely used as bait throughout Australia, where they are used on hooks as live, fresh dead or cut

baits to attract a wide variety of predatory and scavenging species of fish (Ross 1995). Finfish products are

also very popular as bait for traps and pots set for crabs, freshwater crayfish, and saltwater crayfish

(lobsters). Recreational and commercial fishers will use almost any species they can catch at the fishing

location as live or fresh baitfish under some circumstances, but at other times particular species of baitfish

are targeted (Ross 1995). Alternatively, frozen baitfish (usually saltwater species) are widely available at

commercial fishing co-operatives, processors and thousands of retail outlets (tackle shops, service stations,

supermarkets) throughout Australia. Availability of live finfish for bait at retail outlets in Australia is very

limited, unlike in some other countries such as the USA (Goodwin et al. 2004), though it is known that

ornamental fishes are sometimes purchased for use as bait in freshwater and estuarine areas (Arthington and

Blühdorn 1995, Lintermans 2004).

2.1.1 Salmonids

Kewagama Research (2002) estimated that around 0.4% of respondents in Australia, mainly in NSW/ACT

and Victoria, used salmonids (trout Oncorhynchus mykiss, Salmo trutta or salmon Salmo salar) for bait in

2002. The survey found one angler who used personally caught salmonid fish for live bait, and another who

used trout off-cuts sold as bait at a commercial fish-out facility. Use of salmonid products as bait remained

virtually identical in 2006 (Kewagama Research 2007), when it was calculated that overall around 0.5% of

respondents from NSW/ACT, WA and Tasmania used salmonid products as bait, suggesting their use as bait

is very uncommon in Australia.

2.1.2 Saltwater finfish

Kewagama Research (2002) estimated that around 58.6% of respondents throughout all states used saltwater

finfish as bait or berley in 2002. Use of saltwater finfish as bait was highest in Tasmania (84.5% of

respondents), followed by WA (74.0% of respondents) and the NT (66.9% of respondents). The proportion

of respondents using saltwater finfish as bait remained similar in 2006, when 61% of respondents throughout

_________________________________________________________________________________________________________________________________________________________________________________


35

www.digsfish.com

all states used saltwater finfish as bait (Kewagama Research 2007). Usage rates remained reasonably stable

amongst the states with the top three highest users remaining Tasmania (81.6% of respondents), followed by

WA (78.7% of respondents) and the NT (66.8% of respondents) (Kewagama Research 2007).

Some of the more popular species and groups of saltwater finfish used as bait in Australia in both

commercial and recreational fisheries include Arripidae (Australian salmon, tommy ruff), Atherinidae

(hardyheads, silversides, whitebait), Carangidae (trevallies, jack mackerels, horse mackerels), Clupeidae

(herrings, pilchards), Congoridae (Conger eels), Emmelichthyidae (Redbait, Emmelichthys nitidus),

Engraulidae (anchovies, sprats), Gempylidae (barrcouta), Gerridae (silver biddies), Hemirhamphidae

(garfish), Labridae (wrasse), Leiognathidae (ponyfishes), Lethrinidae (emperors), Lutjanidae (snappers),

Monacanthidae (leatherjackets), Moridae (rock cods), Muglidae (mullet), Nemipteridae (whiptails),

Scombridae (bonitos, mackerels, tunas), tailor (Pomatomus saltatrix), Sillaginidae (whitings), Sparidae

(breams) and many others (Ross 1995).

2.1.3 Freshwater finfish

The use of live freshwater finfish as bait in Australian inland waters has declined in recent years because of

changes in legislation, as it is now illegal in some states or regions (NSW DPI 2010, Northern Territory

Government 2010), although it still may occur illegally. Kewagama Research (2002) estimated that around

1.3% of respondents used freshwater finfish as bait or berley in 2002. Use of freshwater finfish as bait was

highest in Victoria (2.7% of respondents), followed by QLD (1.8% of respondents) and NSW/ACT (1% of

respondents). Freshwater finfish were not recorded as being used for bait in SA, Tasmania and the NT in

2002, suggesting their use in these regions is uncommon. However in 2006, national usage had dropped to

0.7% of respondents, with 3.2% of respondents in the NT using freshwater fish as bait, as did 1.7% in

Victoria and 0.6% in NSW/ACT, with none of the respondents from other states saying they had used

freshwater fish in the previous 12 months (Kewagama Research 2007).

Freshwater finfish are not generally sold as live or frozen bait in retail outlets (except from ornamental

species sold from pet shops), which leaves fishers to catch them at the fishing site. Some of the more

popular species and groups of native freshwater finfish used as live bait in Australia include Ambassidae

(perchlets), archerfish (Toxotes sp.), bony bream (Nematalosa spp.), Eleotridae (gudgeons), Galaxiidae

(galaxids), Hemirhamphidae (garfish), Muglidae (mullets), Melanotaeniidae (rainbowfish), Retropinnidae

(smelts), and Teraponidae (grunters, including banded grunter (Amniataba percoides), and spangled perch

(Leiopotherapon unicolor)) (Ross 1995). There is also evidence that noxious and/or ornamental fishes such

as Cichlidae (tilapias), European carp (Cyprinus carpio), goldfish (Carassius auratus), Poeciliidae

(gambusia and swordtails), oriental weatherloach (Misgurnus anguillicaudatus) and redfin (Perca fluviatilis)

are sometimes used as bait in freshwater and estuarine areas (Arthington and Blühdorn 1995, Lintermans et

al. 1990, Lintermans 2004). Ornamental fishes available for retail sale in Australia are known to have high

prevalence of infection by a range of disease agents (Wickins et al. 2011). Anglers who use live bait are

prone to discard excess fish either into the waterway where they are fishing or into local dams and ponds to

provide bait for subsequent fishing trips (Lintermans 2004).

_________________________________________________________________________________________________________________________________________________________________________________


36

www.digsfish.com

2.2 Sharks and rays

Kewagama Research (2002) estimated that around 0.6% of respondents used products from sharks and rays

as bait or berley in 2002. Use of elasmobranchs as bait was highest in QLD (1.5% of respondents), followed

by WA (1.2% of respondents) and Tasmania (0.7% of respondents). Elasmobranchs were not recorded as

being used for bait in NSW/ACT, Victoria and the NT, suggesting their use in these regions is very

uncommon. In 2006, usage of elasmobranchs as bait dropped to 0.3% overall, with respondents from only

the NT (1.4%) and QLD (1.1%) reporting their use (Kewagama Research 2002), suggesting use of products

from elasmobranchs as bait or berley is very uncommon in Australia.

2.3 Crustaceans

Crustaceans are very popular baits throughout Australia for use to bait hooks used to catch a wide variety of

finfish (Ross 1995). For targeting estuarine and saltwater finfish, the most commonly used crustaceans are

smaller varieties of penaeid prawns (< 13 cm) that will conveniently fit onto hooks of appropriate size

(Kewagama Research 2002, 2007). Penaeid prawns are retailed as either fresh or frozen bait, or as seafood,

in thousands of retail outlets throughout Australia (Kewagama Research 2007). Fresh (green unfrozen as

well as frozen) penaeid prawns are also available from commercial fishing co-operatives and retail outlets

throughout the country. In general, smaller prawns are obtained from bait suppliers, while larger prawns >10

cm are obtained from seafood suppliers (Kewagama Research 2002, 2007). Live penaeid prawns are also

popular baits, particularly with recreational fishers, who generally catch them near their fishing sites using

scoop, bait or cast nets, or with baited traps (Broadhurst et al. 2004). Small live crabs are collected around

sandy areas and/or rocky reefs and coral reefs and are used for targeting a narrower range of finfish species

which commonly prey on small crabs (Ross 1995). Live saltwater yabbies/nippers (Family Callianassidae)

are extremely popular as bait for a variety of estuarine and inshore finfish species targeted by recreational

fishers along the eastern seaboard (Skilleter et al. 2005). Waste products from saltwater crayfish (Palinurid

lobsters) processing are used by commercial and recreational fishers as bait and berley in WA and other

southern states (Horvat 2010). In freshwater areas, penaeid prawns remain a popular bait with recreational

fishers, but other crustaceans such as live palaemonid shrimp and freshwater crayfish are also used (Ross

1995).

2.3.1 Penaeid prawns and Palaemonid shrimp

Kewagama Research (2002) estimated that around 62.6% of respondents throughout all states in the country

used prawns or shrimp as bait or berley in 2002. Use of prawns/shrimp was highest in NSW (78.8% of

respondents), followed by QLD (67.9%), WA (66.8%), Victoria (46.9%), and the NT (43.1%). In 2006, the

total number of respondents that used prawns/shrimp dropped to 58.4%, but the usage rates remained

reasonably stable amongst the states (77% of respondents in NSW, followed by 75.1% of respondents in

QLD, and 69.9% of respondents in WA), with a drop in their use in Victoria (24.6%) and an increase in

Tasmania (41.3%) (Kewagama Research 2007).

Some of the more popular species of penaeid prawns used as bait in Australia include greasyback prawns

(Metapenaeus bennettae), school prawns (Metapenaeus macleayi), and juvenile banana (Penaeus

_________________________________________________________________________________________________________________________________________________________________________________


37

www.digsfish.com

merguiensis, P. indicus), king (P. plebejus, P. latisulcatus) and tiger prawns (P. esculentus, P. semisulcatus,

P. monodon), however all available species of Penaeus, Metapenaeus and other genera such as

Metapenaeopsis and Parapenaeopsis are all likely to be retained and used by anglers as bait provided they

are an appropriate size for the fish species being targeted (Ross 1995).

With regard to freshwater palaemonid shrimp, the giant freshwater prawn (Macrobrachium rosenbergii) is a

popular bait in northern parts of Australia. However, all available species of Macrobrachium, Palaemon and

other genera such as Metapenaeopsis and Parapenaeopsis are likely to be retained and used by anglers as

bait provided they are an appropriate size for the fish species being targeted (Ross 1995).

2.3.2 Crabs


used crabs as bait or berley in 2002. Use of crabs was highest in Tasmania (7% of respondents), followed by

NSW/ACT (5.9%), Victoria and SA (5.2%), then QLD and WA (3.3%). Use of crabs as bait was

significantly less common in 2006 (2.1% of respondents), with usage rates remaining highest in Tasmania

(4.5%) and Victoria (3%), but dropping in NSW/ACT (2.8%), SA (2.3%), WA (1.8%), QLD and NT (0%)

(Kewagama Research 2007).

Some of the more popular species and groups of crabs used as bait in Australia include rock crabs

(Leptograpsus sp., Paragrapsus spp.), bait crabs (Plagusia chabrus, Plagusia sp.), ghost crabs (Ocypode

spp.) and soldier crabs (Mictyris longicarpus, M. platycheles spp.) (see Bennett 1987). The introduced

European green crab (Carcinus maenus) is common in parts of Victoria, NSW, Tasmania, SA and WA and is

probably also used as bait from time to time. Larger blue crabs (Portunus pelagicus) and mud crabs (Scylla

serrata) are occasionally used live by a very small number of specialist anglers targeting large predatory fish

in estuarine and inshore areas.

2.3.3 Saltwater yabbies/Nippers (Ghost shrimp, Bass Yabbies, Family Callianassidae)


except SA and WA used saltwater yabbies/nippers (Ghost shrimp) as bait in 2002. Use of ghost shrimp as

bait was highest in QLD (35.3% of respondents), followed by NSW/ACT (12.4%), and Victoria (9.4%). Use

of ghost shrimp as bait was more popular in 2006 (18.21% of respondents), with usage rates remaining

highest in QLD (38.3%), NSW/ACT (22.9%) and Victoria (8.9%) (Kewagama Research 2007).

The main species of ghost shrimp used as bait in Australia include Callianassa (Trypea) australiensis,

Biffarius arenosus, and Upogebia spp. (Bennett 1987, Ross 1995, Skilleter et al. 2005).

2.3.4 Crayfish (freshwater and saltwater)

Kewagama Research (2002) estimated that around 8.4% of respondents throughout all states except WA and

NT used freshwater crayfish as bait in 2002. Use of freshwater crayfish as bait predominantly occurred in

_________________________________________________________________________________________________________________________________________________________________________________


38

www.digsfish.com

Victoria (24.9% of respondents), followed by QLD (7.2%) and NSW/ACT (6.3%). Use of freshwater

crayfish as bait was more popular in 2006 (15.4% of respondents), with usage rates remaining highest in

Victoria (35.2%), and NSW/ACT (14.8%), and rising to 9% in SA while remaining stable in QLD (6.7%)

(Kewagama Research 2007). Around 7.6% of respondents in the NT recorded using freshwater crayfish as

bait in 2006. No one in Tasmania reported using freshwater crayfish as bait in 2002 (Kewagama Research

2002), rising to 0.9% in 2006, while no one in WA recorded using freshwater crayfish as bait in either 2002

or 2006 (Kewagama Research 2007), suggesting their use as bait is very uncommon in WA and Tasmania.

The main species of freshwater crayfish used as bait includes the yabby (Cherax destructor), redclaw

crayfish (C. quadricarinatus), marron (C. tenuimanus, C. cainii), though it is likely that a wide variety of

other members of the genus Cherax, Engaeus and Euastacus are regularly used (and translocated) as bait

when they are available (Ross 1995, Coughran and Leckie 2007, Coughran et al. 2009).

Very few (0.3% of respondents) used saltwater crayfish (Palinurid lobsters) products as bait or berley in

2002, with little change (0.4%) in 2006 (Kewagama Research 2002, 2007). The fact that only respondents

from NSW/ACT (0.4%) and QLD (1.1%) were found to use saltwater crayfish as bait or berley in 2002, but

not in 2006, whereas respondents from Victoria (0.92%), SA (0.93%) and WA (0.62%) recorded using

saltwater crayfish products in 2006, (Kewagama Research 2002, 2007) suggests their use for this purpose is

sporadic and uncommon in Australia. However, processing waste from the rock lobster industry is

reportedly used by commercial fishers in Western Australia as berley (Horvat 2010).

The main species of saltwater crayfish used as bait and berley are probably the most readily available

species, which would include the southern rock lobster (Jasus edwardsii), eastern rock lobster (Jasus

(Sagmariasus) verreauxi), western rock lobster (Panulirus cygnus) and other Panulirus spp. in tropical

regions (Kailola et al 1993).

2.4 Molluscs

Bivalve molluscs are very popular baits throughout Australia for use to bait hooks to catch a wide variety of

freshwater, estuarine and saltwater finfish (Ross 1995). A large proportion of the live bivalves used as bait is

likely to be gathered by recreational fishers at or near their fishing sites. Frozen bivalves are also sold as bait

in many retail outlets around the country. Live squid, octopus and cuttlefish are also popular baits,

particularly with recreational fishers, who generally catch them near their fishing sites (Ross 1995). Frozen

squid and octopus are also used throughout the country, being sold as bait or seafood in thousands of retail

outlets throughout Australia. Fresh and frozen molluscan waste products (squid, cuttlefish, octopus, abalone,

scallops) originating from commercial fishing co-operatives and processing plants throughout the country are

also used as both bait and berley.

2.4.1 Abalone (and other Gastropods)

Kewagama Research (2002) estimated that around 0.6% of respondents in every state except SA and NT

used abalone as bait in 2002. Use of abalone as bait predominantly occurred in Tasmania (3.4% of

respondents), followed by NSW/ACT (0.8%) and QLD (0.5%). The popularity of using abalone as bait

_________________________________________________________________________________________________________________________________________________________________________________


39

www.digsfish.com

increased in 2006 (1% of respondents), with usage rates remaining stable with the highest in Tasmania

(3.2%), followed by NSW/ACT (1.8%), and QLD (1.1%) (Kewagama Research 2007).

The abalone products used as bait and berley in Australia are likely to be processing waste of those species

most commonly caught in commercial fisheries, namely blacklip abalone (Haliotis rubra), greenlip abalone

(H. laevigata) and Roes abalone (H. roei). Other gastropods are occasionally used as bait in small quantities

by recreational fishers, including limpets (Cellana spp.) (see Sharpe and Keough 1998).

2.4.2 Bivalves (Cockles, mussels, pipis, scallops, oysters)


used “other shellfish” (bivalve molluscs) as bait in 2002. Use of bivalve molluscs as bait was very popular

in SA (81.3% of respondents), followed by Victoria (68%), Tasmania (21.5%) and QLD (18.5%). The use

of bivalve molluscs as bait increased in 2006 (33% of respondents), with usage rates remaining stable with

the highest usage in SA (88.7%), followed by Victoria (55.1%), QLD (26.1%) and Tasmania (20.6%)

(Kewagama Research 2007).

The main species of bivalves used as bait in Australia are likely to be pipis (Plebidonax deltoides, also

known as Goolwa cockles), which are widely used by recreational fishers along the eastern and southern

coastlines (Ross 1995). They are commercially fished in NSW and SA (Kailola et al. 1993) and are usually

available frozen at retail bait shops throughout the southern parts of the country (Davies et al. 2008), though

recent increasing demand for pipis as food has increased prices and reduced their availability as bait in some

locations (Fergurson and Mayfield 2006). There is evidence that some pipis may have been translocated

from the eastern states into SA for certain competitive fishing events due to shortages of bait pipis in SA

(Sunfish 2008).

Venerid clams, or cockles (Katelysia spp., Anadara spp., Barbatia spp.) and razor shells (Pinna bicolour,

Pinna spp., Isognomon spp.) are used as fresh (live) bait in many states (Ross 1995). Due to increased

demand for pipis as food, cockles are becoming more widely available throughout the country as frozen

product, and are now available at most retail outlets. Freshwater mussels (Family Hyriidae) are used as bait

for a variety of freshwater fish and crustacean species, while marine mussels (Mytilus edulis) are popular

baits or saltwater fish species (Ross 1995). Sydney rock oysters (Saccostrea glomerata) and Pacific oysters

(Crassostrea gigas) are very occasionally used as bait by recreational fishers (Ross 1995). Processing waste

from scallops (Amusium spp., Pecten spp., Chalmys spp. Family Pectinidae) (and also, presumably, from

squid, octopus and cuttlefish processing as well) is sometimes used in berley blocks (Horvat 2010)

2.4.3 Cephalopods

Kewagama Research (2002) estimated that around 43.9% of respondents in all states throughout Australia

used cephalopods (squid, octopus and cuttlefish) as bait in 2002. Use of cephalopods as bait was very high

in the NT (81.6% of respondents), followed by WA (54%), SA (51%) and Victoria (47.5%). The use of

cephalopods as bait increased significantly in 2006 (54.7% of respondents), with usage rates remaining

_________________________________________________________________________________________________________________________________________________________________________________


40

www.digsfish.com

highest in the NT (87.2%), followed by a large increase in Tasmania (72.3%), WA (64.7%), NSW/ACT

(60.9%, from 32.6% in 2002), and SA (59.7%) (Kewagama Research 2007).

The main species of cephalopods used as bait in Australia are likely to be various species of squid that are

available frozen at retail bait shops throughout the country. These are likely to include arrow squid (also

known as Goulds squid Notodarus gouldi), other Notodarus spp., Mitre squid (Loligo spp.), southern

calamari (Sepioteuthis australis), northern calamari (S. lessoniana), Sepia spp. and others (Kailola et al.

1993). Octopus species used as bait and berley are likely to include a range of species within the genus

Octopus, including the southern octopus (O. australis), Maori octopus (O. maorum), pale octopus (O.

pallidus), gloomy octopus (O. tetricus), reef octopus (Octopus cyanea) and others (see Bennett 1987, Kailola

et al. 1993, Norman and Reid 2000).

The main species of cuttlefishes (Sepia spp.) used as bait and berley in Australia probably include the

rosecone cuttlefish (Sepia rozella), giant cuttlefish (Sepia apama), and several others that are commonly

caught in commercial fisheries (Kailola et al. 1993, Scandol et al. 2008).

2.5 Amphibians

Amphibians are not commonly used as bait in Australia. In some jurisdictions, it is illegal to use frogs as

bait, including in Queensland (EPA 2007), NSW (NSW DPI 2010), and Victoria (Victoria DPI 2010).

However the protection of frogs is not specifically mentioned in recreational fishing guides in some of these

jurisdictions, such as in Queensland (DEEDI 2010), and hence it is possible that some frogs are used for bait

in freshwater areas of Queensland by anglers not familiar with the regulations. In Tasmania8, Western

Australia9, South Australia10 and the Northern Territory (Northern Territory Government 2010), use of frogs

as bait does not appear to be specifically prohibited, although frogs may be protected under wildlife

protection acts (e.g. WA). Kewagama Research (2002, 2007) did not specifically address the use of

amphibians as bait in their surveys, and indeed specifically excluded collection of amphibians from their

definition of recreational fishing (Kewagama Research (2002, 2007). However it is possible that some

respondents to these surveys considered that amphibians could be included in the definition of “other aquatic

animals”. Respondents who used “other aquatic animals” as bait or burley in the previous 12 months

represented only 2.1% of respondents from 5 states (Tasmania, NSW/ACT, QLD, SA and WA) in the 2002

survey, dropping to 0.5% of respondents in 2006 from Victoria only (Kewagama Research 2002, 2007).

This, together with the jurisdictional arrangements in Victoria and QLD (which are supported by large fines

if infringing anglers are caught), suggests that use of amphibians as bait in Australia either does not occur, or

else is extremely uncommon.

2.6 Annelids

Annelids are very popular baits throughout Australia for use to bait hooks to catch a wide variety of

freshwater, estuarine and saltwater finfish (Ross 1995, Davies et al. 2008). A proportion of the annelids used

8 http://www.ifs.tas.gov.au/ifs/aboutus 9 http://www.fish.wa.gov.au/sec/rec/index.php 10 http://www.pir.sa.gov.au/fisheries/recreational_fishing/recreational_fishing_guide

_________________________________________________________________________________________________________________________________________________________________________________


41

www.digsfish.com

as bait is likely to be gathered by recreational fishers at or near their fishing sites, however collecting worms

for bait generally requires hard work and/or skill (Bennett 1987), which has lead to the development of a

significant retail market for baitworms (Davies et al. 2008). Indeed, Davies et al. (2008) found that around

73% of anglers in SA buy their worms from bait shops. Live, frozen and preserved (including freeze dried)

annelids are widely available from retail bait suppliers throughout the country or over the internet (Davies et

al. 2008, 2010a)11.

Both oligochaetes and polychaetes are used by recreational fishers as bait (Ross 1995). However,

Kewagama Research (2002, 2007) did not discriminate between the two groups in their survey, hence only

overall data on use of annelids as bait is available. Kewagama Research (2002) found that 27.3% of

respondents from all states had used annelids for bait in the previous 12 months. Use of annelids as bait in

2002 was highest in NSW/ACT (37%), followed by QLD (36%), SA (25.9%), and Victoria (23.7%), while

they were rarely used in WA (3.7%), Tasmania (3.6%) and the NT (1.3% of respondents) (Kewagama

Research 2002). The use of annelids as bait remained similar in 2006 (28.8% of respondents), with usage

rates highest in QLD (42.4% of respondents), followed by NSW/ACT (31.6%), Victoria (27.4%) and SA

(23.3%). Usage of annelids remained low in Tasmania (7.7%), WA (3.8%) and the NT (1.6%) (Kewagama

Research 2007). Davies et al. (2008) examined the bait worm industry in SA and found that total production

with the industry ranged between 7 and 14 tonnes between 2000-2001 and 2007-2008, but that only 1.2

tonnes were likely to be sold through the bait shops they surveyed.

2.6.1 Oligochaetes

Oligochaetes such as earthworms are popular baits for freshwater fishes in many parts of Australia (Ross

1995). A variety of oligochaetes are used as bait, likely including several genera and species within the

Families Acanthodrilidae, Lumbricidae, Octochaetidae and Megascolecidae (Blakemore 1999),12. Data are

not available, however commercially available composting worms, such as Eisenia foetida, Lumbricus

rebellus and Perionyx excavatus are probably used more frequently than native varieties as they are readily

available in many retail outlets throughout Australia.

2.6.2 Polychaetes

Polychaetes are popular baits in estuarine and marine regions throughout most of Australia (Ross 1995). A

large number of different types of polychaetes are used as bait by both commercial and recreational fishers,

including members of the Family Eunicidae, Glyceridae, Onuphidae and Nereididae (Kailola et al. 1993).

Diopatra dentata is a common tube worm which is sometimes used as bait, while in recent years,

aquaculture of the tube worm Diopatra aciculata (F. Onuphidae) has made this species widely available both

as live worms and preserved/freeze dried worms10. Several species of beach worms (F. Onuphidae) are

widely used as bait along the east coast (Paxton 1979, Bennett 1987), including the king worm

(Australonuphis teres), slimy beachworm (A. parateres), stripey beachworm (Onuphis taeniata), giant

beachworm (Hirsutonuphis gygis) and wiry beachworm (H. mariahirstua) (see Paxton 1979, 1996). The

common bait worm (Australonereis ehlersi), is found throughout country and is widely used as bait for many 11 www.aquabait.com.au 12 http://australianmuseum.net.au/Australian-Earthworms

_________________________________________________________________________________________________________________________________________________________________________________


42

www.digsfish.com

species of estuarine fish (Grixti et al. 2007). Marphysa sanguinea, (F. Eunicidae) also known as the blood

worm, or Cribb Island blood worm, is widely used by anglers along the eastern seaboard and is a common

subject of the live worm trade (Kailola et al. 1993). Glycera ovigera (F. Glyceridae) is a popular bloodworm

bait in South Australia (Davies et al. 2008). The Nereid polychaetes Nephtys australiensis and Perinereis

nuntia are found Australia wide (Ranier and Hutchings 1977). Some of these Nereid species are known in

some areas as wrigger worms, and are widely used and translocated along the east coast as a specialist bait

for certain estuarine species (DPIF 2004). Informal interviews with many experienced anglers suggest that

leeches (Hirudinea) are not used as bait in Australia.

2.7 Other

Kewagama Research (2002, 2007) recorded use of “other aquatic animals” in their bait and berley surveys.

These may have includes barnacles (Crustacea), limpets (Molluca), cunjevoi (Ascidacea: Pyuridae),

echinoderms, seaweeds, aquatic insects and others. In 2002, “other aquatic animals” were used by 2.1% of

respondents from 5 states, namely Tasmania (11%), NSW/ACT (4.7%), QLD (0.9%), SA and WA (0.4%)

(Kewagama Research (2002). In 2006, only 0.5% of respondents from one state recorded using “other

aquatic animals” as bait, namely 2.2% of respondents from Victoria (Kewagama Research 2007). These data

suggest that the use of the following groups of aquatic animals as bait in Australia is uncommon.

2.7.1 Echinoderms

The internal organs of echinoderms such as sea urchins are occasionally used by rock fishing enthusiasts as

berley to help target certain demersal fish species that live in and near rocky reefs along the southern half of

the Australian coastline13. The species harvested for bait and berley include common urchins that occur on

rocky reefs, such as the spiny sea urchin (Centrostephanus rodgersii), purple sea urchin (Helicocidaris spp.),

pink sea urchin (Holopneustes pycnotilus), hairy sea urchin (Tripneustes gratilla), and others. Commercial

rock lobster fishers also may use sea urchins as bait, and significant quantities may be used at times. The

quantity of echinoderms used for bait and berley in the recreational fishery is likely to be limited and the

majority (indeed, probably all of the volumes used), are likely to be collected and used at the fishing site.

2.7.2 Cunjevoi (Ascidians)

Ascidians such as cunjevoi (Pyura stolonifera) are used as bait and berley by rock fishing enthusiasts to

target certain demersal fish species that live near rocky reefs along the southern half of the Australian

coastline (Bennett 1987, Ross 1995). The quantity of cunjevoi used is probably relatively small and the vast

majority of it is likely to be collected and used at the fishing site.

Summary

The main groups and species of aquatic animals used as bait and berley in Australia are summarised in Table

2 (over page). 13 http://www.nswfca.com.au/rokgrp03.html

_________________________________________________________________________________________________________________________________________________________________________________


43

www.digsfish.com

Table 2. Commodity List: The species most widely used as bait and berley in Australia.

Scientific name Common Name States where used FINFISH Saltwater finfish F. Arripidae Australian salmon, tommy

ruff NSW, Tas, Vic, SA, WA

F. Atherinidae Hardyheads, silversides, whitebait

All states

F. Carangidae Trevallies, jack mackerel All states F. Congridae Conger eels SA F. Clupeidae Herring, sardines, pilchards All states F. Emmelichthyidae Redbait Tas, SA F. Engraulidae Anchovies, sprats All states F. Gempylidae Barracouta, gemfish NSW, Vic, Tas F. Gerridae Silver biddies All states F. Hemirhamphidae Garfish All states F. Labridae Wrasse NSW, Vic, Tas, SA F. Leiognathidae Ponyfishes QLD, NSW, WA, NT F. Lethrinidae Emperors QLD, NSW, WA, NT F. Lutjanidae Snappers QLD, NSW, WA, NT F. Monacanthidae Leatherjackets NSW, Vic, Tas, SA F. Moridae Rock cods NSW, Vic, Tas, SA F. Muglidae Mullet All states F. Nemipteridae Whiptails QLD, WA, NT F. Scombridae Bonitos, mackerels, tunas All states F. Salmonidae Trout, salmon Tasmania, Vic F. Sillaginidae Whitings All states F. Sparidae Breams All states Tailor Pomatomus saltatrix QLD, NSW, Vic, WA Freshwater finfish F. Ambassidae Perchlets QLD, NSW, Vic, SA, NT, WA Carassius auratus Goldfish QLD, NSW/ACT, Vic, Tas, SA, WA F. Cichlidae Tilapias QLD, Vic, WA Cyprinus carpio European carp QLD, NSW/ACT, Vic, Tas, SA, WA F. Eleotridae Gudgeons All states F. Galaxiidae Galaxids NSW, Vic, SA, Tas, WA F. Hemiramphidae Garfish All states F. Melanotaeniidae Rainbowfish QLD, NSW, Vic, SA, NT, WA Misgurnus anguillicaudatus Oriental weatherloach QLD, NSW/ACT, Vic F. Muglidae Mullet All states Nematalosa spp. Bony bream QLD, NSW, Vic, SA, NT, WA Perca fluviatilis Redfin NSW/ACT, Vic, SA, WA F. Poeciliidae Gambusia and swordtails All states F. Retropinnidae Smelts QLD, NSW, Vic, SA, Tas F. Salmonidae Trout, salmon NSW/ACT, Vic, Tas, SA, WA F. Teraponidae Grunters QLD, NSW, SA, NT, WA Toxotes sp. Archerfish QLD, NT, WA

_________________________________________________________________________________________________________________________________________________________________________________


44

www.digsfish.com

Scientific name Common Name States where used CRUSTACEANS Prawns and shrimp Macrobrachium rosenbergii Giant freshwater prawn QLD, NT, WA Macrobrachium spp. Other shrimp QLD, NSW, WA, NT Metapenaeopsis spp. Other prawns QLD, NT, WA Metapenaeus bennettae Greasyback prawns QLD, NSW, NT, WA Metapenaeus macleayi School prawns QLD, NSW, Vic Metapenaeus spp. Other prawns QLD, NSW, WA, NT Palaemon spp. Other shrimp All states Parapenaeopsis spp. Other prawns QLD, NT, WA Penaeus esculentus, P. semisulcatus, P. monodon

Tiger prawns QLD, NT, WA

Penaeus indicus, P. merguiensis Banana prawns QLD, NT, WA Penaeus plebejus, P. latisulcatus King prawns Vic, SA Penaeus spp. Other prawns All states Crabs Carcinus maenus European green crab NSW, Vic, Tas, WA Leptograpsus spp., Paragrapsus spp.

Rock crabs QLD, NSW, Vic, Tas, SA, WA

Mictyris longicarpus, M. platycheles spp.

Soldier crabs QLD, NSW, Vic, Tas, NT, WA

Ocypode spp. Ghost crabs QLD, NSW, Vic, NT, WA Plagusia chabrus, Plagusia spp. Bait crabs QLD, NSW, Vic, Tas, SA, WA Portunus pelagicus Blue (sand) crab All states Scylla serrata Mud crab QLD, NSW, WA, NT Saltwater yabbies/nippers Biffarius arenosus Yabby/nipper QLD, NSW, Vic Callianassa (Trypea) australiensis

Yabby/nipper QLD, NSW, Vic

Upogebia spp. Yabby/nipper QLD, NSW, Vic Freshwater crayfish Cherax destructor Yabby QLD, NSW, Vic, SA, WA Cherax quadricarinatus Redclaw crayfish QLD, NT, NSW Cherax tenuimanus, C. cainii Marron WA Cherax spp. Freshwater crayfish All states Engaeus spp. Freshwater crayfish NSW, Vic, Tas Euastacus spp. Freshwater crayfish All states Saltwater crayfish/lobsters Panulirus cygnus Western rock lobster WA Panulirus spp. Tropical lobsters WA, NT, QLD Jasus edwardsii Southern rock lobster WA, SA, Vic, Tas Jasus (Sagmariasus) verreauxi Eastern rock lobster Vic, NSW

_________________________________________________________________________________________________________________________________________________________________________________


45

www.digsfish.com

Scientific name Common Name States where used MOLLUSCS Gastropods Cellana spp. Limpets QLD, NSW, Vic, SA, WA, Tas Haliotis laevigata Greenlip abalone WA, SA, Vic, Tas, NSW Haliotis rubra Blacklip abalone WA, SA, Vic, Tas, NSW Haliotis roei Roes abalone WA Haliotis spp. Abalone All states Bivalves Amusium spp., Pecten spp., Chlamys spp.

Scallops All states

Crassostrea gigas Pacific oysters SA, Tas Vic, NSW F. Hyriidae Freshwater mussels All states Katelysia spp., Anadara spp., Barbatia spp.

Cockles NSW, Vic, Tas, SA, WA

Mytilus edulis /M. galloprovincialis

Mussels NSW, Vic, Tas, SA, WA

Pinna bicolour, P. deltoides, Isognomon spp.

Razor shells SA , WA

Plebidonax deltoides Pipis SA, Vic, NSW, QLD, WA Saccostrea glomerata Sydney rock oysters QLD, NSW Saccostrea cuccullata, S. echinata Tropical rock oysters QLD, NT, WA Cephalopods Loligo spp. Mitre squid All states Notodarus gouldi Arrow squid (Goulds squid) All states Notodarus spp. Other squid All states Octopus australis Southern octopus NSW, Vic, SA, Tas Octopus cyanea Reef octopus QLD, WA, NT Octopus maorum Maori octopus NSW, Vic, SA, Tas Octopus pallidus Pale octopus NSW, Vic, SA, Tas Octopus tetricus Gloomy octopus QLD, NSW, WA Octopus spp Other octopuses All states Sepia apama Giant cuttlefish All states Sepia rozella Rosecone cuttlefish QLD, NSW Sepia spp Other cuttlefish All states Sepioteuthis australis Southern calamari NSW, Vic, Tas, SA, WA Sepioteuthis lessoniana Northern calamari QLD, NT, WA ANNELIDS Oligochaetes F. Acanthodrilidae Earthworms QLD, NSW, Vic, Tas, SA, WA Eisenia foetida Tiger worm All states Lumbricus rebellus Red worm All states F. Lumbricidae Earthworms All states F. Megascolecidae Earthworms All states F. Octochaetidae Earthworms QLD, NSW, SA, WA, NT Perionyx excavatus Blue worm All states

_________________________________________________________________________________________________________________________________________________________________________________


46

www.digsfish.com

Scientific name Common Name States where used Polychaetes Australonereis ehlersi Bait worm All states Australonuphis parateres Slimy beachworm QLD, NSW, Vic, SA Australonuphis teres King beachworm QLD, NSW, Vic Diopatra dentata, D. aciculata Tube worm All states F. Eunicidae All states Glycera ovigera Blood worm NSW, Vic, SA, Tas F. Glyceridae NSW, Vic, SA, Tas Marphysa sanguinea Cribb Island blood worm QLD, NSW Nephtys australiensis, Perinereis nuntia

Wriggler worms All states

F. Nephtyidae All states F. Nereididae All states Hirsutonuphis gygis Giant beachworm QLD Hirsutonuphis mariahirstua Wiry beachworm QLD, NSW, Vic Onuphis taeniata Stripey beachworm QLD, NSW, Vic ECHINODERMS Centrostephanus rodgersii Spiny sea urchin NSW, Vic, SA, WA, Tas Helicocidaris spp. Purple sea urchin QLD, NSW, Vic, SA, WA, Tas Holopneustes pycnotilus Pink sea urchin NSW, Vic, SA, WA, Tas Tripneustes gratilla Hairy sea urchin WA, QLD, NSW ASCIDEANS Pyura stolonifera Cunjevoi QLD, NSW, Vic

_________________________________________________________________________________________________________________________________________________________________________________


47

www.digsfish.com

3.0 The methodology used for this Risk Analysis

3.1 Hazard Identification

After defining the commodities of concern (see Table 2), the next step in the RA is to identify the potential

hazards present in those commodities. This process, hazard identification, begins with the compilation of a

list of disease agents known to be associated with the commodities. The criteria for consideration during the

hazard identification process were as follows:

For each organism in the initial hazard list, the following questions were considered:

1. Whether the commodity could potentially introduce the disease agent,

2. If the disease agent is "under official control", by its listing in State or National lists of reportable

diseases (Tables 3a, 3b), or

3. If the disease agent is restricted in its distribution and/or could conceivably cause a detrimental

impact to industry or the environment if infected commodities were translocated into new areas.

For any disease agent, if the answers to any of these questions was ‘yes’, it was classified as a potential

hazard (Figure 2). Each disease agent identified as a potential hazard was then critically evaluated. Any

disease agents considered likely to cause detrimental impacts in Australia based on one or more of the

following criteria were classed as diseases of concern (hazards) that required detailed risk assessment. The

criteria used included whether:

• it would be expected to cause a distinct pathological effect in an infected population; and/or

• it would be expected to cause economic harm (e.g. increased mortality, reduced growth rates,

decreased product quality, loss of market access, increased costs); and/or

• it would be expected to cause damage to the environment and/or endemic species (defined as either

native species that occur naturally in Australia waters, or species that were introduced into Australia

and are now considered to be acclimatised).

The process used for decision making in relation to the hazard identification process is summarised below in

Figure 2. Disease agents that are not considered likely to cause a distinct pathological effect in affected

populations, and/or economic harm, and/or damage to the environment were considered to represent a

negligible risk, and were excluded from further assessment.

_________________________________________________________________________________________________________________________________________________________________________________


48

www.digsfish.com

Figure 2. Flow chart showing the decision making process used to identify potential hazards in the hazard identification step.

No

Identified as a disease of concern requiring detailed

risk assessment

Disease agent expected to cause a distinct pathological effect in affected populations, and/or economic harm, and/or damage to the environment ?

Yes

Is the disease agent listed in Australias national or state reportable disease lists as under “official

control”

Identified as a potential hazard

Is the disease agent restricted in its

distribution within Australia ?

Yes

No

No

Yes

Could the commodity carry the disease agent ? Exclude from further

examination in RA No

Yes

Develop list of disease agents associated with each

commodity

Develop list of commodities

_________________________________________________________________________________________________________________________________________________________________________________


49

www.digsfish.com

Table 3a. National list of reportable diseases of aquatic animals (ie. diseases under official control).

Australia’s National List of Reportable Diseases of Aquatic Animals (Endorsed by Aquatic Animal Health Committee –May 2010)

Listed in the OIE Aquatic Animal Health Code (2010)

Listed regionally (OIE/NACA) (2010)

Exotic to Australia

FINFISH 1. Epizootic haematopoietic necrosis – EHN virus � � 2. Epizootic haematopoietic necrosis – European catfish virus /

European sheatfish virus � �

3. Infectious haematopoietic necrosis � � � 4. Spring viraemia of carp � � � 5. Viral haemorrhagic septicaemia � � � 6. Channel catfish virus disease � � 7. Viral encephalopathy and retinopathy � 8. Infectious pancreatic necrosis � 9. Infectious salmon anaemia � � � 10. Epizootic ulcerative syndrome (Aphanomyces invadans) � � 11. Bacterial kidney disease (Renibacterium salmoninarum) � 12. Enteric septicaemia of catfish (Edwardsiella ictaluri) � 13. Piscirickettsiosis (Piscirickettsia salmonis) � 14. Gyrodactylosis (Gyrodactylus salaris) � � � 15. Red sea bream iridoviral disease � � � 16. Furunculosis (Aeromonas salmonicida subsp. salmonicida) � 17. Aeromonas salmonicida - atypical strains 18. Whirling disease (Myxobolus cerebralis) � 19. Enteric redmouth disease (Yersinia ruckeri – Hagerman strain) � 20. Koi herpesvirus disease � � � 21. Grouper iridoviral disease � �

MOLLUSCS 1. Infection with Bonamia ostreae � � � 2. Infection with Bonamia species 3. Infection with Bonamia exitiosa � � 4. Infection with Mikrocytos mackini � 5. Infection with Marteilia refringens � � � 6. Infection with Marteilia sydneyi 7. Infection with Marteilioides chungmuensis � � 8. Infection with Perkinsus marinus � � � 9. Infection with Perkinsus olseni � � 10. Infection with Xenohaliotis californiensis � � � 11. Akoya oyster disease � � 12. Iridoviroses � 13. Abalone viral mortality � � � 14. Abalone viral ganglioneuritis � �

CRUSTACEANS 1. Taura syndrome � � � 2. White spot disease � � � 3. Yellowhead disease – Yellowhead virus � � � 4. Gill-associated virus 5. Infectious hypodermal and haematopoietic necrosis � � 6. Crayfish plague (Aphanomyces astaci) � � � 7. White tail disease � � 8. Infectious myonecrosis � � � 9. Monodon slow growth syndrome � � 10. Milky haemolymph diseases of spiny lobster (Panulirus sp.) � �

AMPHIBIANS 1. Infection with Batrachochytrium dendrobatidis � � 2. Infection with ranavirus � �

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

50

w

ww

.dig

sfis

h.co

m

Tab

le 3

b.

Sta

te li

sts

of r

epor

tabl

e di

seas

es o

f aqu

atic

an

imal

s (ie

. di

seas

es u

nder

offi

cial

co

ntro

l in

each

State

).

Dis

ease

age

nts

QLD

N

SW

V

IC

AC

T

TA

S

SA

W

A

NT

FIN

FIS

H

1.

Aer

om

on

as

salm

onic

ida

- aty

pica

l str

ains

�

�

�

N

o �

�

�

�

2.

A

qua

tic b

irna

viru

s no

no

no

N

o �

no

no

no

3.

B

act

eria

l kid

ney

dise

ase

(Ren

iba

cter

ium

sal

mo

nin

aru

m) �

�

no

�

no

�

�

�

4.

C

hann

el c

atfi

sh v

irus

dise

ase

�

�

�

�

�

�

�

�

5.

Ent

eric

re

dmou

th d

ise

ase

(Y

ersi

nia

ruck

eri –

Ha

germ

an

stra

in)

�

�

�

�

�

�

�

�

6.

Ent

eric

se

ptic

ae

mia

of c

atfi

sh (

Ed

wa

rdsi

ella

icta

luri)

�

�

�

�

�

�

�

�

7.

E

pizo

otic

ha

em

atop

oie

tic n

ecr

osis

– E

HN

vir

us

�

�

�

No

�

�

�

�

8.

Epi

zoot

ic h

ae

mat

opoi

etic

ne

cros

is –

Eur

ope

an c

atfi

sh v

irus

/

Eur

ope

an

she

atfis

h vi

rus

�

�

�

�

no

�

no

no

9.

Epi

zoot

ic u

lce

rativ

e s

yndr

ome

(Aph

an

om

yces

inva

dan

s)

no

�

�

No

�

�

�

�

10.

Fur

uncu

losi

s (Aer

om

on

as s

alm

on

icid

a subs

p. sa

lmo

nici

da)

�

�

�

�

�

no

�

�

11.

Gro

upe

r iri

dovi

ral d

isea

se

�

�

�

�

�

�

�

no

12.

Gyr

oda

ctyl

osis

(Gyr

oda

ctyl

us

sala

ris)

�

�

�

�

�

�

�

�

13.

Infe

ctio

us h

aem

ato

poie

tic n

ecr

osis

�

�

�

�

�

�

�

�

14

. In

fect

ious

pan

cre

atic

ne

cros

is �

�

�

�

�

�

�

�

15

. In

fect

ious

sal

mon

ana

em

ia �

�

�

�

�

�

�

�

16

. K

oi h

erpe

svir

us d

ise

ase

�

�

�

�

�

�

�

no

17

. K

oi M

ass

mor

talit

y no

no

�

N

o no

no

no

no

18

. O

nco

rhyn

chu

s m

aso

u vir

us d

ise

ase

(OM

VD

) no

no

no

�

�

�

no

�

19

. P

isci

rick

etts

iosi

s (Pis

ciri

cket

tsia

sa

lmo

nis)

�

�

�

�

�

�

�

�

20.

Re

d se

a br

ea

m ir

idov

ira

l dis

ea

se

�

�

�

�

�

�

�

no

21.

Ric

ketts

ia-li

ke o

rga

nism

(R

LO)

of s

alm

onid

s no

no

no

N

o �

no

no

no

22

. S

ea

lice

(Lep

eop

hth

eriu

s sa

lmon

is) no

no

no

N

o �

no

no

no

23

. S

prin

g vi

rae

mia

of c

arp

�

�

�

�

�

�

�

�

24.

Str

ept

ococ

cosi

s (Str

epto

cocc

us

inia

e) no

no

no

N

o no

no

no

�

25

. S

tre

ptoc

occo

sis

of s

alm

onid

s (

Lac

toco

ccu

s g

arvi

eae)

no

no

no

No

�

no

no

no

26.

Vir

al e

nce

pha

lopa

thy

and

ret

inop

ath

y

no

�

�

�

�

�

�

�

27.

Vir

al h

aem

orrh

agi

c se

ptic

ae

mia

�

�

�

�

�

�

�

�

28.

Whi

rling

dis

eas

e (Myx

ob

olus

cer

ebra

lis)

�

�

�

�

�

�

�

�

29.

Whi

te s

turg

eon

irid

ovir

al d

isea

se

no

no

no

�

no

�

no

no

MO

LLU

SC

S

1.

Aba

lone

vir

al g

ang

lione

uriti

s �

�

�

N

o �

�

�

�

2.

A

balo

ne v

iral

mor

talit

y �

�

�

N

o no

�

�

�

3.

A

koya

oys

ter

dise

ase

�

�

�

�

�

�

�

�

4.

B

oca

rdia

kn

oxi

(mud

wor

m)

no

no

no

No

no

�

no

no

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

51

w

ww

.dig

sfis

h.co

m

Dis

ease

age

nts

QLD

N

SW

V

IC

AC

T

TA

S

SA

W

A

NT

5.

Ha

plos

porid

osis

, Ha

plo

spo

ridiu

m n

elso

ni, H

. co

stal

e, H

. sp

. no

no

�

�

�

�

�

�

6.

In

fect

ion

with

Bo

na

mia

exi

tiosa

�

�

�

�

�

�

�

�

7.

In

fect

ion

with

Bo

na

mia

ost

rea

e �

�

�

�

�

�

�

�

8.

In

fect

ion

with

Bo

na

mia

ro

ughl

eyi

�

�

�

No

�

�

�

�

9.

Infe

ctio

n w

ith B

on

am

ia sp

eci

es

�

�

�

No

�

�

�

�

10.

Infe

ctio

n w

ith M

arte

ilia

ref

ring

ens

�

�

�

�

�

�

�

�

11.

Infe

ctio

n w

ith M

arte

ilia

syd

ney

i no

�

�

no

�

�

�

�

12

. In

fect

ion

with

Mar

teili

oid

es c

hun

gm

uen

sis

�

�

�

no

�

�

no

no

13.

Infe

ctio

n w

ith M

ikro

cyto

s m

acki

ni �

�

�

�

�

�

�

�

14

. In

fect

ion

with

Per

kins

us

mar

inu

s �

�

�

�

�

�

�

�

15

. In

fect

ion

with

Per

kins

us

olse

ni

�

�

�

no

�

�

�

�

16.

Infe

ctio

n w

ith X

eno

halio

tis c

alifo

rnie

nsi

s �

�

�

�

�

�

�

no

17

. Ir

idov

iros

es

�

�

�

�

�

�

�

�

18.

Noc

ard

osis

of s

hellf

ish

no

no

no

no

�

no

no

no

19.

Oys

ter

Oe

dem

a d

ise

ase

no

no

no

no

no

no

�

no

20

. O

yste

r V

ela

r V

irus

Dis

ea

se (

OV

VD

) no

no

no

no

no

�

no

no

21

. P

erki

nsu

s spp

. (e

xotic

) no

no

no

no

no

�

�

�

CR

US

TA

CE

AN

S

1.

Ba

culo

vira

l mid

gut g

land

ne

cros

is

no

no

no

�

no

�

no

�

2.

Ba

culo

viru

s pe

naei

(B

P)

/ te

trah

edr

al b

acu

lovi

rosi

s no

no

�

�

no

no

�

�

3.

C

rayf

ish

pla

gue (A

pha

no

myc

es a

sta

ci) �

�

�

�

�

�

�

�

4.

G

ill-a

ssoc

iate

d vi

rus

�

�

�

no

�

�

�

�

* 5.

In

fect

ious

hyp

ode

rma

l and

hae

ma

topo

ietic

ne

cros

is

�

�

�

�

�

�

�

�

6.

Infe

ctio

us m

yone

cros

is

�

�

�

no

�

�

�

no

7.

Mic

rosp

orid

osis

no

no

no

no

no

no

�

no

8.

M

ilky

hae

mol

ymph

dis

ease

s of

sp

iny

lobs

ter

(P

an

uliru

s sp.

) �

�

no

no

�

�

no

no

9.

M

onod

on b

acu

lovi

rus

/ sph

eric

al b

acu

lovi

rosi

s no

no

no

no

no

no

�

�

10

. M

ono

do

n slo

w g

row

th s

yndr

ome

�

no

�

no

no

�

no

no

11

. N

ecr

otiz

ing

hepa

topa

ncre

atiti

s no

no

no

�

no

�

�

�

12

. S

acc

ulin

a spp

. no

no

no

no

no

�

**

no

no

13.

Spa

wne

r is

ola

ted

mor

talit

y vi

rus

no

no

no

no

no

�

no

revi

ew

14

. T

aur

a s

yndr

ome

�

�

�

�

�

�

�

�

15

. W

hite

spo

t di

seas

e

�

�

�

�

�

�

�

�

16.

Whi

te ta

il di

sea

se

�

�

�

no

no

no

no

no

17.

Ye

llow

hea

d di

seas

e –

Yel

low

head

viru

s �

�

�

�

�

�

�

�

AM

PH

IBIA

NS

1.

In

fect

ion

with

Ba

tra

cho

chyt

rium

den

dro

bat

idis

�

�

�

�

�

�

�

2.

In

fect

ion

with

ra

navi

rus

�

�

�

�

�

�

�

* u

nder

rev

iew

, *

* no

t in

clu

ded

in n

ew li

st u

nder

the

live

sto

ck a

ct

_________________________________________________________________________________________________________________________________________________________________________________


52

www.digsfish.com

3.2 Risk assessment

Once the hazards were identified, a risk assessment was carried out on each disease of concern. This RA

was based on a qualitative assessment of the risks involved with translocation of these commodities. The

qualitative RA method addresses risk in a standardised manner (Jones and Stephens 2006, OIE 2010a)

utilising a series of risk assessment processes, namely release assessment; exposure assessment; consequence

assessment; and risk estimation. More detail on each step in the process is included below.

3.2.1 Release assessment

The likelihood that a hazard would be translocated with the commodity is determined through the release

assessment stage of the process. The ‘unrestricted commodities’ considered here are live, fresh dead (green),

frozen (uncooked/green) or preserved/freeze dried fish, crustaceans, molluscs, annelids and other aquatic

animals used as bait or burley. The most likely release pathways for spread of disease from commodities

into the Australian aquatic environment were shown in Figure 1.

Translocation of infected bait and berley potentially presents a direct transmission pathway for the

introduction and establishment of new disease agents into previously disease free waterways (i.e. an

uninfected jurisdiction). In determining the likelihood of viable and infective disease agents being released

into an uninfected jurisdiction via bait and berley products in Australia, one key factor was considered:

1. Infected Bait: The bait, live or dead, or that portion of the animal used for bait and berley must be

infected with viable disease agent(s). Live or fresh dead fish and invertebrates with patent infections

may have extremely high titres of disease agents in their tissues. However, prevalence and intensity

of infection of disease agents in the bait and berley may vary and also would be subject to processing

and environmental conditions which may or may not promote the survival of potential pathogens.

These may include freezing (Archer 2004), thawing, preservation and long-term storage. While little

data are available on the survival of specific pathogens in visceral or muscle tissues, some data exists

on survival of pathogens under conditions of freezing, heating and desiccation in various media.

Survival of viral and microbial pathogens in frozen tissues is well documented.

The likelihood estimations made in this RA for the release assessment (Table 4) are qualitative assessments

based on information available in the scientific (and other) literature, unpublished data, as well as the

professional judgment of the analyst.

The risk assessment for a particular hazard was concluded if the release assessment determined that the

likelihood of release of that hazard was negligible (OIE 2010a).

_________________________________________________________________________________________________________________________________________________________________________________


53

www.digsfish.com

Table 4. Nomenclature for the qualitative likelihood estimations used in this RA.

Likelihood Definition

High The event would be very likely to occur

Moderate The event would occur with an even probability

Low The event would be unlikely to occur

Very Low The event would be very unlikely to occur

Extremely low The event would be extremely unlikely to occur

Negligible The event would almost certainly not occur

3.2.2 Exposure assessment

The exposure assessment examines the likelihood of wild aquatic animals in an uninfected jurisdiction being

exposed to the hazards via infected and released commodities and determines the likelihood of the

establishment of the hazard. The release pathway will be use of the commodity as bait and/or berley and the

likelihood of exposure will depend on several factors relating to the capacity of the disease agent to survive

in the environment in an infective form, the availability of susceptible hosts, the ease of infection of

susceptible hosts, and the likelihood of subsequent transmission of infection to others within a population. In

determining the likelihood of exposure of susceptible hosts to disease agents being released from an infected

jurisdiction to an uninfected jurisdiction via translocation of bait and berley products, the following key

factors were considered relevant:

1. Route of Infection (Oral/Contact): The bait containing viable disease agents must be ingested by a

susceptible host or otherwise come into contact with susceptible fish or invertebrate species.

Infection may occur via the digestive tract, through direct contact with contaminated water via the

skin and gills.

2. Infective Dose: There must be sufficient quantities of viable disease agents to induce an infection

following ingestion or contact with the contaminated bait or berley via the skin and gills or

integument. There is relatively little data on minimum infectious doses required for disease agents

of aquatic animals. In general, titres in muscle tissues tend to be lower whereas titres in visceral

organs and tissues tend to be higher.

Once a hazard is released into the environment, the likelihood of whether the disease agent would survive,

infect susceptible hosts, and become established within a population was expressed qualitatively using the

likelihood estimations in Table 4, based on information available in the scientific (and other) literature,

unpublished data, as well as the professional judgment of the analyst. The likelihoods for the release and

_________________________________________________________________________________________________________________________________________________________________________________


54

www.digsfish.com

exposure assessments were combined using the matrix of ‘rules’ for combining descriptive likelihoods, as

shown in Table 5.

Table 5. Matrix of rules for combining descriptive likelihoods for the release and exposure

assessments.

Liklihood of exposure

High Moderate Low Very Low Extremely low Negligible

High High Moderate Low Very Low Extremely low Negligible

Moderate Low Low Very Low Extremely low Negligible

Low Very Low Very Low Extremely low Negligible

Very Low Extremely low Extremely low Negligible

Extremely low Negligible Negligible

Negligible Negligible

The risk assessment for a particular hazard was concluded if the exposure assessment determined that the

probability of establishment was negligible (OIE 2010a).

3.2.3 Consequence assessment

The consequence assessment estimates the likely magnitude of the consequences of establishment and/or

spread of a hazard into a new area or catchment, and the possible effects of the disease agent on aquatic

animals, the environment, industry and the economy. The qualitative terms used to describe the

consequences of establishment of an unwanted disease agent in this RA are defined in Table 6. These

descriptions are based on information available in other RAs (Jones and Stephens 2006, Biosecurity

Australia 2009), the scientific literature, unpublished data, as well as the professional judgment of the

analyst.

For each hazard of concern, the consequence assessment determined the likelihood of occurrence and the

associated impact for each of two main outbreak scenarios. Either:

1. The disease agent becomes established and spreads throughout populations of susceptible species in

a new region of Australia. This scenario assumes that if an agent were to establish in a local

population it would eventually spread to its natural geographical limits, or;

Like

lihoo

d of

rel

ease

_________________________________________________________________________________________________________________________________________________________________________________


55

www.digsfish.com

2. An index case occurs and infection may even spread to co-habiting animals, but the agent does not

persist in the environment.

Only the first scenario was considered to represent establishment of the disease agent, because the second

scenario would go undetected.

Table 6. Definition of terms used to describe consequences of establishment of unwanted disease agents.

Consequence Definition

Extreme Establishment of disease would cause substantial biological and economic harm at a

regional or even national level, and/or cause serious and irreversible harm to the

environment.

High Establishment of disease would have serious biological consequences (high

mortality or morbidity) and would not be amenable to control or eradication. Such

diseases would significantly harm economic performance at a regional level and/or

cause serious environmental harm which is most likely irreversible.

Moderate Establishment of disease would cause significant biological consequences

(significant mortality or morbidity) and may not be amenable to control or

eradication. Such diseases could harm economic performance at a regional level on

an ongoing basis and/or may cause significant environmental effects, which may or

may not be irreversible.

Low Establishment of disease would have moderate biological consequences and would

normally be amenable to control or eradication. Such diseases may harm economic

performance at a local level for some period and/or may cause some environmental

effects, which would not be serious or irreversible.

Very Low Establishment of disease would have mild biological consequences and would be

amenable to control or eradication. Such diseases may harm economic performance

at a local level for a short period and/or may cause some minor environmental

effects, which would not be serious or irreversible.

Negligible Establishment of disease would have no significant biological consequences and

would require no management. The disease would not affect economic performance

at any level and would not cause any detectable environmental effects.

The risk assessment for a particular hazard was concluded if the consequence assessment determined that the

consequences of introduction were negligible (OIE 2010a).

_________________________________________________________________________________________________________________________________________________________________________________


56

www.digsfish.com

3.2.4 Risk estimation

Risk estimation is the final step involved with each assessment and would be used to determine whether the

extent of the unrestricted risk presented by each disease agent to the aquatic animals, environment, industries

and community of Australia was sufficient to require risk management. ‘Unrestricted risk’ means the

estimated risk if the various bait and berley commodities were to be translocated with no risk management

measures in place. Risk was assessed using the risk estimation matrix in Table 7 which uses a combination of

the qualitative answers given for the combined likelihoods of release and exposure and the significance of

the consequences of establishment of a disease agent to provide an estimate of the risk involved, ranging

from ‘negligible’ through to ‘extreme’.

The appropriate level of protection (ALOP) adopted in this RA is expressed in qualitative terms. Australia’s

ALOP, which reflects community expectations through government policy, is expressed as providing a high

level of sanitary or phytosanitary protection whereby risk is reduced to a very low level, but not to zero. This

definition of ALOP, and its illustration by way of a risk estimation matrix is shown below in Table 7.

Table 7. Risk estimation matrix showing the ALOP utilized for this RA (white squares = very low risk). Any diseases which fall to the right of the ALOP during the RA will require additional risk

management (red font).

Negligible risk

Very low risk

Low risk Moderate risk

High risk Extreme risk

Negligible risk

Very low risk


High risk Extreme risk

Negligible risk

Negligible risk

Very low risk


High risk

Negligible risk

Negligible risk

Negligible risk

Very low risk


Negligible risk

Negligible risk

Negligible risk

Negligible risk

Very low risk

Low risk

Negligible risk

Negligible risk

Negligible risk

Negligible risk

Negligible risk

Very low risk

If either the likelihood of establishment and spread, or the significance of the consequences of establishment

and spread were considered to be negligible, it was considered the unrestricted risk posed by the disease

agent was negligible (rising to very low for extreme consequences of establishment), and there would be no

Consequences of establishment and spread

Negligible Very Low Low Moderate High Extreme

High Moderate

Low Very low

Ext. Low Negligible Li

kelih

ood

of e

stab

lishm

ent a

nd s

prea

d

_________________________________________________________________________________________________________________________________________________________________________________


57

www.digsfish.com

need to implement any additional risk management steps (Table 7). If the consequences of establishment

and spread were considered to be very low, even a high probability of establishment and spread was tolerable

without the need for risk management. If the likelihood of establishment and spread were considered to be

very low, even high consequences of establishment and spread were tolerated without the need for risk

management, but extreme consequences of establishment and spread were considered to exceed the ALOP,

and risk management would be required (Table 7). Alternatively, if the likelihood of establishment and

spread was high, even if the consequences of establishment and spread were considered to be low, this

scenario would exceeded the ALOP and require risk management (Table 7).

3.2.5 Risk mitigation

If the unrestricted risk estimation for any disease agent is determined to be unacceptable (that is above very

low), the threats posed by the commodity will be ranked (high, medium, low) based on the likelihood the

commodity will pose a disease risk when translocated into new areas and catchments. The ranking process

will take into account not only the types of disease agents harboured in the commodity, but also the volume

of the commodity used as well as the specific pathways of use. The ranking results for each commodity will

be organised based on overall risk (e.g. extreme, high, moderate, low) and stratified by pathway (live bait,

fresh dead bait, frozen bait or freeze-dried/preserved commodities). For any diseases with risk estimation

rankings that exceed the ALOP, risk mitigation measures may be necessary to reduce the risk estimate back

to within the ALOP. The risk mitigation processes examined as part of this RA process will relate only to

option evaluation.

Option evaluation

The RA will identify the options available for managing any risks that may exceed the ALOP. The options

could form the basis of a consultation process that engages stakeholders to evaluate the risks involved with

unrestricted movements of bait and berley to assess options that could be used to reduce any risks to an

acceptable level. Input from stakeholders should also be included as they may provide data on the nature of

alternative risk management measures which may be used to achieve the required risk reduction. Where

there are equivalent risk management measures that could be used to achieve the required risk reduction, the

option(s) least restrictive to trade should be employed. The final risk management methods chosen (if any)

would understandably vary on a case-by-case basis, depending on a wide variety of commodity, industry and

region-related factors.

_________________________________________________________________________________________________________________________________________________________________________________


58

www.digsfish.com

4.0 The Risk Assessment

4.1 Hazard Identification

This section will compile a list of disease agents (potential hazards) associated with the bait and berley

commodities being used in Australia (see Table 2 for the list of species considered). I have included in this

section the important disease agents reported from these commodities throughout their national distribution.

Appendix 1 contains a more comprehensive list of the parasites and disease agents recorded from the bait

and berley commodities being used in Australia, while Table 8 contains the list of the most important disease

agents which will be considered during the hazard identification process. Table 8 outlines the known host(s)

for each potential hazard, whether the hazard is infectious, whether it is under official control in Australia

(i.e. whether it is listed in State or National lists of reportable diseases of aquatic animals), whether it is an

OIE or NACA listed disease, and an indication of whether it is considered likely to cause significant disease.

In addition to the list of the most important disease agents (Table 8), it is normal for healthy aquatic animals

to be naturally infected by a variety of protozoan and metazoan parasites and symbionts. Many of these

parasites and symbionts have restricted distributions that are related to the distribution of their hosts, and/or

the biogeography of the regions in which they occur (e.g. Roubal et al. 1983, Byrnes 1986, Hayward 1997),

however most have high host specificity and/or are not known to be associated with disease. Because of this,

Table 9 lists the many parasites and symbionts that are not expected to cause significant disease or

pathological effects, and hence do not meet the criteria required for inclusion as hazards in the RA. Due to

space limitations, many specific parasites and symbionts are not mentioned in Tables 8 and 9, and instead

only higher taxonomic levels (e.g. Monogenea, Digenea, Cestoda, Nematoda, Copepoda, Ciliata,

Microsporea, Myxozoa and others) are included. The reader is referred to specialist papers on parasites (e.g.

Roubal et al. 1983, Byrnes 1986, Hayward 1997, Whittington et al. 2001) and host/parasite checklists (e.g.

Beumer et al. 1982, Lester and Sewell 1989, Fletcher and Whittington 1998, O’Donoghue and Adlard

(2000)) for more specific information relating to parasites of aquatic animals not included in Tables 8 and 9.

A wide range of the normal microbial (viral/bacterial/fungal) flora of aquatic animals are also considered to

be ubiquitous in the marine environment (Table 9). Many of these agents are not expected to cause disease

or even significant pathological effects, while a smaller number (e.g. Saprolegnia spp., Vibrio spp.,

Flavobacteria) are common opportunistic pathogens which are associated with disease only when the host is

compromised or stressed (Roberts 2001, Austin and Austin 2007). Because of this, they are not included as

potential hazards in Table 8, especially as representatives of these agents are known to already occur

throughout the country. However, pathogenic strains with more restricted distributions may still exist for

some of these microbial opportunists, and these are considered in the RA where their existence is known.

Table 8 should, therefore, not be considered as a complete list of organisms associated with aquatic animals

used as bait and berley. Instead, Table 8 represents a list of the important disease agents presently known

from the bait and berley commodities which are considered relevant to the hazard identification step of this

RA as they are likely to be associated with significant disease or pathological effects. Of course, there are

large knowledge gaps in relation to disease agents that infect aquatic animals in Australia (Appendix 1).

There remains a significant risk of transfer of as yet unknown disease agents, even in the absence of their

identification (Gaughan 2002). Because of this an example of an unknown disease agent is also included.

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

59

w

ww

.dig

sfis

h.co

m

Tab

le 8

. L

ist o

f the

mos

t im

port

ant d

isea

se a

gen

ts o

f those

aqu

atic

ani

mal

s in

Aus

tral

ia t

hat

are

used

as

bait

and/o

r be

rley.

Dis

ease

ag

ent

Hos

t(s)

that

are

kno

wn

to b

e us

ed a

s ba

it o

r bu

rley

Dis

ease

ag

ent

is

infe

ctio

us

Und

er

offic

ial

cont

rol

List

ed b

y O

IE

or

NA

CA

Like

ly t

o ca

use

sig

nific

ant

dise

ase

VIR

US

ES

Fin

fish

Aqu

atic

Birn

avi

rus

On

corh

ynch

us

myk

iss

Yes

Y

es

No

Yes

E

pizo

otic

Ha

ema

top

oiet

ic

Nec

rosi

s V

irus

(EH

NV

) O

nco

rhyn

chu

s m

ykis

s, Per

ca f

luvi

atil

is, F

. Poe

cilii

dae

Yes

Y

es

Yes

Y

es

Ha

ema

top

oiet

ic n

ecro

sis

herp

esvi

rus

of g

oldf

ish

Ca

rass

ius

aur

atu

s Y

es

No

No

No

Pilc

hard

her

pes

viru

s (P

HV

) F

. Clu

peid

ae

Yes

N

o N

o Y

es

Aqu

are

oviru

s P

erca

flu

via

tilis,

Sal

mo

sal

ar

Yes

N

o N

o N

o V

iral E

ncep

halo

path

y a

nd

Ret

inop

ath

y (N

oda

viru

s)

F. C

ara

ngid

ae,

F. E

leot

rida

e, F

. Ter

apo

nida

e Y

es

Yes

Y

es

Yes

An

unkn

own

viru

s of

fin

fish

Uns

pec

ified

fin

fish

used

as

bait

or b

erle

y Y

es

No

No

Yes

C

rust

acea

ns

B

aci

llifo

rm v

iruse

s C

her

ax

ten

uim

an

us, C

. ca

inii,

S. se

rra

ta (

SB

V)

Yes

N

o N

o N

o B

enet

tae

bacu

lovi

rus/

P

leje

bu

s ba

culo

viru

s (M

BV

str

ain

s)

Met

ap

ena

eus s

pp., P

ena

eus

ple

bej

us

Yes

Y

es

No

Yes

BM

NV

-like

P

ena

eus

mo

no

do

n, Pen

aeu

s la

tisul

catu

s Y

es

Yes

N

o N

o C

hera

x de

stru

ctor

bac

illi

form

v

iru

s (C

dB

V)

Ch

era

x d

estr

uct

or

Yes

N

o N

o N

o

Che

rax

dest

ruct

or s

yst

emic

p

arv

o-l

ike

vir

us

(Cd

SP

V)

Ch

era

x d

estr

uct

or

Yes

N

o N

o Y

es ?

Che

rax

quad

rica

rin

atu

s b

acil

lifo

rm v

iru

s (C

qB

V)

Che

rax

quad

rica

rin

atu

s Y

es

No

No

No

Gill

Ass

ocia

ted

Viru

s (G

AV

/LO

V)

Pen

aeu

s m

on

od

on, P

ena

eus

escu

len

tus

, Pen

aeu

s m

erg

uie

nsi

s,

Pen

aeu

s spp

. Y

es

Yes

Y

es

Yes

Gia

rdia

viru

s-lik

e vi

rus

(CG

V)

C

hera

x qu

adri

cari

nat

us

Yes

N

o N

o Y

es ?

Ha

emoc

ytic

rod

sha

ped

viru

s

Pen

aeu

s m

on

od

on, P

ena

eus

escu

len

tus

, Pen

aeu

s m

ergu

ien

sis

Y

es

No

No

No

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

60

w

ww

.dig

sfis

h.co

m

Dis

ease

ag

ent

Hos

t(s)

that

are

kno

wn

to b

e us

ed a

s ba

it o

r bu

rley

Dis

ease

ag

ent

is

infe

ctio

us

Und

er

offic

ial

cont

rol

List

ed b

y O

IE

or

NA

CA

Like

ly t

o ca

use

sig

nific

ant

dise

ase

Hep

ato

panc

rea

tic P

arvo

viro

sis

(HP

V)

Ma

cro

bra

chiu

m r

ose

nb

erg

ii, Ma

cro

bra

chiu

m sp

p., P

ena

eus

mo

no

do

n, P

ena

eus

escu

len

tus

, Pen

aeu

s m

erg

uie

nsi

s, P

ena

eus s

pp.,

Po

rtu

nu

s p

ela

gic

us, S

cylla

ser

rata,

Che

rax

quad

rica

rin

atu

s

Yes

N

o N

o Y

es

Infe

ctio

us H

ypod

erm

al a

nd

Ha

ema

top

oiet

ic N

ecro

sis

Viru

s (I

HH

NV

)

Met

ap

ena

eus s

pp., P

ena

eus

mo

no

do

n, Pen

aeu

s es

cule

ntu

s

Yes

Y

es

Yes

Y

es

Lym

pho

id P

arvo

-like

Viru

s (L

PV

) P

ena

eus

mo

no

do

n, Pen

aeu

s es

cule

ntu

s, P

ena

eus

mer

gu

ien

sis

Y

es

No

No

No

Mon

odon

Ba

culo

viru

s (M

BV

) P

ena

eus

mo

no

do

n, P.

escu

len

tus, P

. se

mis

ulc

atu

s, P. m

erg

uie

nsi

s, M

elic

ertu

s la

tisu

lca

tus, P

. p

leje

bu

s, Pen

aeu

s sp.

, Met

apen

aeu

s spp

. Y

es

Yes

N

o Y

es

Mou

rilya

n V

irus

(MoV

)

Pen

aeu

s m

on

od

on, P

ena

eus s

pp.

Yes

N

o N

o Y

es

Par

vovi

rus

Che

rax

quad

rica

rin

atu

s Y

es

No

No

Yes

R

eovi

rus

Che

rax

quad

rica

rin

atu

s Y

es

No

No

No

Scy

lla b

acu

lovi

rus

Scy

lla s

erra

ta Y

es

No

No

Yes

?

Spa

wne

r-Is

ola

ted

Mor

talit

y V

irus

(SM

V)

Pen

aeu

s m

on

od

on, P

ena

eus

mer

gu

ien

sis

, Che

rax

quad

rica

rin

atu

s Y

es

Yes

N

o Y

es

Ma

cro

bra

chiu

m r

ose

nbe

rgii

nod

avi

rus (

MrN

V)

/ W

hite

Ta

il D

isea

se

Ma

cro

bra

chiu

m r

ose

nb

erg

ii Y

es

Yes

Y

es

Yes

Mol

lusc

s

Aba

lone

Vira

l Ga

nglio

neur

itis

Ha

liotis

laev

iga

ta, H

alio

tis r

ub

ra

Yes

Y

es

Yes

Y

es

Dig

estiv

e ep

ithel

ial v

irosi

s A

mu

siu

m sp

p., P

ecte

n spp

. Y

es

No

No

No

Vira

l ga

met

ocyt

ic h

yper

trop

hy C

rass

ost

rea

gig

as

Yes

N

o N

o N

o V

irus-

like

incl

usio

ns

Pin

na

bic

olo

r Y

es

No

No

No

BA

CT

ER

IA

F

infis

h

Aer

om

on

as

salm

oni

cid

a (a

typi

cal)

Ca

rass

ius

aur

atu

s, Cyp

rin

us

carp

io, P

erca

flu

via

tilis,

Sa

lmo

sa

lar

Yes

Y

es

No

Yes

La

cto

cocc

us

ga

rvie

ae

O

nco

rhyn

chu

s m

ykis

s Y

es

Yes

N

o Y

es

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

61

w

ww

.dig

sfis

h.co

m

Dis

ease

ag

ent

Hos

t(s)

that

are

kno

wn

to b

e us

ed a

s ba

it o

r bu

rley

Dis

ease

ag

ent

is

infe

ctio

us

Und

er

offic

ial

cont

rol

List

ed b

y O

IE

or

NA

CA

Like

ly t

o ca

use

sig

nific

ant

dise

ase

Pis

ciri

cket

tsia-

like

orga

nism

s (P

LBs)

of

salm

onid

s S

alm

o s

ala

r Y

es

Yes

N

o Y

es

Yer

sin

ia r

uck

eri

Ca

rass

ius

aur

atu

s, On

corh

ynch

us

myk

iss, S

alm

o s

ala

r Y

es

Yes

N

o Y

es

Cru

stac

eans

Myc

op

lasm

a spp

. P

ena

eus

mo

no

do

n Y

es

No

No

No

Vib

rio

mim

icu

s C

her

ax

qu

ad

rica

rina

tus

Y

es

No

No

No

Ric

kett

sia

-like

org

ani

sms

Ch

era

x q

ua

dri

carin

atu

s, Ch

era

x d

estr

uct

or, C

her

ax s

pp.

Yes

Y

es

Yes

Yes

M

ollu

scs

R

icke

ttsi

a-li

ke o

rga

nism

s H

alio

tis la

evig

ata,

Hal

iotis

ru

bra

, Ha

liotis

spp

., Cra

sso

stre

a g

iga

s, P

inn

a b

ico

lor,

Sa

cco

stre

a g

lom

era

ta Y

es

No

Yes

N

o

FU

NG

I

Fin

fish

E

US

/ Ap

ha

no

myc

es in

vad

an

s

F. A

mba

ssid

ae,

Ca

rass

ius

au

ratu

s, F. C

ichl

ida

e, F

. Ele

otri

dae,

F.

Lutja

nida

e, F

. Mel

ano

taen

iida

e, F

. M

uglid

ae,

N

ema

talo

sa sp

p., F

. S

illa

gini

dae,

F. S

parid

ae,

F. T

era

poni

dae,

T

oxo

tes s

pp

Yes

Y

es

Yes

Y

es

PR

OT

OZ

OA

Fin

fish

E

imer

ia s

pp,

F. A

ther

inid

ae,

F. G

errid

ae,

F. M

uglid

ae,

F. M

onoc

ant

hida

e, F

. S

illa

gini

dae,

Ca

rass

ius

aura

tus, F

. Ter

apo

nida

e Y

es

No

No

No

Go

uss

ia sp

p.

F. C

ara

ngid

ae,

F. S

parid

ae,

F. S

illa

gini

dae,

F.

Poe

cilii

dae

Yes

N

o N

o N

o O

ther

Mic

rosp

orea

F

. Clu

peid

ae,

F. E

leot

rida

e, F

. Eng

raul

ida

e, F

. G

emp

ylid

ae,

F.

Labr

ida

e, F

. Mon

oca

nthi

dae

Y

es

No

No

Yes

Neo

pa

ram

oeb

a spp

. O

nco

rhyn

chu

s m

ykis

s Y

es

No

No

Yes

S

yste

mic

am

oebi

asi

s C

ara

ssiu

s a

ura

tus

Yes

N

o N

o Y

es

Uro

nem

a sp.

, sc

utic

ocili

ate

s F

. Sill

agi

nida

e, F

. Sco

mbr

ida

e Y

es

No

No

Yes

C

rust

acea

ns

A

mes

on s

pp.

Pen

aeu

s m

on

od

on, P

ena

eus

escu

len

tus

, Pen

aeu

s se

mis

ulc

atu

s,

Pen

aeu

s m

erg

uie

nsi

s, P

ort

un

us

pel

ag

icu

s Y

es

Yes

N

o N

o

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

62

w

ww

.dig

sfis

h.co

m

Dis

ease

ag

ent

Hos

t(s)

that

are

kno

wn

to b

e us

ed a

s ba

it o

r bu

rley

Dis

ease

ag

ent

is

infe

ctio

us

Und

er

offic

ial

cont

rol

List

ed b

y O

IE

or

NA

CA

Like

ly t

o ca

use

sig

nific

ant

dise

ase

Hem

ato

din

ium

spp.

P

ort

un

us

pel

ag

icu

s, Scy

lla s

erra

ta Y

es

No

No

Yes

O

ther

Mic

rosp

orid

ia

Ch

era

x d

estr

uct

or, C

her

ax

qu

ad

rica

rina

tus,

Ch

era

x te

nu

ima

nu

s,

C. ca

inii,

Ch

era

x spp

., .M

acr

ob

rach

ium

ros

enb

erg

ii, Pen

aeu

s se

mis

ulc

atu

s, Mel

icer

tus

latis

ulc

atu

s, Pen

aeu

s spp

., Pan

ulir

us

cyg

nu

s, P

an

ulir

us s

pp., P

ort

un

us

pel

agi

cus

Yes

Y

es

No

Yes

Pso

rosp

erm

ium s

pp.

Ch

era

x q

ua

dri

carin

atu

s, Ch

era

x te

nu

ima

nus, C

. ca

inii

Yes

N

o N

o N

o T

hel

oh

an

ia sp

p.

Ch

era

x d

estr

uct

or, C

her

ax

qu

ad

rica

rina

tus,

Ch

era

x te

nu

ima

nu

s,

C. ca

inii,

Ch

era

x spp

., Pen

aeu

s m

on

od

on, P

ena

eus

escu

len

tus

, P

ena

eus

sem

isu

lca

tus

, Pen

aeu

s m

erg

uie

nsi

s, M

elic

ertu

s la

tisu

lca

tus,

Po

rtu

nu

s p

ela

gicu

s

Yes

Y

es

No

Yes

Uni

dent

ified

thr

aus

toch

ytri

d S

cylla

ser

rata

Yes

N

o N

o N

o M

ollu

scs

B

on

am

ia r

ou

ghle

yi S

acc

ost

rea

glo

mer

ata

Y

es

Yes

Y

es

Yes

B

on

am

ia sp

p.

Sa

cco

stre

a g

lom

era

ta

Yes

Y

es

Yes

Y

es

Ha

plos

por

idos

is

Sa

cco

stre

a g

lom

era

ta,

S.

cucc

ulla

ta

Yes

Y

es

No

Yes

M

art

eilia

syd

ney

i (Q

X)

Sa

cco

stre

a g

lom

era

ta

Yes

Y

es

No

Yes

M

art

eilio

ides

bra

nch

alis

Sa

cco

stre

a g

lom

era

ta

Yes

N

o N

o N

o P

erki

nsu

s o

lsen

i, Per

kin

sus

spp.

H

alio

tis la

evig

ata,

Hal

iotis

ru

bra

, Ha

liotis

spp

., Am

usi

um

spp.

, P

ect

en s

pp., K

atel

ysia

spp.

, An

ada

ra s

pp., S

acc

ost

rea

glo

mer

ata

Yes

Y

es

Yes

Y

es

Ste

inh

au

sia

myt

ilovu

m, m

icro

spor

idia

ns

Myt

ilus

edu

lis, S

acc

ost

rea

glo

mer

ata

Yes

N

o N

o N

o

Uni

dent

ified

mic

roce

ll C

rass

ost

rea

gig

as

Yes

Y

es

Yes

Y

es

Ann

elid

a

Ma

rtei

lia s

ydn

eyi (Q

X)

Nep

hty

s a

ust

ralie

nsi

s, Per

iner

eis

nu

ntia,

F. N

epht

yida

e Y

es

Yes

N

o Y

es

ME

TA

ZO

A

F

infis

h

Ben

eden

ia spp

. F

. Ca

rang

ida

e, F

. Lut

jani

dae,

F. S

parid

ae

Yes

N

o

No

Yes

B

oth

rio

cep

ha

lus

ach

eilo

gn

ath

i C

ara

ssiu

s a

ura

tus

, Cyp

rin

us

carp

io, F

. Ele

otri

dae,

F. P

oeci

liida

e, F

. R

etro

pinn

ida

e,

Yes

N

o N

o Y

es

Ca

ligu

s ep

idem

icu

s

F. M

uglid

ae,

F. S

parid

ae

Yes

N

o N

o Y

es

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

63

w

ww

.dig

sfis

h.co

m

Dis

ease

ag

ent

Hos

t(s)

that

are

kno

wn

to b

e us

ed a

s ba

it o

r bu

rley

Dis

ease

ag

ent

is

infe

ctio

us

Und

er

offic

ial

cont

rol

List

ed b

y O

IE

or

NA

CA

Like

ly t

o ca

use

sig

nific

ant

dise

ase

Ca

ma

llan

us

cotti

F. P

oeci

liida

e Y

es

No

No

Yes

C

entr

oce

stu

s spp

. F

. Cic

hlid

ae,

F. P

oeci

liida

e,

Yes

N

o N

o Y

es

Clin

ost

om

um

spp.

F

. Am

bass

ida

e, F

. E

leot

rida

e, F

. Mel

ano

taen

iida

e, F

. Ter

apo

nida

e Y

es

No

No

N

o D

act

ylo

gyr

us s

pp.

Ca

rass

ius

aur

atu

s, Cyp

rin

us

carp

io

Yes

N

o N

o Y

es

Gyr

od

act

ylu

s sp

p.

(Not

G

. sa

lari

s)

Ca

rass

ius

au

ratu

s, F

. E

leot

rida

e,

F.

Lutja

nida

e, M

isg

urn

us

an

gu

illic

au

dat

us,

F. P

oeci

liida

e, F

. Ret

ropi

nnid

ae,

F. S

illa

gini

dae

Y

es

No

No

Yes

Ku

do

a sp

p.

F. C

ara

ngid

ae,

F. C

lupe

ida

e, F

. Eng

raul

ida

e, F

. Ge

mp

ylid

ae,

F.

Sco

mbr

ida

e, F

. Sill

agi

nida

e Y

es

No

No

Yes

Ler

na

ea c

ypri

na

cea

/ Ler

na

ea

spp.

C

ara

ssiu

s a

ura

tus, C

ypri

nu

s ca

rpio,

F. G

ala

xiid

ae

, F

. Poe

cilii

dae,

F

. Ret

ropi

nnid

ae,

F. T

era

poni

dae

Yes

N

o N

o Y

es

Lig

ula

inte

stin

alis

F

. Ga

laxi

ida

e Y

es

No

No

Yes

M

yxo

bo

lus s

pp.

Ca

rass

ius

aur

atu

s, F. E

leot

rida

e, F

. G

ala

xiid

ae

Yes

N

o N

o

Yes

O

ther

Myx

ozoa

F

. Am

bass

ida

e, F

. A

rrip

ida

e, F

. A

ther

inid

ae

, F

. Car

ang

ida

e,

Ca

rass

ius

aur

atu

s, F. C

lupe

ida

e, Cyp

rin

us

carp

io, F

. Ele

otri

dae,

F.

Gem

pyl

ida

e, F

. Ger

rida

e, F

. La

brid

ae,

F. L

ethr

inid

ae,

F.

Lutja

nida

e,

F. M

uglid

ae,

Nem

ata

losa

spp.

, Per

ca f

luvi

atil

is, F

. Ret

rop

inni

dae,

F

. Sco

mbr

ida

e, F

. Sill

agi

nida

e, F

. Spa

rida

e, F

. Ter

apo

nid

ae

Yes

N

o N

o Y

es

Cru

stac

eans

Ca

rcin

on

emer

tes

mits

uku

rii

P

ort

un

us

pel

ag

icu

s Y

es

No

No

No

Olig

ocha

etes

/ P

olyc

haet

es C

her

ax

des

tru

cto

r, Ch

era

x spp

. N

o N

o N

o N

o O

stra

coda

Ch

era

x d

estr

uct

or, C

her

ax

ten

uim

anu

s, C.

cain

ii, C

her

ax s

pp.

No

No

No

No

Sa

ccu

lina s

pp.

Po

rtu

nu

s p

ela

gic

us

Yes

N

o N

o Y

es

Tem

noce

pha

la C

her

ax

des

tru

cto

r, Ch

era

x q

ua

dric

ari

na

tus, C

her

ax

ten

uim

an

us, C

. ca

inii,

Ch

era

x spp

., Eng

aeu

s spp

., Eu

ast

acu

s spp

. N

o N

o N

o N

o

Tur

bella

ria

Po

rtu

nu

s p

ela

gic

us

No

No

No

No

Mol

lusc

s

Bo

cca

rdia

spp.

H

alio

tis la

evig

ata,

Hal

iotis

ru

bra

, Ha

liotis

spp

., Am

usi

um

spp.

, P

ect

en s

pp., C

rass

ost

rea

gig

as, M

ytilu

s ed

ulis

N

o Y

es

No

Yes

Po

lyd

ora

spp.

H

alio

tis la

evig

ata,

Hal

iotis

ru

bra

, Ha

liotis

spp

., Cra

sso

stre

a g

iga

s, M

ytilu

s ed

ulis

, Sa

cco

stre

a g

lom

era

ta, S.

cucc

ulla

ta N

o N

o N

o Y

es

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

64

w

ww

.dig

sfis

h.co

m

Tab

le 9

. D

isea

se a

gent

s no

t inc

lude

d in

Tab

le 8

bec

ause

th

ey a

re u

nlik

ely

to c

ause

ser

ious

dis

ease

and

/or

are

like

ly to

be

ubiq

uito

us.

Viru

ses

Bac

teria

F

ung

i P

roto

zoa

Met

azo

a Ly

mp

hocy

stis

viru

s A

ero

mo

na

s h

ydro

ph

ila A

chyl

a sp

p.

Oth

er C

iliop

hora

A

cant

hoce

pha

la

E

dw

ard

siel

la t

ard

a B

ran

chio

myc

es sp

p.

Am

ylo

od

iniu

m sp

p.

Oth

er C

esto

da

F

lexi

ba

cter

spp.

E

xop

hia

la sp

p.

Cre

pid

oo

din

ium

spp.

O

ther

Cru

sta

cea

F

lavo

ba

cter

ium

spp.

F

usa

riu

m sp

p.

Cry

pto

cary

on

irri

tan

s O

ther

Mon

ogen

ea

Leu

coth

rix s

pp.

Ha

liph

tho

ros s

pp.

Ep

isty

lis s

pp.

Oth

er N

ema

toda

L

act

ob

aci

llus s

pp.

L

ag

enid

ium

spp.

Ic

hth

yob

od

o sp

p.

Tre

ma

toda

Myc

ob

act

eriu

m sp

p.

Pyt

hiu

m sp

p.

Ich

thyo

ph

thir

ius

mu

ltifil

iis

Gre

garin

es

P

seu

do

mo

na

s spp

S

ap

role

gn

ia sp

p.

La

gen

op

hry

s spp

P

inn

oth

eres

spp

T

ena

cib

acu

lum

spp.

Oo

din

ium

spp.

O

cto

lasm

is sp

.

Vib

rio s

pp.

P

isci

no

od

iniu

m sp

p.

Ca

rcin

on

emer

tes s

pp.

T

etra

hym

ena s

pp.

Tri

cho

din

a spp

.

Tri

cho

din

ella

spp.

V

ort

icel

la s

pp.

Zo

oth

am

niu

m spp

.

_________________________________________________________________________________________________________________________________________________________________________________


65

www.digsfish.com

4.2 Elimination of insignificant diseases

Hazard identification for the diseases reported from these commodities identified at least 80 diseases of

potential concern, including 30 viruses, 8 bacterial diseases, 20 protozoan diseases and 21 metazoan diseases

from finfish, crustaceans and molluscs, as well as one fungal disease from finfish (Table 8). However, the

unrestricted risk presented by several of the significant organisms listed in Table 8 that could be translocated

with bait and berley commodities is highly likely to be either within the ALOP or negligible, meaning that

additional risk management measures are not required at this time. Section 4.2 contains a brief discussion of

the reasons why some of these organisms have been excluded from the full risk assessment at this time.

However, it must be considered that knowledge regarding new, and emerging diseases always evolves

rapidly. It is also important to realise that the status of some of the existing disease agents with respect to the

ALOP may change at some time in the future, especially if industries based on movement of live finfish,

crustaceans or molluscs for bait become established, as they have in some other parts of the world (e.g.

USA). Because of this, it is likely that the hazard list, and this RA, will require regular updating to consider

new information on diseases of bait and berley commodities as it becomes available, as well as whenever

there are significant changes to how bait is used in Australia.

4.2.1 Viruses

Finfish

Haematopoietic necrosis herpesvirus of goldfish (Cyprinid herpesvirus-2 (CyHV-2))

Goldfish carry a wide range of disease agents (Diggles 2005). There are several areas of Australia where

populations of goldfish have become established in the wild, and their use as bait is thought to be one avenue

for their introduction and spread (Lintermans 2004). Of the many viral diseases carried by goldfish (Diggles

2005), herpesviruses are amongst the most important, including herpesviral haematopoietic necrosis virus

(CyHV-2) (Jung and Miyazaki 1995). CyHV-2 has been recorded in Australia (Stephens et al. 2004) and it

is known to be widespread in commercial goldfish farms overseas (where most of the goldfish retailed in

Australia originate) at high prevalences (e.g. Goodwin et al. 2009). Hence there is a very high probability

that a high percentage of goldfish used as bait in Australia have been in contact with CyHV-2. However the

narrow host range of CyHV-2, which is known to only cause disease in goldfish, and does not cause clinical

disease in other cyprinids or non-cyprinids (Bergmann et al. 2010). It is not known whether Australian

native fishes are susceptible to CyHV-2, however the narrow host range in cyprinids strongly suggests non-

cyprinids are refractory to infection, indicating that its presence in goldfish used as bait is probably

inconsequential in the Australian context, due to a likely lack of suitable hosts in the wild, hence CYHV-2

will not be considered further in this RA. However, the close relationship between CyHV-2 and other

significant cyprinid herpesviruses (e.g. Koi Herpesvirus CyHV-3) is worth noting (Waltzek et al. 2005).

Indeed, of more significant concern is that goldfish are also now known to be carriers of CyHV-3 (Koi

Herpesvirus), an OIE Listed disease, and that infected carrier goldfish can transmit the infection to naïve

common carp (El-Matbouli and Soliman 2011). CyHV-3 is considered an exotic disease in Australia

(Tables 3a, 3b), and hence will not be considered here.

_________________________________________________________________________________________________________________________________________________________________________________


66

www.digsfish.com

Pilchard herpesvirus (PHV)

Disease caused by a novel herpesvirus caused mass mortalities of up to 75% of the population of pilchards

(Sardinops sagax neopilchardus) throughout the 6000 km range of the species in Australian coastal waters in

1995 and again in 1998/99 (Gaughan et al. 2000; Whittington et al. 2008). Infection of the gill epithelium by

the herpesvirus resulted in severe inflammation, epithelial hypertrophy, and hyperplasia (Whittington et al.

1997). Affected fish died of asphyxiation around 3 to 4 days after initial infection, as evidenced by blood gas

results that indicated hypoxaemia (low oxygen) and hypercapnea (high carbon dioxide) (Whittington et al.

1997). Observations of affected pilchards showed no unusual behavioural changes of fish in the school,

unless they were chased, at which time affected fish would leave the school, begin swimming in an unco-

ordinated manner, and would die within a few minutes (Whittington et al. 1997; BK Diggles, personal

observations of underwater video). The massive extent of the pilchard kills resulted in significant, but

largely unquantified, ecological effects for birds, fishes, and other predators that usually consumed pilchards

(Gaughan et al. 2000). Exposure of a naïve population to an exotic herpesvirus carried by imported frozen

pilchards used as aquaculture feed is considered the most likely cause of both events (Hine and MacDiarmid

1997; Gaughan 2002; Murray et al. 2003; Whittington et al. 2008). While PHV caused significant mortality,

it apparently had high host specificity for pilchards, and was recorded from diseased pilchards throughout the

full range of their distribution in Australia. Because of this, there appears to be little chance that even the

current extensive use of pilchards as bait throughout the country could extend the range of the disease agent

beyond its present distribution, and therefore PHV will not be considered further in this RA.

Aquareoviruses in Atlantic salmon and redfin perch

In Australia, aquareoviruses have been isolated from redfin perch in Victoria, and Atlantic salmon in

Tasmania (Humphrey 1995a, 1995b, AQIS 1999a, 1999b). Both of these species are sometimes used as bait,

however, the discovery of the viruses in each species was incidental, and aquareoviruses are generally not

associated with disease (Roberts 2001). Because these viruses are unlikely to cause disease, they will not be

considered further in this RA.

Crustaceans

Viral diseases of freshwater crayfish

Reviews of the viral diseases of freshwater crayfish were conducted by Evans et al. (1998) and Edgerton et

al. (2002). The relationship between infection and disease remains obscure for the majority of crayfish

viruses (Edgerton et al. 2002). The parvovirus and reovirus observed in Cherax quadricarinatus, as well as

the bacilliform viruses such as Cherax quadricarinatus bacilliform virus (CqBV), Cherax destructor

bacilliform virus (CdBV), and other bacilliform viruses observed in Cherax tenuimanus and C. cainii

appeared to be low virulence and were not always associated with disease, or in the case of CdBV, the one

study reported was too limited to assess whether it caused disease (Edgerton et al. 2002). Because

transmission experiments have not been performed, the full host range of these viruses is unknown and it is

not known whether they pose a threat to the health of other, rare and threatened native crayfish (Evans et al.

1999, Edgerton et al. 2002). Because they do not appear to be directly associated with disease in their

natural hosts, the viruses mentioned above will not be considered in the RA. However, there are three

_________________________________________________________________________________________________________________________________________________________________________________


67

www.digsfish.com

viruses reported from cultured freshwater crayfish that were associated with disease in juveniles

(Giardiavirus-like virus of Cherax quadricarinatus (CGV)), moribund adults (Cherax destructor systemic

parvo-like virus (CdSPV)), or in both juveniles and adults (a parvovirus in C. quadricarinatus (CqPV)).

Because of their association with disease, it is possible that if CGV, CqPV and /or CdSPV were translocated,

they could represent a threat to the health of other crayfish (including rare and threatened native freshwater

crayfish, see Coughran and Leckie 2007, Coughran et al. 2009) or even other crustaceans. Because of this,

CGV, CqPV and CdSPV will be included in the RA as examples of the viral diseases potentially translocated

in freshwater crayfish used as bait.

Viral diseases of prawns

Prawns harbour a wide variety of disease agents, including several viruses that have been particularly

problematic in aquaculture (Biosecurity Australia 2009). However, there are also several viruses of prawns

in Australia which have not been associated with clinical disease, or which are readily controllable using

simple methods such as routine washing of eggs and nauplii in clean seawater. These include a BMNV-like

virus (Lightner 1996), a haemocytic rod shaped virus (Owens 1993), and a lymphoid-parvo-like virus (LPV)

(Owens et al. 1991). A non-occluded bacilliform virus in Australian prawn species was reported by Lightner

(1996) as a BMNV-like virus. The term BMN-type virus is misleading as it gives the impression that this

virus was similar to BMNV that infects Penaeus japonicus in Japan (Lightner 1996), and the term non-

occluded bacilliform virus is more correct. This virus has not been reported in the literature for over 20

years, suggesting that it is not associated with significant disease, and hence it will not be considered in the

RA. Epizootic losses of hybrid Penaeus esculentus x P. monodon were associated with the presence of a

haemocytic rod shaped virus (PHRV) in the gill tissues (Owens 1993). However, a strain of Infectious

Hypodermal and Haematopoietic Necrosis Virus (IHHNV) was also present, and it is not clear that PHRV

was responsible for disease, as the concurrent infection with IHHNV may have lead to an expression of an

otherwise latent virus (Owens 1993). A lack of reports on this virus in recent years suggests it does not

cause significant disease, and hence it will not be considered in the RA. LPV was observed by Owens et al.

(1991) to occur in lymphoid organ, antennal gland, and nerve cord of moribund wild spawners of Penaeus

merguiensis, but was also observed in samples from apparently healthy farmed P. monodon, P. merguiensis

and P. esculentus, as well as P. esculentus x P. monodon hybrids (Owens et al. 1992). LPV appears closely

related to IHHNV (Owens et al. 1991, Lightner 1996), however IHHNV is a much more pathogenic virus

and was most likely responsible for disease in the instance where both were present together in the same

hosts (Owens et al. 1992). Given that there is no evidence that LPV has ever directly caused significant

disease, it will not be considered in the RA.

Viral diseases of crabs

A non-occluded baculovirus (Scylla baculovirus, SBV) was found in juvenile, sub adult and adult mudcrabs

(Scylla serrata) from the Northern Territory (Anderson and Prior 1992). The virus did not cause clinical

disease, the infected crabs were apparently healthy and feeding well, and the distribution of infected

hepatopancreatic epithelial cells was focal. Owens et al. (2010) reported the presence of a similar virus in

broodstock and larvae of S. serrata from near Townsville, North Queensland. There has been no evidence to

date that SBV causes disease in mudcrabs or other crustaceans. Given that there is no evidence that SBV has

ever caused significant disease, it will not be considered in the RA.

_________________________________________________________________________________________________________________________________________________________________________________


68

www.digsfish.com

Molluscs

Molluscs can harbour a range of viruses (Elston 1997). Digestive epithelial virosis associated with small

RNA viruses was first reported in bivalves in New Zealand, where they were associated with mortalities of

mussels, scallops and toheroa (Jones et al. 1996, Hine and Wesney 1997). Histological lesions, similar to

those in New Zealand, were reported from scallops (Pecten alba) from Port Phillip Bay, Victoria, and pearl

oysters (Pinctada maxima) in northern Western Australia, but the presence of virus was not confirmed

(AFFA 2002). There is some confusion over the identity and significance of these small viruses. They have

been associated with digestive sloughing during outbreaks of mortality (Jones et al. 1996, Hine and Wesney

1997) but sloughing of digestive cells also occurs in the absence of disease, and Koch's postulates have not

been fulfilled (Hine and Wesney 1997). The viral particles may be irrelevant to digestive epithelial

degeneration, or they may alter the kinetics of degeneration and renewal, leading to disease (Hine and

Wesney 1997). Because it is not clear that these viruses cause significant disease at this time, they will not

be considered further in this RA.

Viral gametocytic hypertrophy caused by papova-like viruses (Choi et al. 2004) has been recorded in an ever

increasing assortment of bivalve hosts in an increasing number of areas around the world (Garcia et al. 2006,

Cheslett et al. 2009). These viruses cause massive hypertrophy of individual gametes and gametogenic

epithelium by replicating in the host cell nucleus. Host response to infection is usually negligible and the

infection is not known to cause disease or reduced fecundity. In Australia the condition has been reported

from Pacific oysters (Crassostrea gigas), Sydney rock oysters (Saccostrea glomerata), and pearl oysters

(Pinctada maxima) in locations such as QLD, NSW, WA, SA and Tasmania (AFFA 2002, B. Diggles,

personal observations). Apparent increased prevalence of viral gametocytic hypertrophy in recent years

could be due to spread of the disease, increased surveillance of molluscs, increased recognisiton of the

disease by diagnosticians, or a combination of these factors (Garcia et al. 2006). As these viruses do not

appear to have any detrimental effect on the host, and because they already appear to be widely distributed,

they will not be considered further in this RA.

Hine and Thorne (2000) found intranuclear virus-like particles in 1 out of 71 Pinna bicolor (including 65

spat) examined from Dampier archipelago in Western Australia. Eosinophillic inclusions occurred in the

nuclei of diverticular epithelial cells (Hine and Thorne 2000). The affected bivalves were apparently

healthy, and the virus particles were not associated with disease, and because of this, this virus will not be

considered further in this RA.

4.2.2 Bacteria

Crustaceans

Mycoplasmas have been recorded from penaeid prawns in Australia (Biosecurity Australia 2009), however

their presence in infected prawns is not usually associated with serious disease (Biosecurity Australia 2009),

and hence will not be considered further in this RA. However, there are some important bacterial diseases of

crustaceans, including some that are listed by the OIE and NACA, that are associated with infection by

related rickettsial disease agents, hence these will be considered in more detail in the RA.

_________________________________________________________________________________________________________________________________________________________________________________


69

www.digsfish.com

Vibrio mimicus is a bacterium that was implicated in mortalities of Cherax quadricarinatus during

significant mortality events in aquaculture ponds in SE QLD and northern NSW (Eaves and Ketterer 1994).

However, the bacterium was considered to be part of the normal bacterial flora of the aquatic environment in

aquaculture ponds and appeared to be an opportunistic pathogen causing disease only after crayfish were

stressed from overcrowding or poor water quality (Eaves and Ketterer 1994). Because of this, V. mimicus

will not be considered further in this RA.

Molluscs

Procaryote rickettsia-like organisms (RLOs) and chlamydia-like organisms (CLOs) are commonly observed

in the gills and digestive tract of wild and cultured molluscs in Australia and worldwide (Hine and Thorne

2000, AFFA 2002). These agents are sometimes associated with disease (especially when they occur in the

branchial epithelium of scallops), though they are also commonly present in apparently healthy molluscs

(Hine and Diggles 2002a). The taxonomy of RLOs and CLOs remains poorly defined and in bivalves it is

not known whether bacteria infective to different hosts are different species, or the same species with

different levels of pathogenicity in different hosts (AFFA 2002). These bacteria have been noted to infect

scallops, oysters, cockles, clams and mussels worldwide, however in bivalves they have been noted to be

associated with disease only in scallops (Hine and Diggles 2002a, AFFA 2002). However, in univalves

Candidatus Xenohaliotis californiensis, the agent responsible for the OIE and NACA-listed disease

Withering Syndrome of abalone, is also a RLO (Gardener et al. 1995, Friedman et al. 2000). RLOs and

CLOs are common in many bivalve hosts in Australia, and RLOs have also been observed in Australian

abalone (Handlinger et al. 2006). Furthermore, while X. californiensis is an important pathogen of abalone,

and has been spread via movements of infected abalone to several Asian countries (Wetchateng et al. 2010),

this pathogen is considered to be exotic to Australia. Because of these reasons, RLOs from molluscs will not


4.2.3 Fungi

None excluded

4.2.4 Protozoa

Finfish

Coccidia of finfish and crustaceans (Eimeria spp., Goussia spp.)

Coccidians are members of the Apicomplexa (Lom and Dykova 1992). A variety of species of finfish used as

bait in Australia are known to harbour coccidian infections, including Families Atherinidae, Carangidae,

Poeciliidae, Gerridae, Muglidae, Monocanthidae, Sillaginidae, Sparidae, Teraponidae and also Carassius

auratus (see Lom et al. 1992, Lom and Dykova 1995, Molnar and Rohde 1988a, 1988b). The distribution of

many of these parasites in their teleost definitive hosts and other intermediate hosts (e.g. crustaceans) is

largely unknown, however they occur naturally in the internal organs of apparently healthy fish hosts at

_________________________________________________________________________________________________________________________________________________________________________________


70

www.digsfish.com

prevalences of up to 100% (Lom et al. 1992, Lom and Dykova 1995, Molnar and Rohde 1988a, 1988b).

Given that only a relatively small number of fish and crustacean species have been specifically examined for

coccidia in Australia, it is likely that many more species would be discovered if systematic investigations

were undertaken (Molnar and Rhode 1988a). Fish coccidia can cause disease in instances where their hosts

are held at high densities in confinement and the life cycle can be completed (Lom and Dykova 1992).

However, because it appears that these parasites do not have any detrimental effects on their wild hosts in

Australia (either fish or crustaceans), and because they already appear to be widely distributed, they will not


Neoparamoeba spp.

Some salmonid products are used as bait, and it is known that cultured salmonids in Tasmania are adversely

affected by amoebic gill disease (AGD), which is caused by infection of the gills with Neoparamoeba spp.

Recently the aetiological agent of the disease has been identified as Neoparamoeba perurans (Young et al.

2007), a species which was also found in archived samples, confirming that it has been the predominant

aetiological agent of AGD in Tasmania since epizootics were first reported there. Recent studies have

confirmed that N. perurans is a free-living amoeba that occurs naturally in the rearing environment around

seacages containing cultured salmonids not only in Tasmania (Bridle et al. 2010), but in several salmonid

hosts in at least 6 different countries (Young et al. 2008). It appears that these parasites are opportunistic

pathogens that cause disease only in cultured salmonids, and cultured salmonids and their processing wastes

are not sold for bait or berley (Grant Pullen, DPIWE Tasmania, personal communication 8 April 2011),

although fresh and frozen salmon products are widely sold as food fish and some of these products could be

diverted for use as bait. Furthermore, these agents are free living and already appear to be widely

distributed, and because of these reasons, they will not be considered further in this RA.

Crustacea

Psorospermium spp.

Psorospermium spp. are unicellular organisms that parasitize freshwater crayfish. Members of the genus

have been found in 17 species of freshwater crayfish worldwide (Bangyeekhun et al. 2001). A species of

Psorospermium has been identified from freshwater crayfish in Australia (Herbert 1987), however the

pathogenicity of this parasite to the crayfish host is not clear (Edgerton et al. 2002). Some workers have

reported a lack of haemocytic reaction to the organism and taken this to imply little or no pathogenic effect

(Herbert, 1987), however other analysts have stated that the presence of high Psorospermium loads could

increase susceptibility to other pathogens or disease-causing agents (Edgerton et al. 2002). Because it is not

clear that these parasites can cause disease, they will not be considered further in this RA.

Unidentified thraustochytrid of mud crabs

Kvingedal et al. (2006) described a new parasite that infected the eggs of captive Scylla serrata. The

parasite was discovered during a broodstock research program and caused 100% egg mortality (Kvingedal et

al. 2006). DNA analysis indicated that the parasite was related to the thraustochytrids. The origin of the

_________________________________________________________________________________________________________________________________________________________________________________


71

www.digsfish.com

parasite was not determined. A variety of saprophytic organisms, particularly fungi, are opportunistic

pathogens of eggs of crustaceans reared in captivity, including mud crabs (Nakamura et al. 1995, Leano

2002), prawns (Lio-Pio and Sanvictores 1986), crabs (Sparrow 1973), freshwater crayfish (Herbert 1987)

and spiny lobsters (Shields et al. 2006, BK Diggles, personal observations). Infection of eggs by saprophytic

organisms is therefore common, and these organisms tend to originate from the natural environment where

they are usually associated with substrates such as sediments (Kvingedal et al. 2006). Because these

infections are caused by opportunistic pathogens, some of which can be relatively easy to control through

chemotherapy or changes in husbandry practices (e.g. Hamasaki and Hatai 1993), saprobes of crustacean

eggs will not be considered further in this RA.

Molluscs

Steinhausia mytilovum is a microsporidian that has been reported from blue mussels (Mytilus

galloprovincialis) in Cockburn Sound, WA, but not in other mussel growing areas (Jones and Creeper 2006).

Steinhausia –like microsporidians have also been recorded from Sydney rock oysters (Saccostrea glomerata)

in Moreton Bay, QLD (Anderson et al. 1995) and Crassostrea echinata from the Northern Territory (Wolf

1977). Steinhausia mytilovum is a globally distributed microsporidian parasite which infects the oocytes of

blue mussels Mytilus edulis and M. galloprovincialis (See Comtet et al. 2004). These parasites can affect the

condition index of infected mussels (Rayyan and Chintiroglou 2003) and sometimes induce marked

haemocytic infiltration inside affected gonad follicles (e.g. Jones and Creeper 2006), but no conclusive

evidence has been reported regarding the viability of the infected oocytes, or mortality of the host, and

therefore the effect of Steinhausia mytilovum on its host remains unclear (Comtet et al. 2004). Their known

distribution suggests that Steinhausia-like parasites are already distributed in several species of molluscs

throughout Australia. This, combined with the fact that it is unclear whether these parasites can cause

disease, means they will not be considered further in this RA.

4.2.5 Metazoa

Crustaceans

Crustaceans can harbour a wide variety of metazoan organisms (Bower et al. 1994, Shields et al. 2006),

including the nemertean egg predator Carcinonemertes spp., oligochaetes and polychaetes in the gills and on

the carapace, ostracods, copepods, temnocephalans and other turbellaria, as well as digeneans, cestodes and

several other groups listed in Table 9. These groups are mostly free living commensals, symbionts or

parasites that are not known to cause disease, while Carcinonemertes are semi-parasitic egg predators which

do not affect the health of the host, but may reduce fecundity in very high intensity infections in confined

crustaceans, though the infections can be easily treated (Shields et al. 2006). Because of these reasons, they

will not be considered further in this RA

4.3 The diseases of concern to be considered in the RA

The remaining 44 diseases that have been recorded from species on the commodity list (Table 2) are listed in

Table 10 as diseases of concern. Detailed risk assessments will be conducted on these disease agents. As

_________________________________________________________________________________________________________________________________________________________________________________


72

www.digsfish.com

mentioned previously, there are significant knowledge gaps in relation to disease agents that infect aquatic

animals in Australia (Appendix 1). Because of this, there remains a significant risk of transfer of as yet

unknown disease agents, even in the absence of their identification (Gaughan 2002), so an example of

translocation of a new, unknown virus in a finfish commodity is also included (Table 10) so that a scenario

relating to the assessment of risks posed by an unknown disease agent can be explored.

Table 10. The list of diseases of concern to be considered in the detailed risk assessment.

Disease/Disease agent

Disease agent is

infectious

Agent or strains

confined to certain regions

Under official control

OIE-

NACA listed

Expected to cause

significant disease

FINFISH Viruses Aquatic birnavirus � ? � � Epizootic Haematopoietic Necrosis Virus (EHNV)

� � � � �

Viral Encephalopathy and Retinopathy (Nodavirus)

� � � � �

An new unknown virus of finfish � ? ? Bacteria Aeromonas salmonicida (Atypical) / Goldfish ulcer disease

� � � �

Lactococcus garvieae � � � �

Piscirickettsia-like organism (PLBs) of salmonids

� � � �

Yersinia ruckeri � � � �

Fungi Aphanomyces invadans (EUS) � � � � � Protozoans Microsporeans � ? � Systemic amoebiasis of goldfish � ? � Uronema sp, Scuticociliates � ? � Metazoans Benedenia spp. � � � Bothriocephalus acheilognathi � � � Caligus epidemicus/sea lice � ? � Camallanus cotti � ? � Centrocestus spp. � ? � Dactylogyrus spp. � ? � Gyrodactylus spp. � ? � Kudoa spp. and other myxozoa � ? � Lernaea cyprinacea / Lernaea spp. � ? � Ligula intestinalis � ? � CRUSTACEANS Viruses Cherax destructor systemic parvo-like virus (CdSPV)

� ? ?

Giardia-like virus (CGV) � ? ? Gill Associated Virus (GAV) � � � � � Hepatopancreatic Parvovirus (HPV) � � � Infectious Hypodermal and Haematopoietic Necrosis Virus (IHHNV)

� � � � �

Monodon Baculovirus (MBV) � � � � Mourilyan Virus (MoV) � � � Spawner Isolated Mortality Virus (SMV) � � � �

_________________________________________________________________________________________________________________________________________________________________________________


73

www.digsfish.com

Disease/Disease agent

Disease agent is

infectious

Agent or strains

confined to certain regions

Under official control

OIE-

NACA listed

Expected to cause

significant disease

Macrobrachium rosenbergii Nodavirus (MrNV)

� � � � �

Bacteria Rickettsia-like Organisms � ? � � � Protozoans Hematodinium spp. � � �

Thelohania and other microsporidians � ? � �

Metazoans Sacculina spp. � � � �

MOLLUSCS Viruses Abalone Viral Ganglioneuritis (AVG) � � � � � Protozoans Bonamia roughleyi � � � � � Bonamia spp. � � � � � Haplosporidosis � � � � Marteilia sydneyi (QX disease) � � � � Perkinsus olseni / Perkinsus spp. � � � � � Unidentified microcells � � � � Metazoans Boccardia spp. ? � � Polydora spp. ? � ANNELIDS Protozoans Marteilia sydneyi (QX disease) � � � � �

_________________________________________________________________________________________________________________________________________________________________________________


74

www.digsfish.com

5.0 Detailed Risk Assessment

5.1 Infection of finfish and molluscs with Aquatic Birnavirus

5.1.1 Aetiologic agent: Non-enveloped viruses with a double-stranded RNA genome of the genus

Aquabirnavirus within the Family Birnaviridae.

5.1.2 OIE List: No NACA List: No

5.1.3 Australias status: Tasmanian aquabirnavirus (TAB) is an aquatic birnavirus strain (IPNV Genogroup

5) that has been reported from cultured Atlantic salmon and various other species of marine fish in Tasmania

(Crane et al. 2000, Davies et al. 2010b). TAB is listed as a reportable disease in Tasmania (Table 3b).

5.1.4 Epidemiology

Aquatic birnaviruses have been isolated from a large number of marine and freshwater aquatic animals

(McAllister 1993), to the extent that these viruses are considered to be ubiquitous in aquatic environments

worldwide (Reno 1999). Various strains of birnavirus have been described from at least 65 species of fish in

20 families (McAllister 1993), and also from bivalve molluscs and crustaceans (Reno 1999). The type

species for the genus Aquabirnavirus is Infectious Pancreatic Necrosis Virus (IPNV), which causes

infectious pancreatic necrosis (IPN), a significant disease of salmonids (Wolf et al. 1960). The genus

includes both virulent and avirulent viruses with the term ‘infectious pancreatic necrosis’ (IPN) virus being

reserved for those isolates that are pathogenic for species within the Family Salmonidae (McColl et al.

2009). IPN disease is not known to exist in Australia (Herfort 2004), however in 1998 an aquatic birnavirus

was identified in farmed Atlantic salmon (Salmo salar), and wild rainbow trout (Oncorhynchus mykiss),

greenback flounder (Rhombosolea tapirina), cod (Pseudophycis sp.), spiked dogfish (Squalus megalops) and

ling (Genypterus blacodes) in Macquarie Harbour in western Tasmania (Crane et al. 2000). The Tasmanian

aquabirnavirus (TAB) is related to an aquabirnavirus found in marine fish in New Zealand (Tisdall and

Phipps 1987, Davies et al. 2010b). Both isolates appear to have low pathogenicity for salmonids (Crane et

al. 2000, Davies et al. 2010b), and both have never been associated with mortalities in freshwater hatcheries

(McColl et al. 2009). However, a birnavirus was found to be associated with a suspicious outbreak of

disease in juvenile turbot in New Zealand (Diggles et al. 2000) several years after the event (BK Diggles

unpublished), and it is possible that the Australasian birnavirus isolates may be pathogenic in non-salmonid

hosts or even to salmonids under different environmental or husbandry conditions (such as juvenile fish in

hatcheries) (Davies et al. 2010b).

In Japan aquatic birnaviruses cause some of the most important diseases of juvenile yellowtail (Seriola

quinqueradiata), kingfish (S. lalandi aureovittata) and amberjack (S. dumerili) (see Isshiki and Kusuda

1987, Isshiki et al. 2001, Nakajima et al. 1998, Muroga 2001). This suggests that cultured kingfish in

Australia are also likely to be susceptible to aquatic birnaviruses (AQIS 1999b). Aquatic birnaviruses are

known to cause disease almost exclusively in juvenile fish (Novoa et al. 1993, Reno 1999), with yellowtail

less than 10 grams being particularly susceptible in Japan, with moribund juveniles typically exhibiting

anaemic gills, haemorrhaging in the liver, severe ascites, and pancreatic necrosis (Nakajima et al. 1998).

_________________________________________________________________________________________________________________________________________________________________________________


75

www.digsfish.com

Infection is direct via horizontal exposure to viral particles in the water, or by vertical transmission from

infected gametes (McAllister 1999). Juvenile fish that survive infection can be lifelong carriers which shed

the virus via the urine, faeces and sexual products (Reno 1999), however large juveniles and adults exposed

to the virus for the first time may be refractory to infection or spontaneously recover (Novoa et al. 1993), but

may still become infected if they are stressed or compromised by sub-optimal environmental conditions.

Water-born birnaviruses accumulated by bivalves, crustaceans and birds can remain viable when excreted

(Mortensen et al. 1992), and can be subsequently used to infect fish experimentally (Mortensen 1993),

although viral replication does not appear to occur in other hosts, hence the main method of translocation

remains live fish and eggs (Reno 1999).


The TAB continues to be isolated from marine fish on a regular basis in Tasmania (Davies et al. 2010b).

While the TAB found in 2002 was detected during routine health surveillance of cultured seacaged salmon,

this isolate was not recovered from a pinhead fish as was the case with the original isolate in 1998 (Crane et

al. 2000). The isolate in 2002 was from healthy Atlantic salmon, showing no overt signs of disease (Davies

et al. 2010b). Of the other species known to harbour the TAB, only rainbow trout (Oncorhynchus mykiss) is

listed as being used as bait (Tables 2, 8). However, bivalves can also harbour birnaviruses and are effective

carriers (Rivas et al. 1993, Reno 1999), and may act as vectors and/or reservoirs of infection (Rivas et al.

1993, Kitamura et al. 2007). Kitamura et al. (2007) used PCR to determine that aquatic birnavirus was

present in blue mussels Mytilus galloprovincialis at prevalences of between 80 and 100% during the summer

months, with these prevalences being 8 to 10 times higher than those found in susceptible finfish in the same

area at the same time of year. It is therefore possible that bivalves (especially from Tasmania) also could

harbour aquatic birnaviruses, though it appears that the required surveys have not been conducted to confirm

or rule out this possibility.

The highest titres of birnavirus in infected fish occur in the internal organs such as the pancreas, spleen,

kidney, liver and gastrointestinal tract, with low levels present in the fillet (Reno 1999). Aquabirnaviruses

are very persistent in the environment, with minimal loss of infectivity after 10 weeks in filtered seawater at

4 and 10°C, and they are also very resistant to a broad range of disinfectants (Bovo et al. 2005).

Aquabirnaviruses are very tolerant of freezing, with Inouye et al. (1990) finding no loss of infectivity of

IPNV after freezing for 56 days at -20°C, although each freeze/thaw cycle from that temperature slightly

reduces the quantity of viable virus (Mortensen et al. 1998).

Currently it is unknown whether aquatic birnaviruses occur in the waters of mainland Australia, however

there are no barriers to movement of wild fish from areas of Tasmania where aquatic birnavirus is known to

occur in wild fish, and some fish species do move from these areas into mainland waters (e.g. Stanley 1978).

It must also be recognized that aquatic birnavirus is not inactivated by passage through the bird digestive

system and as such, the disease agent can also be spread naturally via mechanical vectors such as sea birds

(Reno 1999). Because of this, aquatic birnaviruses may also occur in the waters of mainland Australia and

indeed, they are considered likely to be present throughout the Southern Ocean (Davies et al. 2010b), though

the required surveys have not been conducted to confirm or rule out this possibility. Besides salmonids and

bivalves, the literature suggests that flatfish (e.g. greenback flounder in Australia) appear particularly

common carriers of aquatic birnaviruses, however no flatfish species are regularly used as bait in Australia.

_________________________________________________________________________________________________________________________________________________________________________________


76

www.digsfish.com

Other groups known to be susceptible to infection by aquatic birnaviruses that are used as bait in Australia

include atherinids, carangids, clupeids, cyprinids (including goldfish), redfin perch, and sparids (Herfort

2004), however birnavirus has not been recorded from any of these groups in Australia to date. Hence based

on the information presently available, there is a likelihood that aquatic birnaviruses could be translocated

via bait only in salmonid products and bivalves. Cultured salmonids and their processing wastes are not sold

for bait or berley (Grant Pullen, DPIWE Tasmania, personal communication 8 April 2011), but they are

widely distributed as food fish and hence some product could be diverted to use as bait or berley around the

country. Kewagama Research (2002, 2006) estimated that 0.4-0.5% of recreational fishers in NSW, Victoria

and ACT used salmonids products as bait, but between 30 and 33% of recreational fishers throughout all

states used bivalve molluscs as bait, their use being most popular in SA, followed by Victoria, QLD and

Tasmania. While there is evidence that some bivalves may have been translocated from the eastern states

into SA for certain competitive fishing events (Sunfish 2008), it is unclear whether bivalves from Tasmania

are being moved to mainland Australia for the same purpose. It is known, however, that processing waste

from scallops (Amusium spp., Pecten spp., Family Pectinidae) is sometimes used in berley blocks (Horvat

2010). Taking into account the quantities of salmonid products and bivalves used as bait or berley (Table 1),

as well as the prevalence and tissue trophism of the disease agent (mainly in internal organs), the likelihood

estimations for the occurrence of aquatic birnavirus in these commodities are listed below.

Release assessment for infection of finfish and molluscs with Aquatic Birnavirus

Commodity

type

Live

finfish

Whole

fresh

dead

finfish

Frozen

whole

finfish

Frozen

fillets

Frozen

fish

heads

Frozen

guts/offal

Live

bivalve

molluscs

Frozen

bivalve

molluscs

Likelihood of

release

Moderate Moderate Moderate Low Low Moderate Moderate Moderate


Marine teleosts and bivalves in Tasmania are already at risk of exposure to the local strain of aquatic

birnavirus. However, it is not known whether teleosts and bivalves in mainland Australia (both marine and

freshwater environments) are naturally exposed to these disease agents. Specific data on production and use

of bait and berley from Tasmania is not available (Grant Pullen, DPIWE Tasmania, personal communication

7 February 2011), however throughout Australia, fresh or frozen whole or processed bait or berley products

are offered for sale via wholesale outlets such as commercial fishing co-operatives, or packaged for resale at

retail outlets such as fishing tackle shops, service stations and supermarkets. Live bait in Australia is usually

only available for sale at specialist fishing tackle shops, being generally restricted to polychaetes and rarely,

some molluscs obtained in small quantities from local suppliers. Finfish for use as live bait are generally

unavailable at retail outlets (except ornamental species sold from pet shops), and most live bait is collected

by recreational fishers at the fishing site. So the quantities of fish and bivalves translocated as live bait is

likely to be small. The majority of the volume of bait translocated throughout the country is frozen,

including frozen whole fish and processed fish products (e.g. heads or guts), and frozen whole bivalve

molluscs. Fresh or frozen Atlantic salmon are also widely sold as food fish throughout Australia, and these

_________________________________________________________________________________________________________________________________________________________________________________


77

www.digsfish.com

could be diverted to use as bait or berley in both freshwater and marine environments around the country.

Because aquatic birnavirus is highly resistant to freezing, and a large percentage of virus is likely to remain

viable after thawing, this suggests that direct pathways exist for translocated bait from these retail outlets to

enter both freshwater and marine environments, potentially exposing wild fish and molluscs to viable aquatic

birnaviruses. Other significant pathways for exposure of fish and molluscs to viable aquatic birnaviruses

would include disposal into natural waters of untreated processing wastes from commercial processing

premises, and use of processing wastes in berley blocks.

Birnavirus infections have been reported from a wide range of species of fish and bivalves, including species

groups that are known to occur in Australian waters. Many species of wild fish and molluscs in Australia are

therefore likely to be susceptible to infection by aquatic birnaviruses in infected bait or berley, but infection

would occur only if sufficient quantities of virus (i.e. an infective dose) was introduced into an area where

susceptible hosts were present. Susceptible fish and bivalves can become infected with aquatic birnavirus

via horizontal transmission through the water (immersion) and also by per-os exposure (Reno 1999). The

infectious dose of birnavirus by the immersion pathway varies according to the strain of virus used and the

species challenged (McAllister and Owens 1995), ranging from more than 103 TCID50/mL for Arctic char

(McAllister et al. 2000), to as low as < 10-1TCID50/mL for Atlantic salmon post smolts exposed to

pathogenic strains of IPNV (Urquhart et al. 2008). The infectious dose of birnavirus by per-os exposure to

infected feed also varies, with Mortensen (1993) requiring a dose of 106 TCID50/g of IPNV obtained from

scallops before successful transmission to brown trout was obtained. However Wechsler et al. (1987)

reported successful transmission of IPNV to striped bass fed brook trout infected with between 2 x 102 and

2 x 105 TCID50/g IPNV.

Clinically diseased fish infected with birnavirus can have very high viral titres in their internal organs (107 -

109 TCID50/g,) as well as high viral shedding rates (Reno 1999, Sommer et al. 2004, Urquhart et al. 2008),

however no fish with clinical disease caused by birnavirus infection have ever been recorded in Australia

(Crane et al. 2000, Davies et al. 2010b). The levels of birnavirus in sub-clinically infected fish can still be

relatively high (102 - 106 TCID50/g , see McAllister et al. 2000), but are usually lower and often around the

limits of detection using cell culture techniques (c. 101 - 102 TCID50/g, Wechsler et al. 1987, Roberts 2001).

Even given that susceptible fish can be infected by immersion at low infective doses (Urquhart et al. 2008),

the likelihood that an infectious dose can be transmitted horizontally into a natural water body via use of bait

that was sub-clinically infected with aquatic birnavirus appears unlikely, unless unusually high volumes of

bait are used in one location. One specific end use where this could occur is use of bait as aquaculture feed,

however this end use is not within the scope of this RA. Nevertheless, fish may still become infected by the

per-os route if they eat whole fish containing virus levels typical of subclinically infected fish (Wechsler et

al. 1987). Given that pathways may exist for translocation and spread of birnavirus into the environment via

use of bait or berley, and acknowledging the broad range of susceptible hosts, the risk of exposure and

establishment is non-negligible, and the likelihood of exposure and establishment of aquatic birnavirus in

new fish and mollusc populations is considered to be Low.


When fish become infected with aquatic birnavirus, mortality is mainly restricted to larval and early juvenile

stages, and disease does not necessarily occur in larger fish. Indeed, many fish experimentally infected with

_________________________________________________________________________________________________________________________________________________________________________________


78

www.digsfish.com

aquatic birnavirus can naturally resolve the infection provided they are healthy and remain unstressed (Reno

1999). However, others fish remain carriers for life, and risk spreading birnavirus to their progeny vertically

via infected gametes. Given that aquabirnavirus is already present in at least some parts of the Australian

environment, and these viruses are unlikely to cause disease in juvenile and adult fish in the wild, the main

consequences appear those related to possible increased mortality of larval and early juvenile stages, which

although never documented in wild fishes, if it occurs it could have some impact on wild fish at the

population level. These viruses may also increased cost of production in finfish aquaculture hatcheries due

to use of infected wild caught broodstock. These viruses are no longer listed by the OIE and hence their

presence is unlikely to have adverse impacts on national or international trade. Considering all of these

factors, establishment of the disease would have mild to moderate biological consequences, which would be

amenable to control, and would not cause irreversible environmental effects. It is therefore estimated that

the consequences of introduction of birnavirus strains into different parts of the Australian environment via

use of bait or berley would likely be Low.


The unrestricted risk associated with aquatic birnavirus is determined by combining the likelihood of release

and exposure (from Table 5) with the consequences of establishment (Table 7). The unrestricted risk

estimate for aquatic birnavirus does not exceed the ALOP for any of the commodity types, suggesting that

additional risk management for this disease agent is not required at this time.

Risk estimate for infection of finfish and molluscs with Aquatic Birnavirus

Commodity

type

Live

finfish

Whole

fresh

dead

finfish

Frozen

whole

finfish

Frozen

fillets

Frozen

fish heads

Frozen

guts/offal

Live

bivalve

molluscs

Frozen

bivalve

molluscs

Combined

likelihood of

release and

exposure

Low Low Low Very low Very low Low Low Low

Consequences

of

establishment

Low Low Low Low Low Low Low Low

Risk

estimation

Very

Low

Very

Low

Very Low Negligible

risk

Negligible

risk

Very Low Very

Low

Very

Low

_________________________________________________________________________________________________________________________________________________________________________________


79

www.digsfish.com

5.2 Infection of finfish with Epizootic Haematopoietic Necrosis Virus (EHNV)

5.2.1 Aetiologic agent: EHNV, a virus with a double stranded DNA genome of the genus Ranavirus, in the

Family Iridoviridae.

5.2.2 OIE List: Yes NACA List: Yes

5.2.3 Australias status: IHNV has been reported in Victoria, NSW, ACT and SA (Whittington et al.

2010), and it is a reportable disease in all states excluding the ACT (Table 3b)

5.2.4 Epidemiology

Epizootic haematopoietic necrosis virus (EHNV) is a member of the Ranavirus genus (Family Iridoviridae)

and has a known distribution confined to freshwater areas in south eastern Australia, where it causes

mortalities in wild redfin perch (Perca fluviatilis) and farmed rainbow trout (Oncorhynchus mykiss)

(Langdon et al. 1986, 1988, Langdon and Humphrey 1987, Whittington et al. 1996, 1999). Disease due to

natural EHNV infections are known from only two teleosts, redfin perch and rainbow trout (Langdon et al.

1986, 1988, Langdon and Humphrey 1987), however, many other species of finfish are susceptible

experimentally (Langdon 1989b, Whittington et al. 2010). Experimental bath exposures demonstrated that

the virus can also cause mortality in Macquarie perch (Macquaria australasica), mosquito fish (Gambusia

affinis), silver perch (Bidyanus bidyanus) and mountain galaxias (Galaxias olidus), while Murray cod

(Maccullochella peeli), and Atlantic salmon (Salmo salar) are also susceptible to infection by bath exposure,

but do not become diseased and may act as asymptomatic carriers (Langdon 1989b). Goldfish (Carassius

auratus), European carp (Cyprinus carpio) and Barramundi (Lates calcarifer) are refractory to infection,

while another catadromous species, Australian bass (Macquaria novemaculeata) is susceptible to infection

only by intra peritoneal inoculation, and hence is probably not a natural carrier of the virus (Langdon 1989b).

In Australia, the redfin perch is an introduced species. It is highly susceptible to EHNV, and the virus is

considered endemic in redfin perch populations in many river systems and impoundments in south eastern

Australia, although the distribution of the virus is discontinuous (Whittington et al. 1996, 2010). Clinical

disease in redfin perch is recognized by epizootic mortality, often affecting a very large proportion of the

population (Langdon and Humphrey 1987). Natural epizootics affecting juvenile and adult redfin perch

occur mostly in summer, probably due to higher water temperatures that increase susceptibility of the host to

infection, as Whittington and Reddacliff (1995) found that redfin are susceptible to EHNV infection between

12 and 21°C, but are refractory to infection at water temperatures between 6–10°C. In areas where EHNV is

endemic, only fingerling and juvenile fish tend to be clinically affected, whereas in newly infected

populations adults are also affected (Whittington et al. 2010). A survey to find potential invertebrate hosts

for EHNV including shrimp, freshwater crayfish, Daphnia and water beetles, was unsuccessful (Langdon

1989b). There is little published information available on the susceptibility of obligate marine species to

EHNV. Red sea bream (Pagrus auratus) in Japan (conspecific to the Australian snapper) is not susceptible

to EHNV by intra peritoneal challenge (Nakajima and Maeno 1998), however, closely related viruses in the

family iridoviridae are known to infect the obligate marine turbot (Scophthalmus maximus) in Europe

(Whittington et al. 1996).

_________________________________________________________________________________________________________________________________________________________________________________


80

www.digsfish.com

Clinically diseased fish exhibit necrotic changes in the kidney, liver and spleen, particularly necrosis of

haematopoietic tissues and other parenchymal tissues (Langdon and Humphrey 1987). Transmission of

EHNV between susceptible hosts within a population occurs horizontally via the water probably through the

gills and skin, and also possibly via ingestion of viral particles (Langdon et al. 1988, Whittington and

Reddacliff 1995), however infection by per-os ingestion of tissues from infected fish has not been reported.

Movement of infected trout fingerlings is probably the most common means of spread of EHNV within the

aquaculture industry (Whittington et al. 1996), while translocation of infected redfin perch is the most likely

mechanism of spread in redfin perch (Whittington et al. 1996, 2010).


EHNV continues to be isolated from redfin perch on a regular basis in many river systems and

impoundments in south eastern Australia, as well as in aquacultured rainbow trout (Whittington et al. 1996,

2010). Of the species known to be susceptible to EHNV, redfin perch, rainbow trout and mosquito fish are

known to be used as bait (Tables 2, 8). EHNV is extremely resistant to many physical and chemical

treatments. The virus is resistant to drying (remaining infective after 113 days at 15°C), and showed no

decrease in titre over 3 months in water at 15°C (Langdon 1989b). It can and remain infective in frozen fish

tissues for more than 2 years, though prolonged freezing did affect the onset of cytopathic effect in cell

culture (Langdon 1989b). Clinically diseased fish infected with EHNV can have very high viral titres in

their internal organs, particularly the spleen, liver and kidney (Langdon 1989b), but considering the tissue

tropism of the virus, it is unlikely to be present at high levels in the fillet.

Silver gulls (Larus novaehollandiae) and great cormorants (Phalacrocorax carbo) were observed feeding on

EHNV affected juvenile redfin perch, and the gastrointestinal contents of these birds were positive for

EHNV (Whittington et al. 1996). EHNV is not immediately inactivated by passage through the bird

digestive system and as such, birds are potential mechanical vectors and may spread the virus by

regurgitation of ingested material within a few hours of feeding, as well as mechanically on feathers, feet and

the bill, though it is unlikely that the virus can survive full passage through the gut and remain infective in

the faeces (Whittington et al. 1996). Kewagama Research (2002) estimated that around 1.3% of fishers used

freshwater finfish as bait or berley in 2002, with their use highest in Victoria (2.7% of respondents),

followed by QLD (1.8% of respondents) and NSW/ACT (1% of respondents). However in 2006, national

usage had dropped to 0.7% of respondents, with 3.2% of respondents in the NT using freshwater fish as bait,

as did 1.7% in Victoria and 0.6% in NSW/ACT, with none of the respondents from other states saying they

had used freshwater fish in the previous 12 months (Kewagama Research 2007). This suggests that use of

freshwater finfish as bait is uncommon.

Freshwater finfish are not generally sold as live or frozen bait or berley in retail outlets (except live

ornamental species sold from pet shops), and recreational anglers usually collect them near the fishing site.

However, the majority of salmonid products used for bait or berley will almost certainly originate from

aquaculture farms, as there are few self-sustaining populations of salmonids in Australia. Taking into

account the quantities of freshwater finfish and salmonid products used as bait or berley (Table 1), and also

the fact that clinically diseased fish that carry high titres of EHNV have been recorded at high prevalences at

times in some parts of the country, the likelihood estimations for the occurrence of EHNV in these

commodities are listed below.

_________________________________________________________________________________________________________________________________________________________________________________


81

www.digsfish.com

Release assessment for infection of finfish with EHNV

Commodity

type

Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release

High High Moderate Very low Low Moderate


Freshwater teleosts in south east Australia in water bodies that contain redfin perch are already at risk of

exposure to EHNV, but it is extremely unlikely that freshwater and marine teleosts in other parts of the

country are naturally exposed to this virus (Whittington et al. 2010). Specific data on use of redfin perch and

rainbow trout as bait or berley is not available, however it is known that these species are not generally sold

as bait either fresh or frozen via wholesale outlets such as commercial fishing co-operatives, or at retail

outlets such as fishing tackle shops, service stations and supermarkets. However, fresh or frozen rainbow

trout are widely sold as food fish throughout Australia, and these could be diverted to use as bait or berley in

both freshwater and marine environments around the country. Because EHNV is highly resistant to freezing

for many months, a large percentage of any virus present in carrier fish is likely to remain viable after

thawing. The scale of use of redfin perch and rainbow trout (as well as other potential carriers such as

mosquito fish) as live bait in Australia remains unknown, but since they are not available commercially, the

quantities translocated are likely to be relatively small. This suggests that direct pathways exist for

translocated infected products used by private fishers to enter both freshwater and marine environments, thus

potentially exposing wild fish to viable EHNV.

Natural EHNV infections have only been reported from redfin perch and rainbow trout, however the full

extent of the range of fish species susceptible to EHNV has not been determined. The virus can infect and

cause mortality in several species of native freshwater fishes, including macquarie perch (Macquaria

australasica), silver perch (Bidyanus bidyanus) and mountain galaxias (Galaxias olidus). Mosquito fish

(Gambusia affinis) are also susceptible, while Murray cod (Maccullochella peelii) and Atlantic salmon

(Salmo salar) can act as carriers (Langdon 1989b). However, infection would occur only if sufficient

quantities of virus (i.e. an infective dose) were introduced into an area where susceptible hosts were present.

Redfin perch are extremely susceptible to EHNV with immersion in as few as 0.08 TCID50/ml being a lethal

dose for 100% of fish (Whittington and Reddacliffe 1995). Macquarie perch were also highly sensitive to

infection, with 100% mortality being recorded in fish exposed to 103 TCID50/mL by immersion (Langdon

1989b). It appears that mosquito fish are also very susceptible to infection, as Langdon (1989b) recorded

90% mortality in mosquito fish exposed by immersion to 103 TCID50/mL EHNV, and 100% mortality in

mosquito fish exposed to EHNV infected redfin perch by co-habitation. Whittington and Reddacliffe (1995)

found that rainbow trout in their experiments were not susceptible to infection solely by immersion, but

Langdon (1988) successfully infected one out of four 3.5-4 cm long rainbow trout via bath inoculation with

103 TCID50/mL at 22°C.

_________________________________________________________________________________________________________________________________________________________________________________


82

www.digsfish.com

Clinically diseased redfin perch infected with EHNV can have viral titres up to 108.5 TCID50/g in the spleen,

and 105 TCID50/g in the liver and head kidney (Langdon 1989b). Given that redfin perch can be infected by

immersion at extremely low infective doses (Whittington and Reddacliffe 1995), and several other fish

species can also become infected at relatively low doses compared to the titres present in clinically diseased

fish, the likelihood that an infectious dose can be transmitted horizontally into a new water body via use of

clinically diseased bait or berley remains high, even if only small quantities of bait or berley are translocated,

and possibly even if subclinically infected fish are used. Given that pathways exist for translocation and

spread of EHNV into the environment via use of bait, and acknowledging that the virus can infect a range of

susceptible hosts, the risk of exposure and establishment is non-negligible, and the likelihood of exposure

and establishment of EHNV in new fish populations is considered to be Moderate.


When fish become infected with EHNV in a new area, mortality can occur in all size classes of fish, and

because of this the virus has the potential to cause significant damage to fish populations (Langdon 1989b,

Whittington 2010). The virus is also known to be pathogenic to some species of threatened native fishes,

and while the full extent of its host range remains unknown, the fact that mosquito fish are susceptible means

that vectors are readily available in many freshwater environments around Australia where EHNV is

currently absent. EHNV is already present in some parts of the Australian environment, where it is known

to cause annual disease outbreaks in wild populations of susceptible fish species. The virus is listed by the

OIE and NACA and is a reportable disease in all States except the ACT (Table 3b). Hence spread of the

virus into new geographic areas is likely to adversely affect trade. Considering all of these factors,

establishment of the disease would have significant or serious biological consequences, which would not be

amenable to control, and could cause irreversible environmental effects as well as economic damage to

fisheries and aquaculture industries. It is therefore estimated that the consequences of introduction of EHNV

into different parts of the Australian environment via use of infected bait or berley would likely be High.


The unrestricted risk associated with EHNV is determined by combining the likelihood of entry and

exposure (from Table 5) with the consequences of establishment (Table 7). The unrestricted risk estimate for

EHNV exceeds the ALOP for all commodity types, suggesting that additional risk management is required

for this disease agent.

_________________________________________________________________________________________________________________________________________________________________________________


83

www.digsfish.com

Risk estimate for infection of finfish with EHNV

Commodity type Live

finfish

Whole

fresh dead

finfish

Frozen

whole

finfish

Frozen

fillets

Frozen

fish heads

Frozen

guts/offal

Combined

likelihood of release

and exposure

Moderate Moderate Low Very Low Low Low

Consequences of

establishment

High High High High High High

Risk estimation High Risk High Risk Moderate

Risk

Low Risk Moderate

Risk

Moderate

Risk

_________________________________________________________________________________________________________________________________________________________________________________


84

www.digsfish.com

5.3 Infection of finfish with Viral Encephalopathy and Retinopathy (Nodavirus)

5.3.1 Aetiologic agent: Non-enveloped viruses with a genome containing two segments of positive sense

single stranded RNA of the genus Betanodavirus in the Family Nodaviridae.

5.3.2 OIE List: No NACA List: Yes

5.3.3 Australias status: Betanodavirus infections have been reported in QLD, NSW, Tasmania, SA, WA

and the NT (Moody and Horwood 2008). These viruses are reportable diseases in all states excluding QLD

(Table 3b).

5.3.4 Epidemiology

Betanodaviruses are the causative agents for viral encephalopathy and retinopathy (VER), a serious disease

of larval and juvenile marine fish that occurs almost worldwide (OIE 2010b). Clinical signs of the disease

are most commonly observed in larvae and fry and are due to damage to the nervous tissue of the spinal cord,

brain and retina caused by heavy vacuolation and necrosis of the tissues of the central nervous system

(Munday et al. 1992, 2002, Munday and Nakai 1997). Affected fish display abnormal behaviour, including

spiral swimming and rapid uncoordinated darting movement, with mass mortalities occurring over a short

period of time (Munday et al. 1992). Colour changes (lightening or darkening of diseased fish), cessation of

feeding and increased susceptibility to cannibalism may also be observed (Moody and Horwood 2008). The

onset of mortalities due to betanodavirus infection are usually observed between 1 and 40 days post hatching

in larval fish, with mortalities commonly approaching 100%, though susceptibility decreases as the age of

the fish increases (Moody and Horwood 2008). Fish that survive epizootics as juveniles can harbour sub-

clinical infections as adults, and the virus appears to be transmitted vertically to their progeny via eggs or

sexual fluids (Munday et al. 2002). More than 37 species of fish from all continents (excluding Africa) are

known to be susceptible to nodaviruses (Moody and Horwood 2008) and this number is bound to rise as

more species are cultured. Some species such as the gilt head sea bream Sparus aurata, become infected, but

do not exhibit clinical signs of this disease, and hence can act as asymptomatic carriers (Castric et al. 2001).

In Australia, disease caused by nodavirus has been reported from Australian bass (Macquaria

novemaculeata), barramundi (Lates calcarifer), barramundi cod (Cromileptes altivelis), goldspotted rockcod

(Epinephelus coioides), flowery cod (Epinephelus fuscoguttatus) and striped trumpeter (Latris lineata) from

marine aquaculture facilities in NSW, NT, QLD, SA and Tasmania, and from sleepy cod (Oxyeleotris

lineolatus) from a freshwater aquaculture facility in QLD (Moody and Horwood 2008). Other species from

which nodavirus has been suspected (without disease) include eel tailed catfish (Tandanus tandanus)

(Munday et al. 2002), while juvenile cultured kingfish from SA were weakly positive for Australian bass

nodavirus nucleic acid using nested PCR, but no virus was cultured in cell culture (unpublished laboratory

reports provided to BK Diggles from AAHL). Silver trevally (Pseudocaranx dentex) are highly susceptible

to nodavirus infection, however no studies appear to have been conducted to determine whether nodavirus

occurs naturally in wild P. dentex in Australia. It therefore appears highly likely that many other species of

finfish in Australia are susceptible to nodavirus infection.

_________________________________________________________________________________________________________________________________________________________________________________


85

www.digsfish.com


Nodavirus continues to be isolated from cultured marine fish in Australia on a regular basis. Nodavirus

infections have been reported from finfish in all states except Victoria and the ACT (Moody and Horwood

2008). While no phylogenetic information is available for WA isolates, all the other studied Australian

isolates have been identified as members of the RGNNV genotype (Moody and Horwood 2008). However,

the isolates form two distinct strains within this genotype; one group of isolates from NSW and SA and a

second group of isolates from QLD, the NT and Tasmania (Moody and Horwood 2008). As the effect of the

different strains on native finfish species is unknown, controls are in place to minimise risk of escape of virus

from aquaculture facilities or via translocation of infected stock in restocking programs. Exclusion of the

virus from aquaculture premises, good hygiene and reduced stocking densities has contributed to decreasing

the incidence of nodavirus outbreaks in recent years (Moody and Horwood 2008). It is likely that wild fish

are the main reservoir of infection, and hence use of nodavirus RT-PCR -negative broodstock has reduced

the occurrence of disease in cultured larvae (Moody and Horwood 2008).

The highest titres of nodavirus in both clinically diseased and carrier fish occur in the tissues of the brain,

and retina, where the majority of active viral replication occurs, however the virus has also been detected in

other internal organs (Korsnes et al. 2009). Nodavirus is persistent in the environment, remaining infective

after 6 months in seawater and 3 months in freshwater at 15°C, but exhibiting complete loss of viability after

6 months in freshwater (Frerichs et al. 2000). These viruses are also reasonably resistant to a broad range of

disinfectants (Frerichs et al. 2000, Bovo et al. 2005). Nodavirus is very tolerant of freezing, with Frerichs et

al. (2000) finding no loss of infectivity after freezing for 1 year at -20°C.

Nodavirus is likely to occur naturally in many species of marine fish throughout Australian waters. Families

of fishes used as bait in Australia that are susceptible to infection by nodavirus include carangids, elotrids

and teraponids (Table 8). Taking into account the large quantities of marine finfish used as bait or berley

(Table 1), and the prevalence of the disease agent, the likelihood estimations for the occurrence of aquatic

birnavirus in these commodities are listed below.

Release assessment for infection of finfish with Nodavirus

Commodity

type

Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release

High High High Low High Moderate


Marine teleosts in most parts of Australia are already at risk of exposure to local strains of nodavirus.

However it is not known whether significant differences in pathogenicity or host specificity exist between

_________________________________________________________________________________________________________________________________________________________________________________


86

www.digsfish.com

the different nodavirus strains that are present in different parts of the country (Moody and Horwood 2008),

or even whether teleosts in freshwater environments are naturally exposed to these disease agents. While

nodavirus infections were originally considered a disease of marine fishes only, recently they have caused

serious disease in the culture of freshwater fishes such as tilapia (Bigarr et al. 2009). Throughout Australia,

fresh or frozen whole or processed marine fish are offered for sale as bait via wholesale outlets such as

commercial fishing co-operatives, or packaged for resale at retail outlets such as fishing tackle shops, service

stations and supermarkets. Species known to be particularly susceptible to nodavirus, such as cultured

barramundi, are widely sold as fresh or frozen food fish throughout Australia, and these could be diverted to

use as bait or berley in both freshwater and marine environments around the country. In Australia, finfish for

use as live bait are not generally available for retail sale (except ornamental species sold from pet shops). So

the quantity of baitfish translocated live is likely to be small, although recreational anglers may still catch

their own live bait at the fishing location. The majority of the volume of bait translocated throughout the

country is frozen, including frozen whole fish and processed fish products (e.g. heads or guts). Because

nodavirus is stable when frozen (Frerichs et al. 2000), a large percentage of virus is likely to remain viable

after thawing. Direct pathways therefore exist for translocated bait or berley from these retail outlets to enter

both freshwater and marine environments, potentially exposing wild fish to viable nodavirus. Other

pathways for exposure of wild fish to viable nodavirus include disposal into natural waters of untreated

processing wastes from commercial processing premises, and use of processing wastes in berley blocks.

Nodavirus infections have been reported from a wide range of fish species, including many species groups

that occur in Australian waters. These species are therefore likely to be susceptible to infection by nodavirus

carried in infected bait, but infection would occur only if sufficient quantities of virus (i.e. an infective dose)

were introduced into an area where susceptible hosts were present. Susceptible fish can become infected

with nodavirus via horizontal transmission through the water with entry via the skin, intestine or nasal cavity,

as well as by vertical transmission (Munday and Nakai 1997). The infectious dose of nodavirus by the

immersion pathway will probably vary depending on the strain of virus used and the species challenged, but

the literature shows the disease is consistently transferred at waterborne concentrations of 103 TCID50/mL, a

level which caused 100% mortality in larval and juvenile barramundi within 4 days (Parameswaran et al.

2008), and up to 75% mortality in 0.3 – 1.0 g wolfish (Anarhichas minor) larvae (Sommer et al. 2004). It is

also notable that concentrations of 1.6 x 104 TCID50/ml were recorded by Nerland et al. (2007) in rearing

water where epizootics had been recorded in halibut (Hippoglossus hippoglossus) larvae.

Clinically diseased fish infected with nodavirus can have high viral titres in internal organs (>108 TCID50/g,

see Bigarr et al. 2009), and while only larvae and early juveniles usually become clinically diseased, juvenile

fish are commonly used as live or frozen bait (B.K. Diggles, personal observations). The levels of nodavirus

in sub-clinically infected fish can also be relatively high compared to the infectious dose (103 - 105 TCID50/g,

see Castric et al. 2001) despite the absence of disease. Because susceptible fish can be infected by

immersion at around 103 TCID50/mL, it is possible that an infectious dose could be transmitted horizontally

into the water in the vicinity of a bait that was clinically, or even sub-clinically infected with nodavirus. Fish

could also theoretically be exposed to an infectious dose via the per-os route if they consumed enough bait

fish containing virus levels typical of clinically or sub-clinically infected fish. Given that pathways exist for

translocation and spread of nodavirus into the environment via use of bait or berley, and acknowledging the

broad range of susceptible hosts, the risk of exposure and establishment is non-negligible, and the likelihood

of exposure and establishment of nodavirus in new fish populations is considered to be Moderate.

_________________________________________________________________________________________________________________________________________________________________________________


87

www.digsfish.com


When fish become infected with nodavirus, mortality is mainly restricted to larval and early juvenile stages,

and disease does not necessarily occur in larger fish. However, infection of larger fish can still occur without

disease. Infected fish can remain carriers for life, and can transfer the virus to their progeny vertically via

infected gametes. These viruses are unlikely to cause disease in adult fish in the wild, hence the main

consequences appear those related to possible increased mortality of larval and early juvenile stages (which

could have impacts on wild fisheries at the population level), as well as increased costs of production in

finfish aquaculture hatcheries due to use of infected wild caught broodstock. Because at least two strains of

nodavirus are already present in the Australian environment, the full extent of any increases in mortality rates

that may occur due to translocation of the disease agents via bait is difficult to assess. These viruses are no

longer listed by the OIE, but remain listed by NACA and on State reportable disease lists, hence their spread

can still have adverse impacts on trade. Considering all of these factors, establishment of the disease would

most likely have mild to moderate biological consequences, which may be amenable to control, and are not

likely to cause irreversible environmental effects. It is therefore estimated that the consequences of

introduction of nodavirus strains into different parts of the Australian environment via use of infected bait or

berley would likely be Low.


The unrestricted risk associated with nodavirus is determined by combining the likelihood of entry and


nodavirus exceeds the ALOP for live, whole fresh dead, and whole frozen finfish, and frozen fish heads,

suggesting that additional risk management is required for this disease agent in these commodities.

Risk estimate for infection of finfish with Nodavirus

Commodity type Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen

fish heads

Frozen

guts/offal

Combined


and exposure

Moderate Moderate Moderate Low Moderate Low

Consequences of

establishment

Low Low Low Low Low Low

Risk estimation Low risk Low risk Low risk Very low

risk

Low risk Very low

risk

_________________________________________________________________________________________________________________________________________________________________________________


88

www.digsfish.com

5.4 Emergence of a new previously unknown virus of finfish

5.4.1 Aetiologic agent: A previously undescribed virus that infects finfish.


5.4.3 Australias status: Unknown distribution and not listed as a reportable disease in any State (Table 3b).

5.4.4 Epidemiology

Despite an increasing amount of research on aquatic animal health in recent times, and the existence of a

broad base of knowledge accumulated over many years (e.g. see review by Humphrey 1995a, 1995b), there

is still a significant lack of knowledge and understanding of the full range of parasites and disease agents of

aquatic animals in Australia. For example, Dove and O’Donoghue (2005) studied the trichodinid ciliate

ectoparasites of introduced and native freshwater fishes. They found that 81% (17) of the 21 species of

Trichodina found in 33 species of fish examined were undescribed, and used a simple formula to estimate

that the biodiversity of these parasites in Australian freshwater fish may approach 150 species, a number that

approached the total number of trichodinid species then described from all freshwater hosts at that time

(Dove and O’Donoghue 2005). Similarly, Bott et al. (2005) studied the digenean fauna of bivalves on the

Great Barrier Reef, and compared their findings to the number of known digenean species recorded at that

time from final hosts (fish and birds) that occur in the region. They deduced that, at best, only around 10%

of the digenean fauna from the Great Barrier Reef was known (Bott et al. 2005). These examples are

probably typical, and highlight the incompleteness of our knowledge of the parasites and disease agents of

Australias aquatic fauna. Because of this, it is virtually certain that disease risks are present during

translocation of bait products, even in the absence of identification of known pathogens in the commodity

(Gaughan 2002).

Gaughan (2002) suggests that a RA should embrace case histories linked to unidentified pathogens. To

accommodate this view, this RA will now examine the translocation of a hypothetical, previously

undescribed, virus in finfish to examine whether risks posed by unidentified pathogens can be adequately

assessed by the RA process. For the sake of this exercise, the hypothetical virus will be carried by redbait

(Emmelichthys nitidus), a species which is captured in an expanding midwater trawl fishery off Tasmania at

depths of 100 to 500 meters (Neira et al. 2009), product from which is used in mainland Australia as feed for

southern bluefin tuna, and as bait in lobster fisheries in SA and WA.


There is very little known about the parasites and nothing known of the virology of the Family

Emmelichthyidae (bonnetmouths). Redbait resemble clupeids or jack mackerel in both appearance and

ecological niche (Welsford and Lyle 2003), and the parasite fauna of E. nitidus (a monogenean, several

digeneans, cestodes and anisakid nematodes, see Korotaeva (1975)) suggests that these fish are plankton

feeders. Indeed, redbait consume mostly zooplankton, primarily crustaceans, and form an important part of

the diet of large predatory fish, seals and seabirds and are likely to be a key species in the continental shelf

_________________________________________________________________________________________________________________________________________________________________________________


89

www.digsfish.com

pelagic ecosystem (Welsford and Lyle 2003). However, this tells us very little about whether E. nitidus

harbours any disease agents of concern that should be considered in a RA, particularly microparasites such as

viruses, bacteria or protozoa. Large differences in the size at first maturity in different locations near

Tasmania (Ewing and Lyle 2009) suggest that the redbait population may be structured, a finding that may

have important ramifications in relation to the extent of the disease risks posed by translocation of redbait.

For the sake of this exercise, it is assumed that redbait harbour a pathogenic iridovirus that affects primarily

larvae and juveniles. It will also be assumed that the prevalence in the population is only moderate, but

many adult fish are sub-clinical carriers of the disease. Like other iridoviruses, the new virus mainly infects

internal organs and is stable in frozen tissue. Because large quantities of redbait could be used as bait in rock

lobster fisheries, the likelihood estimations for the occurrence of the hypothetical new virus in these

commodities would be as follows.

Release assessment for a new previously unknown virus of finfish

Commodity

type

Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release

High High High Low Low High


If redbait populations have significant population structuring (Ewing and Lyle 2009), it is very possible that

redbait and other teleosts in mainland Australia have either not been exposed to the new virus, or not been

exposed to Tasmanian strains of the new virus. If teleosts on the mainland have not been previously exposed

to the new virus, these naïve populations may have a lower infective dose and/or may be more likely to be

susceptible to disease compared to fishes from regions where the virus is endemic. Use of infected redbait as

lobster bait as opposed to aquaculture feed will reduce the chances of disease transmission, because in the

rock lobster fishery the bait is spread very sparsely over a very large area (2.5-5.4 kg of bait per day per km2

of ocean floor, see Jones and Gibson 1997). However, fish potentially susceptible to the new virus can enter

lobster pots and eat the infected redbait, and/or remain in or near pots in close vicinity of the infected redbait,

meaning that the chances of establishment of an index case of infection are not negligible. The chances of

establishment of the index case will depend on the volumes of bait translocated, as well as the way the bait is

processed and handled, and how used bait is disposed of at sea. The new virus, like most other iridoviruses,

is stable when frozen, and hence a large percentage of virus is likely to remain viable after thawing. This

suggests that a direct pathway exists for viruses from translocated redbait to enter the marine environment,

thus potentially exposing susceptible wild fish to viable iridovirus. Other significant pathways for exposure

of marine (and even freshwater) fish to the new iridovirus would include disposal into natural waters of

untreated processing wastes from commercial processing premises, and use of processing wastes in berley

blocks.

_________________________________________________________________________________________________________________________________________________________________________________


90

www.digsfish.com

Because the virus is new, the range of species likely to be susceptible to infection would be unknown.

However, it would be safe to assume that redbait in mainland Australian waters would be susceptible, and

that that they may not have been previously exposed to Tasmanian strains of the virus. It may also be

possible for fishes with similar ecological niches in the new location, e.g. pilchards (Sardinops

neopilchardus) to also be susceptible to the new virus. However, infection would occur only if sufficient

quantities of virus (i.e. an infective dose) were introduced into an area where susceptible hosts were present.

Susceptible fish could become infected with the new virus via horizontal transmission through the water, but

the likelihood of this happening is not possible to determine without an understanding of the infectious dose

required, and the levels of virus present in clinically diseased and subclinically infected redbait. At this stage

the RA would be significantly compromised by a lack of information, unless data from other iridovirus

agents was substituted. If this was the case, and the hypothetical virus produced infections with viral

burdens and infective dose characteristics similar to other virulent iridoviruses (such as EHNV), given that

pathways exists for translocation and spread of the new virus into the environment inhabited by susceptible

hosts via use of bait, the risk of exposure and establishment would be non-negligible, though it would not be

possible to accurately estimate the likelihood of exposure and establishment. However, for the sake of this

exercise, given the end use as lobster bait, and based on information on transmission of other iridoviruses so

as to obtain a measure of the combined likelihood of release and exposure (Table 5), it will be assumed that

the likelihood of exposure and establishment is considered to be Low.


The consequences of establishment of a new virus, and whether additional risk management may be

required, would depend heavily on the range of susceptible hosts, the pathogenicity of the virus to

susceptible hosts, the importance of fisheries that may exist for susceptible hosts, the ecological position of

susceptible hosts in marine ecosystems, and the conservation status of susceptible hosts. It is also not

possible to determine whether the virus would cause disease to such an extent that it would be listed as a

notifiable disease, and thus potentially have adverse effects on trade. All of this information cannot be

estimated without first knowing the virus exists and which hosts are susceptible. Hence it is the consequence

assessment stage of the RA that is most severely handicapped by the absence of identification of known

pathogens in the commodity.


The unrestricted risk associated with the hypothetical unknown virus would be determined by combining the

likelihood of entry and exposure (from Table 5) with the consequences of establishment (Table 7).

However, because the unrestricted risk estimate relies heavily on the assessment of the consequences of

establishment to determine whether additional risk management is required, it will depend heavily on the

identity of the susceptible hosts and the pathogenicity of the virus to those hosts. Unfortunately, this

information cannot be estimated without first knowing the virus exists, and which hosts are susceptible. This

scenario demonstrates that a scientific risk analysis for a hypothetical unknown virus is not possible,

however this process of undertaking a RA on an unknown new virus is valuable because it highlights the

type of information required to make such an assessment possible. In this case, the information needed

would be obtained through surveillance in the form of screening of statistically defined numbers of redbait

for viruses by cell culture and/or molecular techniques in order to detect, isolate and study unknown viruses.

_________________________________________________________________________________________________________________________________________________________________________________


91

www.digsfish.com

If a virus was isolated, data on the levels of virus present in subclinically infected and/or clinically diseased

redbait could be obtained, and further experiments may be able to determine the viruses identity, infectious

dose and pathogenicity to key fish species from the region receiving the translocated bait. All of these data

could then be used to inform a RA process to help determine the risk posed by translocation of the new virus

with the commodity.

So this hypothetical scenario highlights the importance of active, rather than passive, disease surveillance, an

activity that could be implemented in a structured manner either in the early stages of development of

fisheries for new species likely to be used as bait or berley, or whenever significant quantities of bait or

berley are being translocated to a new geographical area.

Risk estimate for a new previously unknown virus of finfish

Commodity type Live

finfish

Whole

fresh dead

finfish

Frozen

whole

finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Combined


and exposure

Low Low Low Very Low Very Low Low

Consequences of

establishment

Unknown Unknown Unknown Unknown Unknown Unknown

Risk estimation Not

possible

Not

possible

Not

possible

Not

possible

Not

possible

Not

possible

_________________________________________________________________________________________________________________________________________________________________________________


92

www.digsfish.com

5.5 Infection of finfish with Aeromonas salmonicida (Goldfish ulcer disease, GUD)

5.5.1 Aetiologic agent: Atypical strains of Aeromonas salmonicida, which are gram negative bacteria in the

Family Aeromonadaceae.


5.5.3 Australias status: Aeromonas salmonicida (atypical variants) have been reported in QLD, NSW,

Victoria, SA and the ACT. They are reportable disease agents in all states excluding the ACT (Table 3b).

5.5.4 Epidemiology

“Atypical” variants of the Aeromonas salmonicida bacterium affect mainly non-salmonids and differ in

several characteristics from Aeromonas salmonicida ssp. salmonicida, a bacterium exotic to Australia that

causes furunculosis in salmonids. Infection with atypical A. salmonicida causes ulcerative dermatitis, but

does not necessarily result in the acute mortality and septicaemia characteristic of typical furunculosis,

manifesting instead mainly as external lesions and ulceration (Wiklund and Dalsgaard 1998). Three

biovariants of atypical Aeromonas salmonicida have been isolated in Australia, firstly A. salmonicida var.

Nova which causes goldfish ulcer disease (GUD) in goldfish (Carassius auratus) and silver perch (Bidyanus

bidyanus) (Trust et al. 1980, Humphrey and Ashburner 1993), a second variant from greenback flounder

(Rhombosolea tapirina) which causes ulcer disease of flounder (Whittington et al. 1995), and a third (A.

salmonicida var. Acheron) from Atlantic salmon (Salmo salar) that causes marine aeromonad disease of

salmonids (MAS) (Carson and Gudkovs 2001). Goldfish ulcer disease was introduced into Australia in 1974

with goldfish broodstock imported from Japan into a commercial goldfish farm in Victoria (Trust et al. 1980,

Humphrey and Ashburner 1993). From there, the bacterium was translocated within Victoria and into 4

other States by 1977 through sales of goldfish via the aquarium industry (Trust et al. 1980, Humphrey and

Ashburner 1993, Whittington et al. 1987, 1995). Spread of GUD into the wild was facilitated by release of

effluent water from affected goldfish farms, release of live infected goldfish into waterways and use of

goldfish as bait (Whittington and Cullis 1988, Humphrey and Ashburner 1993). Outbreaks of GUD in

Australia have been reported in most parts of the Murray-Darling River system, where the spread of the

bacterium has been facilitated by movements of feral populations of goldfish and carp (Cyprinus carpio), in

which the bacterium is enzootic (Humphrey and Ashburner 1993, Whittington et al. 1995).

In 1993 the second atypical A. salmonicida variant was recovered from ulcerative dermal lesions and kidney

of greenback flounder cultured in seawater in Tasmania (Whittington et al. 1995, Carson and Gudkovs

2001). Atlantic salmon (Salmo salar) and striped trumpeter (Latris lineata) have been infected by co-

habitation with the flounder variant (Herfort 2004). The bacteria isolated from the greenback flounder

differs genetically from isolates from goldfish and silver perch, and probably originated from the marine

environment (Whittington et al. 1995). Atlantic salmon cultured in seacages in Tasmania have also been

infected by the third variant (A. salmonicida var. Acheron), causing MAS (Carson and Gudkovs 2001),

which has a clinical presentation similar in some respects to furunculosis (Herfort 2004). Other species

susceptible to infection with atypical A. salmonicida that occur in Australia include redfin perch (Perca

fluviatilis), roach (Rutilus rutilus) and silver biddy (Gerres ovatus) (Herfort 2004).

_________________________________________________________________________________________________________________________________________________________________________________


93

www.digsfish.com


Atypical variants of A. salmonicida occur in many freshwater rivers and lakes in south eastern Australia and

the Murray Darling system, as well as marine waters of Tasmania, where the Acheron biovariant is currently

restricted to a relatively small salmon production area. There appears to be no published data to indicate

whether these bacteria occur in the marine waters of mainland Australia, however there are no barriers to

migration of marine fish from Tasmania to mainland Australia, and some species do migrate between these

two areas (Stanley 1978). The Nova biovariant is very pathogenic for salmonids, but its effects on native

finfish species are mostly unknown. This has lead to strict controls on movements of goldfish to reduce the

risk of translocation of the Nova biovariant from south east Australia into Tasmania and WA.

Infected fish may carry A. salmonicida covertly in sub-clinical infections at very low intensities that are

extremely hard to detect, even with PCR (Carson and Gudkovs 2001). However, carriers of A. salmonicida

usually revert to a diseased state during stressful events, and as such "stress tests" have been devised to

improve the chances of detecting carrier fish (Wiklund and Dalsgaard 1998). The highest numbers of

bacteria in clinically diseased fish occur in the skin gills and muscle tissue affected by ulcers, though the

bacterium can also be isolated in other internal organs such as the kidney (Whittington and Cullis 1988,

Wiklund and Dalsgaard 1998). Though an obligate fish pathogen, the bacterium may be able to persist in the

environment for long periods in water and sediments, from 60 days (Wiklund and Dalsgaard 1998) to 9

months (Hine and McDiamant 1997). Silver perch became infected at an aquaculture farm where clinical

GUD was recorded 7 years previously, however clinically normal goldfish and carp present in other parts of

the farm may have acted as carriers during the interim period (Whittington et al. 1995). These bacteria are

susceptible to a broad range of antibiotics and disinfectants (Hine and McDiamant 1997), though resistance

to some antibiotics has developed. Vaccines against A. salmonicida have been successfully developed,

including against the Acheron biovariant. These bacteria are very tolerant of freezing, though there may be

some loss of viability upon thawing (Hine and MacDiarmid 1997).

Atypical variants of A. salmonicida are known to occur naturally in a limited range of fish species in certain

regions of Australia. Families of fishes used as bait or berley in Australia that are susceptible to infection by

A. salmonicida include goldfish, carp, redfin perch, and salmonids (Table 8). Taking into account the

relatively small quantities of these that are used as bait or berley (Table 1), and the prevalence of the disease

agent, the likelihood estimations for the occurrence of atypical variants of A. salmonicida in these


Release assessment for infection of finfish with Aeromonas salmonicida (atypical variants)

Commodity

type

Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release

High High Moderate Moderate Moderate Moderate

_________________________________________________________________________________________________________________________________________________________________________________


94

www.digsfish.com


Freshwater teleosts in the south east regions of Australia are already at risk of exposure to endemic variants

of A. salmonicida carried by feral populations of goldfish and carp. Several species of marine fish are also

exposed to variants of A. salmonicida in the Tasmanian marine environment. Throughout Australia, fresh or

frozen whole or processed marine fish are offered for sale as bait or berley via wholesale outlets such as

commercial fishing co-operatives, or packaged for resale at retail outlets such as fishing tackle shops, service

stations and supermarkets. However none of the obligate marine fish species known to be infected by

atypical A. salmonicida variants in Tasmania are used as bait or berley in any significant quantity. Similarly,

the quantities of translocated fish sold as live bait is likely to be negligible, however recreational fishers may

still catch their own live bait at the fishing location, and a small proportion of anglers may still persist in

unlawfully translocating live goldfish, redfin or carp as bait or berley in freshwater regions (Whittington and

Cullis 1988, Humphrey and Ashburner 1993), while silver biddy are a popular live bait in east coast estuaries

(B.K. Diggles, personal observations). Cultured salmonids and their processing wastes are not sold for bait

or berley (Grant Pullen, DPIWE Tasmania, personal communication 8 April 2011), but fresh or frozen

Atlantic salmon are widely distributed as food fish and hence some product could be diverted to use as bait

or berley around the country. Because atypical A. salmonicida is stable when frozen, a large percentage of

virus is likely to remain viable after thawing, this suggests that direct pathways exist for fresh or frozen

salmon products used as bait or berley to enter both freshwater and marine environments, thus potentially

exposing wild fish to viable atypical A. salmonicida. Other significant pathways for exposure of marine and

freshwater fish to atypical A. salmonicida would include disposal into natural waters of untreated processing

wastes from commercial processing premises.

Many fish species in regions where this bacterium is not enzootic are likely to be susceptible to infection by

A. salmonicida carried in infected bait or berley, but infection would occur only if sufficient quantities of

bacteria (i.e. an infective dose) were introduced into an area where susceptible hosts were present.

Whittington and Cullis (1988) used the Nova biovariant from goldfish to find the 10 day LD50 for Atlantic

salmon via IP injection was 7.4 x 10-3 cfu, 3 x 10-2 cfu for brown trout, 3.7 x 102 cfu for brook trout and 6.4 x

103 cfu for rainbow trout. The infective dose for brown and rainbow trout by immersion was 105 - 106

cfu/ml., but this route was not successful for infecting Atlantic salmon in their experiments, although they

reported horizontal transfer of A. salmonicida by co-habitation to 5/195 control fish placed in the same tanks

as infected fish (Whittington and Cullis 1988). In another study done around the same time, Carson and

Handlinger (1988) found the LD50 of 30-40 gram Atlantic salmon by IP injection of A. salmonicida from

goldfish was 3 cfu, while a bath of 8 x 105 cfu/ml was lethal to 80-90% of salmon (Carson and Handlinger

1988). These data confirm that some salmonids are extremely susceptible to A. salmonicida infection from

goldfish.

Clinically diseased fish infected with atypical A. salmonicida can have large numbers of bacteria in skin and

muscle lesions and internal organs. In contrast, infected fish may carry A. salmonicida covertly in sub-

clinical infections at very low intensities that are extremely hard to detect (Carson and Gudkovs 2001). The

minimum dose required to infect local fish species with A. salmonicida by immersion have not been

published, however the results of Whittington and Cullis (1988) suggest the disease can be transferred

horizontally by cohabitation of susceptible hosts with infected fish. It appears possible, therefore, that an

infectious dose could be transmitted horizontally via the water in the immediate vicinity of bait or berley that

_________________________________________________________________________________________________________________________________________________________________________________


95

www.digsfish.com

was clinically infected with atypical A. salmonicida, but this would probably not occur if the bait or berley

was sub-clinically infected. Fish could also theoretically be exposed to an infectious dose via the per-os

route if they consumed a bait fish containing virus levels typical of clinically infected fish. Clinically

infected fish are likely to be aquacultured only, and some states have bans on use of aquacultured products as

bait (T. Hawkesford, personal communication). However, given that pathways still exist for translocation

and spread of atypical A. salmonicida into the environment via use of bait and berley, and acknowledging

that very susceptible hosts do exist, but they may require exposure to reasonably high concentrations of the

bacterium before they become infected by the immersion route, the risk of exposure and establishment is

non-negligible, and the likelihood of exposure and establishment of atypical A. salmonicida in new fish

populations is considered to be Moderate.


When naïve susceptible fish become infected with atypical A. salmonicida, disease and mortality can occur

in fish of all age classes. Infected fish that survive the infection can remain carriers for life. Disease caused

by atypical A. salmonicida is usually not a severe problem in wild fish and establishment is unlikely to have

impacts on wild fisheries at the population level. The main consequences appear those related to possible

increased mortality as well as increased costs of production in finfish aquaculture, where use of antibiotics or

production of effective vaccines would be required to prevent significant losses. Because three variants of

the bacterium are already present in the Australian environment, the full extent of any increases in mortality

rates that may occur due to translocation of the disease agents via bait or berley is difficult to assess. These

bacteria are not listed by the OIE, or NACA, but remain on State reportable disease lists, hence their spread

can still have adverse impacts on trade. Considering all of these factors, establishment of the disease would

most likely have mild to moderate biological consequences for aquaculture, which may be amenable to

control, and are not likely to cause irreversible environmental effects. It is therefore estimated that the

consequences of introduction of atypical A. salmonicida into different parts of the Australian environment

via use of infected bait or berley would likely be Low.


The unrestricted risk associated with atypical variants of A. salmonicida is determined by combining the

likelihood of entry and exposure (from Table 5) with the consequences of establishment (Table 7). The

unrestricted risk estimate for atypical variants of A. salmonicida exceeds the ALOP for live finfish and whole

fresh dead finfish, suggesting that additional risk management is required for this disease agent in these

commodities.

_________________________________________________________________________________________________________________________________________________________________________________


96

www.digsfish.com

Risk estimate for infection of finfish with Aeromonas salmonicida (atypical variants)

Commodity type Live

finfish

Whole

fresh

dead

finfish

Frozen

whole

finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Combined


and exposure

Moderate Moderate Low Low Low Low

Consequences of

establishment


Risk estimation Low Risk Low Risk Very Low

Risk

Very Low

Risk

Very Low

Risk

Very Low

Risk

_________________________________________________________________________________________________________________________________________________________________________________


97

www.digsfish.com

5.6 Infection of salmonids with Lactococcus garvieae

5.6.1 Aetiologic agent: Lactococcus garvieae, a gram positive bacterium in the Family Streptococcaceae.


5.6.3 Australias status:. Reported in Tasmania and Victoria, and is a reportable in Tasmania (Table 3b).

5.6.4 Epidemiology

Lactococcus garvieae is an opportunistic pathogen of marine and freshwater fish worldwide (Eldar et al.

1999, Vendrell et al. 2006). This bacterium has a wide host range which includes both aquatic and terrestrial

vertebrates (including cattle and humans), and aquatic invertebrates (Eldar et al. 1996, Eldar et al. 1999,

Vendrell et al. 2006). It is a facultatively anaerobic, non-motile, non-spore forming, gram-positive ovoid

coccus, different variants of which occur in cultured fish in Japan, Europe and Australia, with the Australian

variant being isolated from diseased rainbow trout in freshwater hatcheries in Tasmania and Victoria (Carson

et al. 1993). The emergence of the disease in fish farms is associated with poor water quality and or high

temperatures (Vendrell et al. 2006). Lactococcus garvieae is prominently associated with mastitis in dairy

cows, however activities such as dairy farming in the catchment are not necessarily related to outbreaks of L.

garvieae in fish farms, because isolates from cows and fish are usually genetically distinct (Foschino et al.

2008). However, at least one case exists where a fish isolate had the same biotype as a cow isolate,

suggesting that outbreaks of the same strains in both terrestrial animals and fish could be epidemiologically

related (Vela et al. 2000). Studies have shown a relatively high level of biochemical homogeneity among the

fish strains from different parts of the world, suggesting that different biotypes should not be established

since they would lack epidemiological and taxonomic value (Vendrell et al. 2006).

In Tasmania, disease associated with L. garvieae infection was observed in sea run rainbow trout

(Oncorhynchus mykiss) on at least two occasions in the 1980’s and 1990’s. Disease episodes occurred soon

after sea transfer of covertly infected rainbow trout from hatcheries with a known history of disease due to L.

garvieae, but no transfer of disease to other species of marine fish was recorded (J. Carson, Chief

Microbiologist, Fish Health Unit, DPIWE Tasmania, personal communication 2002). No other infections

with L. garvieae appear to have been recorded in Australian fishes, however in Japan enterococcal infection

caused by Lactococcus garvieae is the major bacterial disease of cultured yellowtail (Seriola

quinqueradiata) (see Kusuda and Salati 1993, 1999). Outbreaks of enterococcal disease in cultured

yellowtail in Japan occur mainly in summer in seacaged juveniles and are associated with poor water quality

and high stocking densities (Kusuda and Salati 1999). This suggests that kingfish (Seriola lalandi)

aquacultured in South Australia and other states may also be susceptible to this bacterium if cultured at high

densities during the summer months. Tilapia and Macrobrachium spp. are two other species that occur in

Australia and are known to be susceptible to L. garvieae (see Vendrell et al. 2006).

_________________________________________________________________________________________________________________________________________________________________________________


98

www.digsfish.com


Outbreaks of L. garvieae on rainbow trout farms in Victoria and Tasmania suggest that the bacterium occurs

naturally in some freshwater environments in the south east of the country. The bacterium is pathogenic for

rainbow trout held in aquaculture under stressful conditions, but it has not been recorded from other native

finfish species in Australia, or Tilapia, nor from freshwater prawns (Macrobrachium spp.) which are another

known susceptible host (Vendrell et al. 2006). The outbreaks of disease soon after sea transfer of rainbow

trout in Tasmania shows that fish exposed in the hatchery can carry L. garvieae covertly in sub-clinical

infections at low intensities. The highest numbers of bacteria in clinically diseased fish usually occur in the

kidney, but the septicaemic nature of the infection means that most internal organs can be infected, including

the brain (Carson et al. 1993, Vendrell et al. 2006). Disease occurs in fish of all sizes (Vendrell et al. 2006).

Cultured rainbow trout and their processing wastes are not widely sold for bait or berley, and Kewagama

Research (2002, 2006) estimated that only 0.4-0.5% of recreational fishers in NSW, Victoria and ACT used

salmonid products as bait (Table 1). However, rainbow trout are widely distributed as food fish and hence

some of this product could be diverted to use as bait or berley around the country.

The bacterium has been isolated from water samples, and can probably persist in the environment for long

periods in both water and sediments. However introduction of sub clinically infected, asymptomatic carrier

fish is the most common route of spread of the disease, with transmission mainly horizontal through the

water between fish that co-habit the same ponds, or by oral or feco-oral routes (Vendrell et al. 2006). These

bacteria are very tolerant of freezing, remaining viable in frozen fish for at least 6 months (Vendrell et al.

2006). These bacteria are susceptible to a broad range of antibiotics and disinfectants, though resistance to

some antibiotics has developed, while vaccines, probiotics and phage therapy have all been successfully

utilised to control the disease (Vendrell et al. 2006).

Lactococcus garvieae is known to occur naturally in some freshwater environments in the south east of the

country, but to date is only known to cause disease in rainbow trout cultured under sub-optimal conditions

(Table 8). Taking into account the relatively small quantities of these that are used as bait or berley (Table

1), but recognising that some of the product sold as food fish could be diverted to use as bait or berley

around the country, the likelihood estimations for the occurrence of Lactococcus garvieae in these


Release assessment for infection of salmonids with Lactococcus garvieae

Commodity

type

Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release

Moderate Moderate Low Very Low Very Low Low

_________________________________________________________________________________________________________________________________________________________________________________


99

www.digsfish.com


Freshwater teleosts in the south east regions of Australia are already at risk of exposure to Lactococcus

garvieae, but no outbreaks of disease have been recorded in any other species except cultured rainbow trout.

The quantities of rainbow trout sold as live bait is likely to be negligible, however recreational fishers may

still catch their own live bait at the fishing location, and a small proportion of anglers may use juvenile live

rainbow trout as bait in freshwater regions. Fresh or frozen cultured rainbow trout are widely sold as food

fish throughout Australia, and some of this could be diverted to use as bait or berley in both freshwater and

marine environments around the country. Because L. garvieae is stable when frozen, a large percentage of

virus is likely to remain viable after thawing, this suggests that direct pathways exist for frozen rainbow trout

products used as bait or berley to enter both freshwater and marine environments, thus potentially exposing

wild fish and crustaceans to viable L. garvieae. Other significant pathways for exposure of fish and

crustaceans to L garvieae would include disposal into natural waters of untreated processing wastes from

commercial processing premises.

Some fish species from regions in Australia where this bacterium is not known to be enzootic (e.g. yellowtail

kingfish, Tilapia and Macrbrachium spp.) are likely to be susceptible to infection by L. garvieae carried in

infected bait or berley, but infection would occur only if sufficient quantities of bacteria (i.e. an infective

dose) was introduced into an area where susceptible hosts were present. Eldar and Ghittino (1999) found the

L. garvieae LD50 for rainbow trout via IP injection was 1.25 x 101 cfu, suggesting the bacterium was very

pathogenic for trout exposed via this route. The LD50 for tilapia via IP injection with L. garvieae was much

higher at 1.4 x 105 cfu (Evans et al. 2009). Chang et al. (2002) found that immersion of rainbow trout in 1 x

106 cfu/ml was sufficient to kill 100% of fish in 1 week. The minimum doses required to infect susceptible

fish and crustacean species with L. garvieae by immersion do not appear to have been established.

Clinically diseased fish infected with L. garvieae can have large numbers of bacteria in their internal organs.

In contrast, many infected fish may carry L. garvieae covertly in sub-clinical infections at very low

intensities (Vendrell et al. 2006). It appears possible, therefore, that an infectious dose could be transmitted

horizontally via the water in the immediate vicinity of bait or berley that was clinically infected with L.

garvieae, but this would probably not occur if the bait or berley was sub-clinically infected. Fish could also

theoretically be exposed to an infectious dose via the per-os route if they consumed a bait fish containing

bacterial levels typical of clinically infected fish. Clinically infected ranbow trout are likely to be

aquacultured only, and some states have bans on use of aquacultured products as bait (T. Hawkesford,

personal communication), but rainbow trout products remain available for bait and berley use if diverted

from product retailed for human consumption. Given that pathways exist for translocation and spread of L.

garvieae into the environment via use of bait or berley, and acknowledging that susceptible hosts do exist in

Australia, but they may require exposure to reasonably high concentrations of the bacterium before they

become infected by the immersion route, the risk of exposure and establishment is non-negligible, and the

likelihood of exposure and establishment of L. garvieae in new fish populations is considered to be Low.


Lactococcus garvieae is considered an opportunistic pathogen that rarely, if ever, causes disease in healthy

unstressed fish. Disease outbreaks associated with L. garvieae are almost exclusively associated with

_________________________________________________________________________________________________________________________________________________________________________________


100

www.digsfish.com

cultured fish when they are injured or held at high densities under poor conditions. Avoidance of disease

outbreaks due to opportunistic bacteria such as L. garvieae is, therefore, based mainly on use of sound

husbandry practices, and as such is not simply related to the presence or absence of the disease agent.

Disease caused by L. garvieae is therefore unlikely to be a problem in wild fish and crustaceans and its

presence is unlikely to have impacts on wild fisheries at the population level. The main consequences appear

those related to possible increased mortality as well as increased costs of production in finfish aquaculture,

where use of antibiotics or production of effective vaccines would be required to prevent significant losses if

culture conditions are allowed to become sub-optimal. Because these bacteria is already present in the

Australian environment, the full extent of any increases in mortality rates that may occur due to translocation

of the disease agents via bait is difficult to assess. These bacteria are not listed by the OIE, or NACA, and

remain on Tasmanias reportable disease list only, hence their spread is unlikely to have large impacts on

trade. Considering all of these factors, establishment of the disease would most likely have mild to moderate

biological consequences, which may be amenable to control, and are not likely to cause irreversible

environmental effects. It is therefore estimated that the consequences of introduction of L. garvieae into

different parts of the Australian environment via use of infected bait or berley would likely be Low.


The unrestricted risk associated with Lactococcus garvieae is determined by combining the likelihood of

entry and exposure (from Table 5) with the consequences of establishment (Table 7). The unrestricted risk

estimate for Lactococcus garvieae does not exceed the ALOP for any of the commodity types, suggesting

that additional risk management for this disease agent is not required at this time.

Risk estimate for infection of salmonids with Lactococcus garvieae

Commodity type Live

finfish

Whole

fresh dead

finfish

Frozen

whole

finfish

Frozen

fillets

Frozen

fish heads

Frozen

guts/offal

Combined likelihood

of release and

exposure

Low Low Low Very Low Very Low Low

Consequences of

establishment


Risk estimation Very low

risk

Very low

risk

Very low

risk

Negligible

risk

Negligible

risk

Very low

risk

_________________________________________________________________________________________________________________________________________________________________________________


101

www.digsfish.com

5.7 Infection of salmonids with Piscirickettsia-like bacteria (PLBs)

5.7.1 Aetiologic agent: Piscirickettsia-like gram negative obligate intracellular bacteria that are members

of the Class Gammaproteobacteria.


5.7.3 Australias status: PLB infections have been reported from cultured Atlantic salmon in Tasmania, and

this disease is reportable in Tasmania (Table 3b).

5.7.4 Epidemiology

In Tasmania, low level mortalities (<5%) in sea cage cultured Atlantic salmon were associated with the

discovery of a rickettsia like organism (RLO) (Elliot 2001) with close affinities to Piscirickettsia salmonis

(see Corbeil et al. 2005, Corbeil and Crane 2009). The onset of disease usually occurs after transfer of fish

from freshwater to seawater net pens (Corbeil and Crane 2009). The source of the disease remains unknown,

but in other parts of the world it has been suggested that wild marine fish are likely candidates as reservoirs

for Piscirickettsia-like bacteria (PLB) (Mauel and Miller 2002). Closely related PLBs have caused disease

in white seabass (Atractoscion nobilis) in California (Chen et al. 2000a), grouper (Epinephelus

melanostigma) in Taiwan (Chen et al. 2000b), and tilapia (Cichlidae) in several countries including Taiwan,

USA and central America, though the disease agents in tilapia may be more closely related to Francisella

than Piscirickettsia (see Colquhoun and Duodu 2011). Indeed, the emergence of Piscirickettsia-like and

Francisella-like gram negative obligate intracellular bacteria has had adverse effects on the profitability and

productivity of an increasing number of marine fish culture industries worldwide (Mauel and Miller 2002,

Colquhoun and Duodu 2011).

Infections with PLBs result in systemic granulomatous infections which affect internal organs, particularly

the spleen, kidney and liver, but skin, gills and virtually every other organ can be damaged, including the

brain (Mauel and Miller 2002, Colquhoun and Duodu 2011). Mortality rate is variable, depending on the

identity of the affected host, the pathogen, and rearing conditions. Coho salmon (Oncorhynchus kisutch)

appear to be particularly susceptible to P. salmonis with heavy mortalities of over 90% being reported in

coho salmon cultured in seawater in Chile, while in Norway mortality in infected Atlantic salmon can be as

low as 0.06% (Mauel and Miller 2002).

Introduction of live, sub-clinically infected, asymptomatic carrier fish is the most common route of spread of

disease due to PLBs, with transmission mainly horizontal through the water between fish that are held in the

same water body at high densities (Mauel and Miller 2002, Colquhoun and Duodu 2011). These bacteria are

somewhat susceptible to antibiotic therapy, but this does not control the disease, while recombinant vaccines

are being developed to better control the disease (Corbeil and Crane 2009).

_________________________________________________________________________________________________________________________________________________________________________________


102

www.digsfish.com


Outbreaks of PLB in seacage cultured Atlantic salmon in Tasmania suggest that PLBs occur naturally in

some areas of the marine environment in the south east of the country. However, there is no evidence to date

indicating whether PLBs occur in the marine waters of mainland Australia, although there are no barriers to

migration of marine fish from Tasmania to mainland waters, and some species do migrate between these two

areas (Stanley 1978). In Australia, PLBs can be pathogenic for salmonids held in aquaculture, but they have

not been observed to cause disease in other native finfish species. However, sciaenids such as white seabass

are known to be susceptible to PLB infection (Chen et al. 2000a), and mulloway (Argyrosomus

hololepidotus) is a sciaenid that is cultured in some areas of mainland Australia. Hence it is likely that

susceptible hosts for PLBs do occur in mainland Australia, with some occurring under conditions known to

favour outbreaks of disease due to PLBs.

Given that PLBs are obligate intracellular disease agents, they appear less likely to survive for long periods

in the marine environment than other heterotrophic bacteria (AQIS 1999b). Indeed, some isolates of PLBs

from marine fish are immediately inactivated in freshwater (Lannan and Fryer 1994, Chen et al. 2000a), but

may survive for up to 2 weeks in seawater, with higher titres remaining at 5°C compared to 15°C after 14

days (Lannan and Fryer 1994). Water temperature is important for disease transmission, with bacterial

replication optimal between 15 and 18°C, becoming greatly retarded above 20°C and below 10°C, with

replication ceasing above 25°C (AQIS 1999b). The highest numbers of bacteria in clinically diseased fish

usually occur in the spleen, kidney, and liver, but the septicaemic nature of the infection means that most

internal organs can be infected (Mauel and Miller 2002). Fish of all ages are susceptible to the disease

(Corbeil and Crane 2009). Cultured salmonids and their processing wastes are not sold for bait or berley

(Grant Pullen, DPIWE Tasmania, personal communication 8 April 2011), but fresh or frozen (but not live)

Atlantic salmon are widely distributed as food fish and hence some product could be diverted to use as bait

or berley around the country. However, a large proportion of PLBs are inactivated by freezing (AQIS

1999b), with freezing reducing infectivity of the PLB from white seabass by 100 fold (Chen et al. 2000a).

PLBs probably occur naturally in the marine environment off Tasmania, but to date they have only infected

cultured Atlantic salmon (Table 8). Taking into account the relatively small quantities of these that are likely

to be used as bait or berley (Table 1), and the prevalence of the disease agent, the likelihood estimations for

the occurrence of Piscirickettsia-like bacteria in these commodities are listed below.

Release assessment for infection of salmonids with PLBs

Commodity

type

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release

Moderate Very low Extremely

low

Very low Very low

_________________________________________________________________________________________________________________________________________________________________________________


103

www.digsfish.com


Several species of marine fish are likely to be exposed to the PLB in the Tasmanian marine environment,

however the disease agent has only been recorded from cultured Atlantic salmon. Some marine fish species

in regions where PLBs are not enzootic are likely to be susceptible to infection by PLBs carried in infected

bait or berley, but infection would occur only if sufficient quantities of bacteria (i.e. an infective dose) were

introduced into an area where susceptible hosts were present.

Susceptible fish can become infected with PLBs via horizontal transmission through the water with entry via

the skin, intestine and gills (Smith et al. 2004), but it is not known whether vertical transmission occurs

(Corbeil and Crane 2009). Smith et al. (1996a) found that the LD50 for coho salmon infected with P.

salmonis via IP injection was 101.9 TCID50/g, and 102.4 TCID50/g for rainbow trout, and virulence of other

PLBs injected via the IP route was variable (c. <103 - 105 TCID50/fish) depending on the isolate used (House

et al. 1999). Bath immersion is the route of infection most applicable to assessing the risk of exposure via

use of bait and berley, and Birbeck et al. (2004) found that infectivity of a virulent (LD50 by IP injection <2

x 103 rickettsial units) Scottish isolate of P. salmonis in Atlantic salmon via the bath immersion route was

much reduced, with a 1 hr bath challenge in 105 TCID50/mL causing only 10% mortality after 18 days, while

direct application of 2.5 x 106 TCID to the skin failed to induce any mortalities within 42 days. In contrast,

Smith et al. (2004) used coho salmon exposed to 103.7 TCID50/fish of P. salmonis to determine that contact

with skin (32% cumulative mortality) was most effective entry portal, followed by intestine (16% mortality)

and gills (12% mortality).

Clinically diseased fish can have large numbers of PLBs in internal organs, gills and skin, but infected fish

can also carry PLBs covertly in sub-clinical infections at very low intensities (Arkush et al. 2006). The

minimum dose required to infect susceptible Australian native fish species with PLBs by immersion have not

been determined, however the disease can be transferred horizontally by cohabitation of susceptible hosts

with infected fish (Mauel and Miller 2002). It may be possible, therefore, that an infectious dose could be

transmitted horizontally via the water in the immediate vicinity of bait or berley that was clinically infected

with PLBs, but this would probably not occur if the bait or berley was sub-clinically infected. Clinically

infected fish are likely to be aquacultured only, but cultured salmonids and their processing wastes are not

sold for bait or berley (Grant Pullen, DPIWE Tasmania, personal communication 8 April 2011). However,

fresh or frozen Atlantic salmon are widely distributed as food fish and some product could be diverted to use

as bait or berley. Pathways therefore still exist for translocation and spread of PLBs into the environment via

use of bait or berley, and susceptible hosts may exist in parts of the country where PLBs are not known to be

endemic, but susceptible hosts may require exposure to reasonably high concentrations of the bacterium

before they become infected by the immersion route. Nevertheless, the risk of exposure and establishment is

non-negligible, and the likelihood of exposure and establishment of PLBs in new fish populations is

considered to be Low.


Disease outbreaks associated with PLBs are almost exclusively associated with cultured fish when they are

held at high densities, therefore RLBs are unlikely to be problematic in wild fish and their presence is

unlikely to have impacts on wild fisheries at the population level. The main consequences appear those

_________________________________________________________________________________________________________________________________________________________________________________


104

www.digsfish.com

related to possible increased mortality as well as increased costs of production in finfish aquaculture, where

use of antibiotics or production of effective vaccines would be required to prevent significant losses if

culture conditions are allowed to become sub-optimal. Because these bacteria are already present in some

parts of the Australian environment, the full extent of any increases in mortality rates that may occur due to

translocation of these disease agents via bait is difficult to assess. These bacteria are not listed by the OIE, or

NACA, and remain on Tasmanias reportable disease list only, hence their spread is unlikely to have much

impact on trade. Considering these factors, establishment of the disease would most likely have mild to

moderate biological consequences, which may be amenable to control, and are not likely to cause irreversible

environmental effects. It is therefore estimated that the consequences of introduction of PLBs into different

parts of the Australian environment via use of infected bait or berley would likely be Low.


The unrestricted risk associated with Piscirickettsia-like bacteria is determined by combining the likelihood

of entry and exposure (from Table 5) with the consequences of establishment (Table 7). The unrestricted

risk estimate for Piscirickettsia-like bacteria does not exceed the ALOP for any of the commodity types,

suggesting that additional risk management for this disease agent is not required at this time.

Risk estimate for infection of salmonids with PLBs

Commodity type Whole fresh

dead finfish

Frozen

whole finfish

Frozen fillets Frozen fish

heads

Frozen

guts/offal

Combined likelihood of

release and exposure

Low Very low Extremely

low

Very low Very low

Consequences of

establishment

Low Low Low Low Low


risk

Negligible

risk

Negligible

risk

Negligible

risk

Negligible

risk

_________________________________________________________________________________________________________________________________________________________________________________


105

www.digsfish.com

5.8 Infection of finfish with Yersinia ruckeri (Yersiniosis)

5.8.1 Aetiologic agent: Yersinia ruckeri, a gram negative bacterium within the family Enterobacteriaceae.


5.8.3 Australias status:. Reported in Tasmania, Victoria and NSW, and is not a reportable disease in any

state, although enteric redmouth disease caused by Y. ruckeri (Hagerman strain) is a reportable disease in all

states (Table 3b).

5.8.4 Epidemiology

The bacterium Yersinia ruckeri causes disease in cultured freshwater fishes worldwide, mainly salmonids,

but also eels, goldfish, carp and others (Tobback et al. 2007, Carson and Wilson 2009). Infection with Y.

ruckeri results in bacterial septicaemia and disease is most commonly detected due to exophthalmos and

blood spots in the eye. The severity of the disease is dependant upon environmental conditions, being most

problematic at higher water temperatures and stocking densities, as well as the virulence of the variant of the

bacterium involved (Tobback et al. 2007). Acute infections in trout with the 'Hagerman’ strain are referred

to as enteric red mouth (ERM), however in Australia the 'Hagerman’ strain is considered exotic (Humphrey

et al. 1987), and a milder form of the disease that occurs in Atlantic salmon and rainbow trout is termed

yersiniosis (Carson and Wilson 2009). In rainbow trout infected with Y. ruckeri, the disease can affect fish

of all age classes. In larger fish the disease is chronic, but acute disease outbreaks with cumulative losses

reaching 35% can occur in small fish up to fingerling size, especially if they are reared in suboptimal

conditions at elevated temperatures or with poor water quality (Carson and Wilson 2009). Survivors of

disease outbreaks can become asymptomatic carriers of the bacterium (Hunter et al. 1980).

The first signs of the disease observed in juvenile salmonids are increased mortalities, followed by changes

in behaviour including swimming near the surface, moving sluggishly, darkening and inappetence, followed

by development of a marked unilateral or bilateral exophthalmos, often with patches of haemorrhagic

congestion on the iris of the eye (Carson and Wilson 2009). In rainbow trout, subcutaneous haemorrhage in

the mouth and throat is strongly indicative of the disease and hence the term enteric red mouth. In clinically

diseased fish, Y. ruckeri may occur in kidney, liver, spleen and the distal portion of the gastro-intestinal tract,

a site from which bacteria may be excreted to the water column (Carson and Wilson 2009). Transmission of

Y. ruckeri is direct and horizontal with the primary source being large numbers of bacteria shed in the faeces

of infected or carrier fish. In clinically diseased fish, bacteria are detected in the blood, while histologically,

salmon fry may contain overwhelming numbers of bacteria with high concentrations detectable in

macrophages of kidney and liver sinusoids (Carson and Wilson 2009).


Known hosts for Yersinia ruckeri in Australia include cultured Atlantic salmon, with rare isolations from

rainbow, brown and brook trout (Carson and Wilson 2009), but other known susceptible species that occur in

Australia include redfin perch, eels, carp and goldfish (Herfort 2004). In Australia, two biovariants of Y

_________________________________________________________________________________________________________________________________________________________________________________


106

www.digsfish.com

ruckeri are known to occur: serotype O1b, (biotype 1) and serotype O1, non-O1b, (biotype 2) (Carson and

Wilson 2009). Outbreaks of yersinosis in Australia have been reported in salmonids in Tasmania, Victoria

and NSW (Humphrey et al. 1987, Langdon 1988, Carson and Wilson 2009), and given its ubiquitous

distribution worldwide, it is likely that the bacterium is enzootic in the Australian environment in other

regions as well.

Infected fish may carry Y. ruckeri in sub-clinical infections at very low intensities, with carriers usually

reverting to a diseased state during stressful events (Hunter et al. 1980). The highest numbers of bacteria in

clinically diseased fish usually occur in the internal organs such as the kidney, liver, spleen and

gastrointestinal tract, while the bacterium is mainly present in the lower intestine of carrier fish (Hunter et al.

1980). The bacterium is able to persist in the environment for long periods in water (>4 months, see Hine

and McDiarmid 1997), sediments and biofilms (Tobback et al. 2007). Yersinia ruckeri is susceptible to a

broad range of antibiotics and disinfectants, though resistance to some antibiotics has developed (Tobback et

al. 2007). Vaccines against Y. ruckeri have been successfully developed, and disease control using

probiotics has also been achieved (Tobback et al. 2007). These bacteria are tolerant of freezing and will

remain viable in frozen fish for at least 6 months (Anderson et al. 1994), though as for other bacteria, there

may be some loss of viability upon thawing.

Those fishes used as bait or berley in Australia that have been recorded as being infected by Y. ruckeri

include goldfish, and salmonids (Table 8), while other bait species known to be susceptible to the bacterium

include carp and redfin perch. Live goldfish may be illegally used as bait by some recreational fishers

(Lintermans 2004), while cultured salmonids and their processing wastes are not sold for bait or berley, but

fresh or frozen trout and Atlantic salmon are widely distributed as food fish and hence some of this product

could be diverted to use as bait or berley around the country. Taking these factors into account, the

likelihood estimations for the occurrence of Y. ruckeri in these commodities are listed below.

Release assessment for infection of finfish with Yersinia ruckeri

Commodity

type

Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release

Moderate Moderate Low Very Low Very Low Low


Freshwater teleosts in the south east regions of Australia are already at risk of exposure to Yersinia ruckeri,

but no outbreaks of disease have been recorded in any species other than cultured salmonids. No salmonids

are known to be sold as live bait, however recreational fishers may still catch their own live bait at the

fishing location, and a small proportion of anglers may use live rainbow trout or goldfish as bait in

freshwater regions. Fresh or frozen cultured Atlantic salmon and rainbow trout sold as foodfish could also

_________________________________________________________________________________________________________________________________________________________________________________


107

www.digsfish.com

be diverted to use as bait in both freshwater and marine environments around the country. Because Y.

ruckeri is stable when frozen, a large percentage of bacteria are likely to remain viable after thawing. This

shows that direct pathways exist for viable Y. ruckeri to enter both freshwater and marine environments in

several products that could be used as bait or berley. Other likely pathways for exposure of marine and

freshwater fish to Y. ruckeri would include disposal into natural waters of untreated processing wastes from

commercial processing premises.

At least some species of fish in Australia (e.g. redfin perch, carp) are likely to be susceptible to infection by

Y. ruckeri carried in infected bait or berley, but infection would occur only if sufficient quantities of bacteria

(i.e. an infective dose) was introduced into an area where susceptible hosts were present. Hunter et al.

(1980) observed that live carriers transmitted Y. ruckeri to clinically healthy fish via co-habitation when the

temperature was raised to 25°C, whereas unstressed carrier fish did not. The infectious dose for transmission

of Y. ruckeri to rainbow trout by immersion is high, in the order of 107 - 108 CFU/mL at 16 to 23°C, which

caused between 0 and 30% mortality depending on the biovariant of the bacterium used (Tobback et al.

2009). The route of infection of fish exposed to high doses of Y. ruckeri via bath immersion was mainly the

gills, followed by the skin and gut, with rapid spread and persistence of infection of internal organs only

occurring in fish exposed to virulent strains (Tobback et al. 2009). The bacterium was reisolated from the

internal organs of clinically normal fish at concentrations of between 102 - 103cfu/g of tissue, and from

clinically diseased fish at 107 - 108cfu/g of tissue (Tobback et al. 2009).

Clinically diseased fish infected with Y. ruckeri can have large numbers of bacteria in their internal organs,

but many infected fish may carry Y. ruckeri covertly in sub-clinical infections at low intensities (Hunter et al.

1980, Tobback et al. 2009). It appears theoretically possible, therefore, that an infectious dose could be

transmitted horizontally via the water in the immediate vicinity of bait or berley that was clinically infected

with Y. ruckeri, but this would probably not occur if the bait or berley was sub-clinically infected. Fish

could also theoretically be exposed to an infectious dose via the per-os route if they consumed a bait fish

containing bacterial levels typical of clinically infected fish. Clinically infected fish are likely to be

aquacultured only, and some States have bans on use of aquacultured products as bait, but salmonid products

remain available for bait or berley use if diverted from product retailed for human consumption. Whether

any clinically infected salmonids are sold for human consumption is not known at this time. Given that

pathways exist for translocation and spread of Y. ruckeri into the environment via use of bait, and

acknowledging that susceptible hosts do exist in Australia, but they may require exposure to reasonably high

concentrations of the bacterium before they become infected by the immersion or per-os routes, the risk of

exposure and establishment is non-negligible, and the likelihood of exposure and establishment of Y. ruckeri

in new fish populations is considered to be Very low.


Yersinia ruckeri is considered an opportunistic pathogen that rarely causes disease in healthy unstressed fish.

Disease outbreaks associated with Y. ruckeri are almost exclusively associated with cultured salmonids when

they are injured or held at high densities under poor conditions. Avoidance of disease outbreaks due to

opportunistic bacteria such as Y. ruckeri is, therefore, based mainly on encouragement of sound husbandry

practices, and as such is not necessarily related to the simple presence or absence of the disease agent. Y.

ruckeri is highly unlikely to cause disease in wild fish and its presence is unlikely to have impacts on wild

_________________________________________________________________________________________________________________________________________________________________________________


108

www.digsfish.com

fisheries at the population level. The main consequences appear those related to possible increased mortality

as well as increased costs of production in salmonid aquaculture, where use of antibiotics or production of

effective vaccines would be required to prevent losses if culture conditions are allowed to become sub-

optimal. Because these bacteria is already present in the Australian environment, the full extent of any

increases in mortality rates that may occur due to translocation of the disease agents via bait is difficult to

assess. These bacteria are not listed by the OIE, or NACA, or on any State reportable disease list, hence

their spread is unlikely to have adverse impacts on trade. Considering all of these factors, establishment of

the disease would most likely have mild biological consequences, which are amenable to control, and are not

likely to cause irreversible environmental effects. It is therefore estimated that the consequences of

introduction of Yersinia ruckeri into different parts of the Australian environment via use of infected bait or

berley would likely be Very low.


The unrestricted risk associated with Yersinia ruckeri is determined by combining the likelihood of entry and


Yersinia ruckeri does not exceed the ALOP for any of the commodity types, suggesting that additional risk

management for this disease agent is not required at this time.

Risk estimate for infection of finfish with Yersinia ruckeri

Commodity type Live

finfish

Whole

fresh dead

finfish

Frozen

whole

finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Combined


and exposure

Very Low Very Low Very Low Extremely

Low

Extremely

Low

Very Low

Consequences of

establishment

Very Low Very Low Very Low Very Low Very Low Very Low

Risk estimation Negligible

risk

Negligible

risk

Negligible

risk

Negligible

risk

Negligible

risk

Negligible

risk

_________________________________________________________________________________________________________________________________________________________________________________


109

www.digsfish.com

5.9 Infection of finfish with Aphanomyces invadans (Epizootic Ulcerative Syndrome)

5.9.1 Aetiologic agent: Aphanomyces invadans, an oomycete fungus from the Family Saprolegniaceae.


5.9.3 Australias status: Reported in QLD, NSW, Victoria, SA, NT and WA and is a reportable disease in

all states excluding QLD and the ACT (Table 3b).

5.9.4 Epidemiology

Systemic mycoses consistent with epizootic ulcerative syndrome (EUS) caused by the fungus Aphanomyces

invadans were first reported in Australia in the early 1970’s (McKenzie and Hall 1976, Fraser et al. 1992,

Arthur and Subasinghe 2002). The emergence of EUS throughout Australia and Asia represented a regional

panzootic caused by the spread of a single clone of A. invadans throughout the region (Callinan et al. 1995,

Lilley and Roberts 1997, Lilley et al. 1997, Roberts 2001). Rhabdoviruses have also been associated with

some EUS outbreaks, and secondary gram-negative bacteria invariably infect EUS lesions, however A.

invadans is recognised as the primary disease agent (OIE 2010b). EUS is mainly problematic in Australasia

and the Indo-Pacific region (Fraser et al. 1992, Callinan et al. 1995, Lilley and Roberts 1997), however

similar ulcerative mycotic diseases were recorded from estuarine fish along the east coast of the United

States for a number of years (Dykstra et al. 1989), and A. invadans was also eventually implicated in that

region (Vogelbein et al. 2001, Blazer et al. 2002, Saylor et al. 2010). Over 76 species of wild and cultured

fishes are known to be naturally affected, though some species, including carp (Cyprinus carpio), Nile tilapia

(Oreochromis niloticus) and milkfish (Chanos chanos) are considered to be resistant to infection (OIE

2010b). The disease is characterised by reddened ulcerative dermal lesions that become visible on the

surface of affected fish a few weeks after periods of heavy rainfall (Fraser et al. 1992, Kiryu et al. 2003).

The presence of the fungus causes an invasive necrotising dermatitis with a prominent granulomatous host

response in affected muscle, and can result in epizootic mortalities of up to 100% in affected fish populations

that occur in water less than 9 ppt salinity over a wide temperature range (18 - 30°C) (Fraser et al. 1992,

Callinan et al. 1995, Lilley and Roberts 1997, Saylor et al. 2010). It is believed that only the zoospores are

capable of attaching to the damaged skin of fish and germinating into hyphae (OIE 2010b). If the zoospores

cannot find the susceptible species or encounter unfavourable conditions, they can form secondary zoospores

which can encyst in the water or pond environment until conditions favour their activation. How the

Aphanomyces pathogen or its spores survive after the outbreak is still unclear as outbreaks usually occur

about the same time every year in endemic areas (OIE 2010b). Treatment is only applicable to captive fish

and often has limited success because antifungal agents tend to be toxic to the fish, or do not penetrate the

body deeply enough to reach the pathogen, which invades deep into the muscle tissue and internal organs

(Campbell et al. 2001, Saylor et al. 2010).

Because A. invadans is not tolerant of salt and can only be transmitted effectively in brackish and

freshwaters, the spread of the disease agent throughout many parts of Australia and Asia has most likely

been through translocation of live fish (Callinan et al. 1995, Lilley et al. 1997), though natural movements of

wild fish during flood events could also explain some of this spread. There is no information to indicate that

_________________________________________________________________________________________________________________________________________________________________________________


110

www.digsfish.com

fish can be lifelong carriers of A. invadans. Most EUS infected fish die, and although some mild or even

moderately infected fish can recover, they are unlikely to be lifelong carriers (OIE 2010b).


In Australia EUS is primarily a disease of estuarine fish, particularly mullet (Mugil cephalus), yellowfin

bream (Acanthopagrus australis) and whiting (Sillago spp.), however, EUS has been reported from a wide

range of wild fishes (Pearce 1990, Humphrey and Pearce 2004) and also cultured silver perch (Bidyanus

bidyanus) and Murray cod (Maccullochella peelii) (NACA 2010). EUS has been recorded from Queensland,

NSW, the Northern Territory and Western Australia and is restricted to areas of low salinity (0 - 9 ‰).

Mullet are highly susceptible to EUS (McKenzie and Hall 1976, Fraser et al. 1992, Roberts 2001) and this

species is very popular for use as live and fresh dead bait for a variety of estuarine fish and crab species

throughout Australia (Ross 1995). Whiting (Sillago ciliata) and bream (Acanthopagrus australis) are also

commonly used as bait and are sometimes translocated into different river systems in southern QLD and

northern NSW through movements of live fish by recreational fishers (Diggles, personal observation). The

families and species of fishes known to be susceptible to EUS and used as bait in Australia (Table 8) include

F. Ambassidae, goldfish, F. Cichlidae, F. Eleotridae, F. Lutjanidae, F. Melanotaeniidae, F. Muglidae,

Nematalosa spp., F. Sillaginidae, F. Sparidae, F. Teraponidae, and Toxotes spp.

Estuarine and freshwater finfish are not generally sold as live bait in retail outlets, but some (e.g. mullet,

goldfish, Nematalosa spp., F. Sillaginidae, F Teraponidae) are sometimes collected at the fishing site and

used as live bait by recreational fishers. Fresh dead fish are sometimes available locally from commercial

fishing co-operatives, while frozen baitfish (e.g. mullet) are used in large quantities and are widely available

at thousands of retail outlets (tackle shops, service stations, supermarkets) throughout Australia (Kewagama

Research 2002, 2007, Table 1). Hyphae of A. invadans that penetrate the muscle of infected fish are well

protected from desiccation and chemical treatments (Campbell et al. 2001, Saylor et al. 2010) and like other

Aphanomyces spp. are likely to survive in fresh chilled product for 2 or more weeks (Oidtmann et al. 2002).

Zoospores of A. invadans can encyst and survive in the environment for at least 19 days provided salinity

does not exceed 4 ppt (Kiryu et al. 2003, OIE 2010b). However, it is unlikely that A. invadans is resistant to

freezing, as has been shown that hyphae and spores of a related fungus (Aphanomyces astaci) are inactivated

by freezing to -20°C for 72 hours (Oidtmann et al. 2002), and indeed, freezing at -5°C for 72 hours is also

effective for inactivating both spores and hyphae of Aphanomyces spp. (Oidtmann et al. 2002). Taking this

information into account, together with the large quantities of susceptible fishes used as bait or berley, the

likelihood estimations for the occurrence of A. invadans in these commodities are listed below.

Release assessment for infection of finfish with Aphanomyces invadans

Commodity

type

Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release

High Moderate Extremely

low

Extremely

low

Extremely

low

Extremely

low

_________________________________________________________________________________________________________________________________________________________________________________


111

www.digsfish.com


Freshwater and estuarine teleosts in several regions of Australia are already at risk of exposure to A.

invadans, but it is not known to what extent that freshwater and estuarine teleosts in other parts of the

country are exposed to A. invadans by natural movements of fish. Mullet and other baitfish known to be

susceptible to A. invadans are sold frozen throughout the country via retail outlets such as fishing tackle

shops, service stations and supermarkets. However freezing for more than 3 days is likely to deactivate both

hyphae and zoospores, virtually eliminating the risk of transfer of infection via frozen bait. In contrast,

movements of live fish and fresh (chilled, unfrozen) product from areas where A. invadans is known to occur

could transmit the disease agent to new regions, but only if infected bait was used in low salinity waters at

temperatures near the optima for the pathogen (18 – 30°C) (Kiryu et al. 2003, 2005). Infection by A.

invadans is horizontal through the water via contact with infective zoospores which cease to become

infective at salinities between 2 and 4 ppt (Blazer et al. 2002; Kiryu et al. 2003, 2005). Nevertheless, A.

invadans has a wide host range, and it is likely that susceptible hosts will occur in at least some areas where

the disease agent is currently not recorded. However, infection would occur only if sufficient quantities of

viable zoospores (i.e. an infective dose) were introduced into an area where susceptible hosts were present.

Aphanomyces invadans is extremely virulent to some hosts, with the LD50 (lethal dose killing 50% of

exposed fish) for menhaden via inoculation of zoospores into the skin being 9.7 zoospores (Kiryu et al.

2003), however, in that study some fish receiving an estimated single zoospore developed infections that

resulted in death (Kiryu et al. 2003). Aphanomyces invadans was also pathogenic via the immersion route,

but a much higher dose was required, with fish exposed to 100 zoospores/ml exhibiting 14% lesion

prevalence and 11% mortality (Kiryu et al. 2003). Mortality rates increased, however, in fish which had

existing skin damage prior to exposure (handled by a net), with those exposed to the same concentration of

zoospores experiencing significantly higher lesion prevalence (64%) and mortality (64%) (Kiryu et al. 2003).

Unfrozen bait has a limited shelf life and is usually only supplied by local fisheries co-operatives and is

unlikely to be translocated long distances. Similarly, finfish for use as live bait are generally not

commercially available in Australia, and the quantities of translocated live fish are likely to be relatively

small. However, A. invadans is a highly infectious, highly pathogenic, highly invasive primary pathogen in

susceptible fish species (Kiryu et al. 2003). Because of this, the risk of exposure and establishment is non-

negligible, and the likelihood of exposure and establishment of A, invadans in new fish populations is

considered to be Moderate.


When fish become infected with A. invadans in a new area, mortality can occur in all size classes of fish, and

therefore this disease has the potential to cause significant damage to fish populations. While the full extent

of its host range remains unknown, A. invadans is known to cause mortality in a wide range of freshwater

and estuarine fishes, including commercially important fisheries and aquaculture species. EUS is listed by

the OIE and NACA and is a reportable disease in all States except QLD and the ACT (Table 3b). Hence the

spread of this pathogen can still have adverse impacts on national or international trade. Considering all of

these factors, establishment of the disease in new areas would have significant biological consequences,

which may not be amenable to control, and could cause significant environmental effects. It is therefore

_________________________________________________________________________________________________________________________________________________________________________________


112

www.digsfish.com

estimated that the consequences of introduction of A. invadans into different parts of the Australian

environment via use of infected bait would likely be Moderate.


The unrestricted risk associated with Aphanomyces invadans is determined by combining the likelihood of


estimate for Aphanomyces invadans exceeds the ALOP for live finfish and whole fresh finfish, suggesting

that additional risk management is required for this disease agent in these commodities.

Risk estimate for infection of finfish with Aphanomyces invadans

Commodity type Live

finfish

Whole

fresh dead

finfish

Frozen

whole

finfish

Frozen

fillets

Frozen

fish heads

Frozen

guts/offal

Combined


and exposure

Moderate Moderate Extremely

low

Extremely

low

Extremely

low

Extremely

low

Consequences of

establishment

Moderate Moderate Moderate Moderate Moderate Moderate

Risk estimation Moderate

risk

Moderate

risk

Negligible

risk

Negligible

risk

Negligible

risk

Negligible

risk

_________________________________________________________________________________________________________________________________________________________________________________


113

www.digsfish.com

5.10 Infection of finfish with Microsporidians

5.10.1 Aetiologic agent: Intracellular protozoan parasites of fishes from the Class Microsporea.


5.10.3 Australias status: Microsporeans have been reported from finfish in all states, and no microsporeans

from finfish are listed as reportable diseases in Australia (Table 3b).

5.10.4 Epidemiology

Microsporeans are obligate, intracellular parasites that infect arthropods, fish, and mammals (Lom and

Dykova 1992). In fish, microsporidian infections can occur in various tissues, often in grossly visible cysts

(xenomas) in the gills, musculature or internal organs. The lifecycle is usually direct, with horizontal

transmission via ingestion of infective spores in the water (Kent et al. 1995) after spore release from skin,

faeces and urine of live hosts or after decomposition of dead hosts (Lom et al. 2000). An intermediate host

may be involved in the lifecycle of some microsporidian species (Vossbrinck et al. 1998) and crustaceans

may act as vectors (Canning and Lom 1986). Host specificity varies, but can be low with some

microsporidian species infecting a wide range of hosts (Lom and Dykova 1992). Free spores are resistant to

external conditions and can remain infective for many months, during which time they may be ingested by

predators or scavengers, while infection can also occur through predation on infected individuals (Lom and

Dykova 1992). In some parts of the world, microsporidian parasites have caused serious disease and reduced

the marketability of product obtained from populations of wild and cultured fish (Kent et al. 1995,

Sutherland 2002, Miller 2009). Affected fish can exhibit anomalous behaviour, become emaciated, and

develop grossly visible lesions and/or discolouration of affected organs (Lom and Dykova 2002).


In Australia, a range of finfish species used as bait are known to harbour microsporidian infections (Table 8),

including members of the Families Clupeidae, Eleotridae, Engraulidae, Gempylidae, Labridae, and

Monocanthidae, while a range of other hosts are also likely to be infected by microsporidians that are

currently unknown and/or undescribed. These host species will be found in a range of environments

throughout the country, however the extent of the distributions of the various species of microsporidian

parasites remains largely unknown. Due to the large numbers of fishers who utilise marine and freshwater

finfish as bait (Table 1, Kewagama Research 2002, 2007), large volumes of fish species likely to be

susceptible to microsporidians are being used as bait and/or berley around the country. Most of this is likely

to be frozen whole baitfish, though use of lesser quantities of fresh chilled baitfish, and small quantities of

live bait collected by recreational fishers, is also likely to occur.

The lifecycle of most microsporidians that infect fish is direct, with horizontal transmission via exposure to

infective spore stages by immersion or per-os routes. The likelihood of release will depend on the ability of

infective stages to remain viable under the conditions of use of their hosts as bait or berley, and it appears

that microsporidian spores can remain viable in the natural environment for months to years. For example,

_________________________________________________________________________________________________________________________________________________________________________________


114

www.digsfish.com

spores of Loma salmonae remained viable when stored in freshwater or seawater at 4°C for up to 95 days

(Shaw et al. 2000), and spores of Glugea stephani remained viable after 17 months at 5°C (Amigo et al.

1996). In contrast, L. salmonae spores were unable to infect salmon after being frozen at -20°C for 24 hours

(Shaw et al. 2000). However, it appears that the viability of microsporidian spores after freezing varies

widely, depending on the species studied. Amigo et al. (1996) found 3.6% of spores of G. stephani remained

infective after being frozen at -19°C for 24 hours. Similarly, spores of Nosemea apis from honey bees

remained viable after 24 hours at -20°C, but spores from Encephalitozoon spp. that infect terrestrial

vertebrates became deactivated after identical treatment (Li and Fayer 2006). In contrast, Koudela et al.

(1999) found that spores of Encephalitozoon cuniculi survived freezing for 24 hours at -20°C and remained

infective. Clearly, given the paucity of data available for the freeze resistance of spores from

microsporidians that infect fishes in Australia, it is not possible to make general conclusions as to whether

freezing is likely to deactivate spores of some or all of these disease agents.

Taking into account the large quantities of finfish products used as bait or berley (Table 1), and also the fact

that microsporidian infections have been recorded from a range of fishes used as bait throughout the country,

but usually at low prevalences, the likelihood estimations for the occurrence of viable microsporidian

parasites in these commodities are listed below.

Release assessment for infection of finfish with microsporidians

Commodity

type

Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release

High Moderate Low Low Low Low


Teleosts throughout marine and freshwater environments in Australia are likely to be already at risk of

natural exposure to infective stages of microsporidians. However, large quantities of fresh or frozen whole

or processed marine fish are offered for sale as bait or berley via wholesale outlets such as commercial

fishing co-operatives, or packaged for resale at retail outlets such as fishing tackle shops, service stations and

supermarkets. Finfish infected with microsporidians often show grossly visible signs of disease such as

xenomas in the flesh, which would be identified and rejected during normal quality control procedures used

for fresh or frozen food fish, largely eliminating the pathway of exposure via diversion of food fish for use as

bait or berley. However, even heavily infected fish with gross signs of disease would still be harvested and

considered acceptable for use as bait. This suggests that direct pathways exist for translocated fish products

infected by microsporidians to enter both freshwater and marine environments via their use as bait or berley,

thus potentially exposing potentially susceptible wild fish to viable infective stages of novel microsporidians.

However, infection and establishment would occur only if sufficient quantities of infective stages (i.e. an

_________________________________________________________________________________________________________________________________________________________________________________


115

www.digsfish.com

infective dose) were introduced into an area where susceptible hosts were present under conditions suitable

for transmission.

Most microsporidians are transmitted directly (Kent et al. 1995), but the minimum infective dose of viable

spores required for successful transmission has not been determined for the majority of species, and will also

probably vary depending on the identity of the host and its immune status (Shaw et al. 2001, Rodriguez-

Tovar et al. 2011). Infection can be achieved by co-habitation with infected fish, with ingestion of spores

occurring via the per-os route (Shaw et al. 2001). After ingestion, the spores enter the stomach, germinate in

response to the pH change and the sporoplasm penetrates an epithelial cell of the gut wall (Kent and Speare

2005). This is followed by multiplication within the host cell by merogony, which is then followed by

sporogony (Lom and Dykova 1992, Shaw et al. 2001, Kent and Speare 2005, Rodriguez-Tovar et al. 2011).

When this sequence of infection is considered, it is clear that infection can be achieved after exposure to a

dose as small as a single viable spore, though the dose required to cause host mortality will depend on many

factors (Rodriguez-Tovar et al. 2011). One xenoma from a lightly infected fish may contain thousands of

spores, while one heavily infected fish with many xenomas may contain literally millions of spores (Lom and

Dykova 1992).

Given that pathways exist for translocation and spread of viable microsporidians into the environment via

use of bait, and acknowledging that some microsporidians can infect a range of susceptible hosts (but others

may not), and the infective doses required for transmission may be very small in comparison to the parasite

burden carried by a single infected fish, the risk of exposure and establishment is non-negligible, and the

likelihood of exposure and establishment of microsporidians in new fish populations is considered to be

Moderate.


Fish of all size classes can become infected with microsporidians. In susceptible species, disfigurement and

reduction of market value of affected fishes can result in economic losses, while heavy mortalities of wild

and cultured fish have been recorded in some parts of the world where microsporidian outbreaks have

occurred. This suggests that some microsporidians have the potential to cause significant damage to fish

populations and aquaculture, though there are examples of successful treatment of microsporidian infections

through use of vaccines and immunostimulants (Rodriguez-Tovar et al. 2011). While the full range of

microsporidian parasites and their susceptible hosts in Australia remain to be determined, microsporidians

have already been recorded from a wide range of teleosts in both freshwater and marine environments. No

microsporidian diseases of teleosts are listed by the OIE or NACA and none are reportable diseases in any

State (Table 3b). Hence the spread of these disease agents is unlikely to have adverse impacts on trade.

Considering all of these factors, establishment of microsporidians in new areas would have moderate

biological consequences, which may be amenable to control, and could cause some unwanted environmental

effects. It is therefore estimated that the consequences of introduction of microsporidians into different parts

of the Australian environment via use of infected bait would likely be Low.

_________________________________________________________________________________________________________________________________________________________________________________


116

www.digsfish.com


The unrestricted risk associated with microsporidian infection is determined by combining the likelihood of


estimate for microsporidians exceeds the ALOP for live finfish and whole fresh dead finfish, suggesting that

additional risk management is required for these disease agents in these commodities.

Risk estimate for infection of finfish with microsporidians

Commodity type Live

finfish

Whole

fresh dead

finfish

Frozen

whole

finfish

Frozen

fillets

Frozen

fish heads

Frozen

guts/offal

Combined likelihood

of release and

exposure


Consequences of

establishment


Risk estimation Low Risk Low Risk Very Low

Risk

Very Low

Risk

Very Low

Risk

Very Low

Risk

_________________________________________________________________________________________________________________________________________________________________________________


117

www.digsfish.com

5.11 Systemic amoebic infections of goldfish

5.11.1 Aetiologic agent: Amoebae-like protozoa that infect goldfish and other finfish.


5.11.3 Australias status: Systemic amoebiasis has been reported in goldfish from QLD, NSW and Victoria

and is not a reportable disease in any State (Table 3b).

5.11.4 Epidemiology

Systemic granulomatous infections caused by enigmatic protozoa have been reported from goldfish

(Carassius auratus) in many countries (Voelker et a. 1977, Dykova et al. 1996), including Australia

(Langdon 1988). Various species of amoebae have been implicated, including Naegleria spp. and

Acanthamoeba spp. (see Wilson et al. 1995), Vannella platypodia and Rosculus ithacus (See Dykova et al.

1996, 1998), and members of the Family Hartmannellidae (see Voelker et al. 1977). Some earlier

descriptions classified the agents involved as “Dermocystidium-like” (see Kovacs-Gayer et al. 1986,

Langdon 1988, Lansberg and Paperna 1992). One of the granulomatous diseases in Australian goldfish is

due to infection by Endolimax nana, an amoeba with affiliations to the Entamoebidae (Pyecroft 2008). The

prevalence of E. nana increases with increasing water temperatures, although granulomas may be present

year round (Pyecroft 2008). Experimental infections suggest that transmission with E. nana is direct and that

infection can occur via IP injection as well as the oral route (Pyecroft 2008). The main target organ is the

kidney, though the spleen and other internal organs including the brain may also be involved in more

advanced infections (Landsberg and Paperna 1992, Dykova et al. 2001). Prevalence can approach 100% and

morbidity and mortality rates of affected fish can be high in rearing systems where fish are held at high

densities. Free living amoebae are commonly detected in aquaria, for example DeJonckheere (1979) found

100% of aquaria maintained between 28 and 37°C had high levels of amoebae, mainly the genera Naegleria

and Acanthamoeba. It is likely, therefore, that amoebic infections in goldfish are due to opportunistic

invasion of stressed hosts by free living amoebae (Dykova et al. 2001).


In Australia, as for diseases of other species of ornamental fishes (Wickins et al. 2011), amoebic infections

can occur at high prevalences in goldfish in retail stores. It is known that goldfish are sometimes used as live

bait by recreational fishers in freshwater areas (Whittington and Cullis 1988, Humphrey and Ashburner

1993, Lintermans 2004), though the quantity used is likely to be small. This suggests there is a direct

pathway for release of amoebae from goldfish into the environment via their use as bait. There is little

information published in relation to host specificity of free living amoebae capable of colonising fish,

although their ability to do so is probably related to environmental conditions that determine the number of

amoebae present in the water, the route of entry as well as the immune status of the host. There is little

information available on their resistance to physical and chemical treatments or freezing. Taking into

account this information and the relatively small quantities of goldfish used as bait, as well as the high

_________________________________________________________________________________________________________________________________________________________________________________


118

www.digsfish.com

prevalence of the disease agent, the likelihood estimations for the occurrence of amoebae in these


Release assessment for systemic amoebic infections of goldfish

Commodity type Live

goldfish

Whole fresh

dead goldfish

Likelihood of release High Moderate


Freshwater teleosts in several regions of Australia where feral populations of goldfish have established are

already at risk of exposure to amoebae carried by goldfish, but freshwater teleosts throughout the country

are also highly likely to be naturally exposed to free living amoebae in the environment. It is not known

whether other species of freshwater teleosts are susceptible to amoebic infections, but granulomatous

amoebic infections of internal organs are uncommon in native freshwater fishes, suggesting that the disease

is mainly problematic only in goldfish held at high densities. Movements of live goldfish via their use as

bait could transmit the disease agents to new regions, but the disease is already likely to be widespread

throughout the ornamental goldfish industry and these fish are sold throughout the country. Natural infection

by free living amoebae in wild fish is possible (Taylor 1977), but amoebic infections in wild fish are

extremely uncommon and directly related to poor water quality or water contamination (Taylor 1977).

Use of goldfish as live bait in Australia is illegal, but may still occur on a small scale, and the quantities of

translocated live fish are likely to be relatively small. The amoebic infections in goldfish are due to invasion

of compromised fish by opportunistic free living amoebae that are also likely to be widespread in the

environment. These disease agents are likely to be ubiquitous and become problematic only in situations

where water quality becomes degraded. Because of this, the additional risks of exposure and establishment

of amoebae from goldfish used as bait appear Negligible, the combined likelihood of release and exposure

are therefore negligible, and no further analysis is required.

_________________________________________________________________________________________________________________________________________________________________________________


119

www.digsfish.com

5.12 Infection of finfish with scuticociliates (including Uronema spp.)

5.12.1 Aetiologic agent: Scuticociliates (Protozoa: Ciliophora) that infect finfish.


5.12.3 Australias status: Scuticociliate infections have been reported in QLD, SA and Tasmania and this

disease is not a reportable disease in any State (Table 3b).

5.12.4 Epidemiology

Systemic infections by scuticociliates are problematic in the aquaculture of a wide variety of species of

marine fishes in many parts of the world (Alvarez–Pellitero et al. 2004, Kim et al. 2004, Smith et al. 2009),

including Australia (Munday et al. 1997). Under certain circumstances, these free living ciliates can infect

the gills, skin, muscle and internal organs (including the brain) and can be highly histophagous, rapidly

destroying infected tissues and causing epizootic mortalities. Several genera and species of scuticociliates

have been implicated in systemic infections of farmed fish in recent years, with the majority of important

pathogens placed within the family Philasteridae (Gao et al. 2010). The main pathogens include Miamensis

avidus (syn. Philasteroides dicentrarchi, see Parama et al. 2006) which occurs in juvenile seahorses

(Rossteuscher et al. 2008), seabass (Dicentrachus labrax) and turbot (Scophthalmus maximus) in Europe,

olive flounder (Paralichthys olivaceus) in Korea (Jung et al. 2007), and kingfish (Seriola lalandi) and groper

(Polyprion oxygeneios) fingerlings in New Zealand (Smith et al. 2009), Uronema marinum in many species

worldwide, including groper in New Zealand (Smith et al. 2009), and U. nigricans in many species

worldwide, including southern bluefin tuna (Thynnus maccoyi) in Australia (Munday et al. 1997).

Free living scuticociliates feed mainly on bacteria, but are opportunistically histophagous (Crosbie and

Munday 1999). Juvenile marine finfish in aquaculture systems appear to be particularly susceptible to

scuticociliate infection during early growout, a stage where fish are held at high densities in tanks that

usually contain nutrient enriched water containing high numbers of bacteria. The enrichment of the rearing

environment is probably due to the high availability of fish food during processes such as feeding of live

foods and weaning onto formulated pellet diets. Indeed ciliate densities in rearing tanks can reach 104

ciliates/L (Smith et al. 2009) or more, which are levels high enough to facilitate horizontal infection of fish

by ciliates that switch to a histophagous mode of nutrition (Jung et al. 2007, Song et al. 2009). Besides

environmental variables, the immune status of the host is also likely to be important (Munday et al. 1997).

Control of the disease can be difficult and depends on reducing nutrient loads in rearing tanks,

chemotherapeutic control of ciliate numbers and boosting immune performance of the fish (Munday et al.

1997, Crosbie and Munday 1999). The increasing frequency of reports of disease caused by scuticociliates

in recent years may be due to increased activity in culturing marine finfish globally, though reductions in

inshore water quality (e.g. organic enrichment) may also account for some of the observed changes, given

that this would favour the ciliates and potentially immunosuppress juvenile fish, facilitating infection by

opportunistic pathogens.

_________________________________________________________________________________________________________________________________________________________________________________


120

www.digsfish.com


Scuticociliate infections have been observed in aquacultured southern bluefin tuna in South Australia

(Munday et al. 1997), barramundi (Lates calcarifer) and barramundi cod (Chromileptes altivelis) in QLD

(B.K. Diggles and M. Landos, personal observations) and striped trumpeter in Tasmania. These ciliates

occur naturally in the environment and can infect a wide range of fish species that occur in marine waters

around the country. The prevalence and intensity of natural scuticociliate infections in wild fish in Australia

are unknown, but are likely to be very low, especially in comparison to those of diseased cultured fish.

Different species or strains of scuticociliates may exist in different parts of the country, although their

distribution is not known at this time. Scuticociliates may occur in fish used as live bait by recreational

fishers, though because live finfish are not commercially available as bait (except ornamental species sold

from pet shops), the quantity of live bait that is translocated is likely to be relatively low. The majority of the

volume of bait and berley translocated throughout the country is frozen, including frozen whole fish and

processed fish products (e.g. heads or guts). Cryopreservation of scuticociliates is possible by freezing them

in specialized cryopreservation media, but conventional freezing without cryopreservatives deactivates

ciliates by cell rupture due to ice formation (Anderson et al. 2009). There is little information published in

relation to host specificity of scuticociliates capable of colonising fish, although their ability to do so is

probably related to environmental conditions that determine the number of ciliates present in the water, the

route of entry as well as the immune status of the host.

Taking into account that juvenile aquacultured marine fish are generally not permitted to be used as live bait

or fresh unfrozen bait or berley in most/all jurisdictions in Australia, and the likely low prevalence of the

disease agent in wild fish, the likelihood estimations for the occurrence of scuticociliates in these


Release assessment for infection of finfish with scuticociliates

Commodity

type

Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release

Low Low Negligible Negligible Negligible Negligible


Marine and freshwater teleosts throughout Australia are already at risk of natural exposure to free living

scuticociliates in the environment. They are facultative parasites of compromised fish and scuticociliate

disease is problematic only in circumstances where large numbers of juvenile fish are held at high densities

in nutrient enriched water typical of aquaculture rearing environments. Movements of live and whole fresh

dead fishes via their use as bait could theoretically transmit these disease agents to new regions, but the

agents are already likely to be free living and widespread throughout the country. Also, infection and

_________________________________________________________________________________________________________________________________________________________________________________


121

www.digsfish.com

establishment would occur only if sufficient quantities of scuticociliates (i.e. an infective dose) were

introduced into an area where susceptible hosts were present.

The quantities of scuticociliates required to successfully transmit infection horizontally via the immersion

route have been determined for some host/parasite combinations. Jung et al. (2007) found that Miamiensis

avidus successfully invaded olive juvenile flounder (mean length: 14.9 cm; mean weight: 26.8 g) after bath

exposure to 2 × 103ciliates/ml , resulting in 85% mortality after 8 to 10 days. Higher doses (2 × 104 and 2 ×

105 ciliates/ml) caused 100% mortality (Jung et al. 2007). Song et al. (2009) found that juvenile olive

flounder exposed by bath immersion to several strains of M. avidus had mortalities ranging between 60 to

100% when subjected to 3-4 × 103 ciliates/ml in fish ranging between 8 and 40 g. Fish infected by

immersion had many M. avidus in gills, fins, skin muscle, brain and intestine accompanied by necrosis and

haemorrhages, fish exposed to Uronema marinum by both IP injection and bath immersion showed less than

30% mortality, with no ciliate invasion in the skin, gills or brain (Song et al. 2009).

Scuticociliates are free living opportunistic pathogens that are likely to be widespread in the natural

environment. Infections of wild fishes with scuticociliates are uncommon, due to the lack of eutrophic

conditions in the wild, and if infections occur in wild fish they are likely to be very low intensity. High

intensity infections that could promote horizontal transmission of disease via use of bait and berley are only

observed in juvenile aquacultured fishes held at high densities in degraded water quality. The ciliates do not

tolerate freezing, and live and fresh dead juvenile aquacultured fishes are generally not permitted to be used

as bait or berley in most/all jurisdictions in Australia. Because they are not commercially available, the

quantities of these commodities that are translocated are therefore likely to be negligible. Because of this,

the additional risks of exposure and establishment of scuticociliates into new fish populations appear to be

Negligible, the combined likelihood of release and exposure are therefore negligible, and no further analysis

is required at this time. However, this risk assessment would need to be revised if industries based on

distribution of juvenile finfish grown in aquaculture facilities specifically for live bait markets are permitted

to be developed in the future.

_________________________________________________________________________________________________________________________________________________________________________________


122

www.digsfish.com

5.13 Infection of finfish with monogeneans

5.13.1 Aetiologic agent: Infections with Benedenia spp, Gyrodactylus spp, Dactylogyrus spp. and other

monogenean ectoparasites (Phylum Platyhelminthes, Subclass Monogenea) of marine and freshwater fishes.

5.13.2 OIE List: Yes (Gyrodactylus salaris only) NACA List: Yes (Gyrodactylus salaris only)

5.13.3 Australias status: Monogeneans have been reported from freshwater and marine finfish in all States,

and infection by Gyrodactylus salaris is a reportable disease in all States (Table 3b).

5.13.4 Epidemiology

Monogeneans are parasitic flatworms which have been recorded from a wide range of marine and freshwater

fish species throughout Australia. Though a number of species of Gyrodactylus have been recorded from

Australia (Dove and Ernst 1998, Ernst et al. 2000), Gyrodactylus salaris has not been recorded from

Australian salmonids (Herfort 2004), and this species will not be considered further in this risk assessment.

Monogeneans have direct lifecycles with horizontal transmission of infective oncomiracidium larvae that

hatch from eggs deposited by adult worms living on the external surfaces, gills, mouth and nares of teleosts

and elasmobranchs (Kearn 1986). The combination of the single host lifecycle with direct transmission of

infective stages, together with the high fecundity of adult worms, means that hyperinfections can quickly

result when infected fish are held in confinement, often resulting in morbidity and mortalities due to

osmoregulatory dysfunction, anaemia, and secondary bacterial infection (Thoney and Hargis 1991).

The majority of monogeneans have high host specificity (Whittington et al. 2000), and many are site specific

within particular microhabitats on one host (Ernst and Whittington 2001). These highly specialized

characteristics tend to greatly reduce the risk of transfer of these parasites to new hosts when they are

translocated into new regions. However, host specificity may not be strict between closely related species of

hosts, which may be able to share limited numbers of parasites when held at close quarters in confinement

(Ernst and Whittington 2001). There are also a few species of monogeneans that have very low host

specificity and infect a broad range of unrelated hosts (Whittington and Horton 1996, Blazek et al. 2008).


Infections of monogeneans are common on wild fish throughout Australia, and while prevalences of

infection may be high in some circumstances, the intensity of natural monogenean infections in wild fish

tend to be very low, especially in comparison to those of diseased cultured fish. Different species of

monogeneans exist on various hosts in different parts of the country (e.g. Byrnes 1986, Hayward 1997),

although the identity and distribution of many species of monogeneans is not known at this time.

Monogeneans are, therefore, highly likely to occur on teleosts used as bait. The majority of the volume of

bait and berley translocated throughout the country is frozen, including frozen whole fish and processed fish

products (e.g. heads or guts). However, freezing quickly deactivates adult monogeneans (Jones and Gibson

1997), while their eggs are also sensitive to desiccation (Ernst et al. 2005, Chen et al. 2010) and for marine

species, low salinity water typically encountered by fish products held on ice also inactivates eggs (Ernst et

_________________________________________________________________________________________________________________________________________________________________________________


123

www.digsfish.com

al. 2005, Chen et al. 2010). It is not known whether monogenean eggs are resistant to freezing, but this is

considered highly unlikely due to ice formation inside the cells of the oncomiracidium. Monogeneans do not

survive for any length of time upon the death of their host (BK Diggles, personal observations), hence the

presence of viable monogeneans on fresh dead finfish is very unlikely, though monogenean eggs may be able

to survive low temperatures typical of those used to increase the shelf life of chilled fish (c. 4°C) for short

periods. The main route for translocation of viable juvenile and adult monogeneans therefore appears to be

via live finfish. Live baits are mainly used by recreational fishers, though because live finfish are not

commercially available as bait, they are usually caught at the fishing site and the quantity of live bait that is

translocated is likely to be relatively low. Because of this, and taking into account the high prevalence of

these disease agents, the likelihood estimations for the occurrence of viable monogeneans eggs or worms in

these commodities are listed below.

Release assessment for infection of finfish with monogeneans

Commodity

type

Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release

High Very low Negligible Negligible Negligible Negligible


Marine and freshwater teleosts throughout Australia are already at risk of exposure to monogenean parasites.

They are natural parasites of wild fishes and cause disease only in circumstances where their hosts are

confined in captivity at the high densities typical of aquaculture and ornamental rearing environments.

Movements of live and whole fresh dead fishes via their use as bait could theoretically transmit these disease

agents to new regions, but the agents mostly have very high host specificity and are unlikely to spread to a

wide range of hosts. Infection and establishment would occur only if live monogenean parasites or viable

eggs were introduced into an area where susceptible hosts were present. However, monogeneans are

hermaphroditic and infections can become established if susceptible hosts are exposed even to only one

viable larvae from one egg (BK Diggles, personal observations), hence the translocation of some types of

fish products with viable monogeneans worms or eggs remains a route of introduction and infection of fish in

new geographic areas.

Because live baitfish are not commercially available (except ornamental species sold from pet shops), fish

used for live bait are usually caught by recreational fishers at or near the fishing location, hence the

quantities of these commodities that are translocated any large distances are likely to be low. Only small

quantities of fresh unfrozen fish are generally available from commercial fishing co-operatives for local use

as bait, and the vast majority of bait and berley that is translocated throughout the country is frozen whole or

processed marine fish, in which monogeneans will be unviable. However, monogenean infections can

become established even if susceptible hosts are exposed to only one viable larvae from one egg (BK

Diggles, personal observations), hence the translocation of fish products with viable monogenean worms or

eggs remains a potential route of introduction and infection of fish in new geographic areas, even if small

_________________________________________________________________________________________________________________________________________________________________________________


124

www.digsfish.com

volumes of fish are translocated. Because of this, the additional risk of exposure and establishment is non-

negligible, and the likelihood of exposure and establishment of monogeneans in new fish populations is



Wild fish of all size classes can become infected with monogeneans, however infections of wild fish do not

usually kill them or reduce the market value of affected fishes. However, some monogeneans have the

potential to cause significant damage to confined fish in aquaculture and aquaria, and while there are many

examples of successful treatment of monogeneans, their treatment does add significantly to production costs

of affected aquaculture industries (Ernst et al. 2002). While the full range of monogenean parasites and their

susceptible hosts in Australia remain to be determined, these parasites have already been recorded from a

wide range of teleosts in both freshwater and marine environments. Besides the exotic G. salaris, no

monogenean disease agents are listed by the OIE or NACA and none are reportable diseases in any State

(Table 3b). Hence the spread of these disease agents is unlikely to have adverse impacts on trade.

Considering all of these factors, establishment of monogeneans in new areas would have mild to moderate

biological consequences, which would be amenable to control, and could cause some unwanted

environmental effects. It is therefore estimated that the consequences of introduction of monogeneans into

different parts of the Australian environment via use of infected bait would likely be Low.


The unrestricted risk associated with monogeneans is determined by combining the likelihood of entry and


monogeneans does not exceed the ALOP for any of the commodity types, suggesting that additional risk

management for these disease agents is not required at this time.

Risk estimate for infection of finfish with monogeneans

Commodity type Live

finfish

Whole

fresh dead

finfish

Frozen

whole

finfish

Frozen

fillets

Frozen

fish heads

Frozen

guts/offal

Combined


and exposure

Low Very low Negligible Negligible Negligible Negligible

Consequences of

establishment


Risk estimation Very

low risk

Negligible

risk

Negligible

risk

Negligible

risk

Negligible

risk

Negligible

risk

_________________________________________________________________________________________________________________________________________________________________________________


125

www.digsfish.com

5.14 Infection of finfish and molluscs with digeneans

5.14.1 Aetiologic agent: Digenean trematodes (Phylum Platyhelminthes, Subclass Digenea) (including

Centrocestus formosanus) that utilise freshwater molluscs as the first intermediate host and fish as the second

intermediate hosts.


5.14.3 Australias status: Centrocestus formosanus has been reported in QLD (Evans and Lester 2001,

Dove 2000). These disease agents are not reportable in any State (Table 3b).

5.14.4 Epidemiology

Digenean trematodes are endoparasitic helminths which have been recorded from a wide range of marine and

freshwater fish species throughout Australia. Their indirect lifecycle requires a molluscan first intermediate

host with plankton eating fishes as final hosts, or second intermediate hosts in some lifecycles where final

hosts include larger fishes, birds and mammals. Under most circumstances, the multi host lifecycles of these

parasites reduce the risk of their translocation, because additional hosts need to occur in the receiving

environment in order to complete the life cycle. However, invasive digenean parasites have low host

specificity, because low host specificity is a prerequisite for invasiveness (Bauer 1991). Centrocestus

formosanus is a digenetic trematode with a freshwater snail first intermediate host, various species of fish as

second intermediate hosts, and piscivorous birds and mammals as the final host (Mitchell et al. 2005). The

worm encysts as metacercaria in the gills of a wide range of freshwater fishes, including species used as bait

in Australia such as goldfish, European carp, atherinids, elotrids, mullet, poeciliids and cichlids (Scholz and

Salgado-Maldonado 2000, Evans and Lester 2001, Mitchell et al. 2005). Centrocestus formosanus originates

from Taiwan, but has been spread worldwide with the movements of snail intermediate hosts, bird definitive

hosts and/or ornamental fishes (Scholz and Salgado-Maldonado 2000, Evans and Lester 2001, Font 2003,

Mitchell et al. 2005). This species has been detected at a prevalence of 100% in poeciliids sampled from

ornamental finfish wholesalers in Australia (Evans and Lester 2001), and also at high prevalences in wild

populations of poeciliids (Dove 1998, Dove 2000). This parasite is virtually non host specific for fish second

intermediate hosts, but is more specific for the first intermediate host, which is usually the snail Melanoides

tuberculata (see Vogelbein and Overstreet 1988, Scholz and Salgado-Maldonado 2000, Mitchell et al. 2005).

Melanoides tuberculata, a member of the family Thiaridae, has been confirmed as being introduced into

tropical Australia, and there are number of species of native Thiarid snails in Australia which could also act

as intermediate hosts for this parasite, which uses water birds (herons and egrets) and mammals (including

mice, rats, cats, rabbits and even humans) as the final host (Mitchell et al. 2005).

Another notable trematode in freshwater fishes in Australia is Clinostomum complanatum, a digenetic

trematode with fish second intermediate hosts and bird final hosts. This worm encysts as large yellow

metacercaria in the fillet muscle of the fish, reducing their marketability (Matthews and Cribb 1998) and

potentially causing zoonotic disease if a human consumes undercooked infected fish (Aohagi et al. 1992). In

Australia, Clinostomum spp. have been recorded from several teleosts used as bait, including members of the

Families Ambassidae, Eleotridae, Melanotaeniidae, and Teraponidae (Appendix 1).

_________________________________________________________________________________________________________________________________________________________________________________


126

www.digsfish.com


Digeneans have been recorded from a wide range of marine and freshwater fish species throughout Australia,

however it is likely that only a fraction of the total number of native species that exist in this country have

been identified to date (Bott et al. 2005). Different species of digeneans exist on various hosts in different

parts of the country (e.g. Bray and Cribb 1998), and the identity and distribution of many species of

digeneans is not known at this time. Digeneans are, therefore, highly likely to occur in teleosts and molluscs

used as bait. For introduced species such as C. formosanus, which has likely been introduced via ornamental

finfish (Evans and Lester 2001), the parasite has been recorded in populations of poeciliids that have

established in the wild (Dove 2000). It is likely, therefore, that C. formosanus would be present in poeciliids

and perhaps other species of freshwater fishes used as bait or berley. Freshwater finfish are not generally

sold as live or frozen bait in retail outlets (except ornamental species sold from pet shops), and the majority

of the volume of bait and berley translocated throughout the country is frozen, including frozen whole fish

and processed fish products (e.g. heads or guts). However, freezing quickly deactivates adult digeneans as

well as metacercariae (Jones and Gibson 1997), disrupting the lifecycle. Digeneans do not survive for very

long after the death of their host at normal environmental temperatures (BK Diggles, personal observations),

hence the presence of viable digeneans in fresh dead finfish is unlikely, though their viability may be

extended for short periods by the low temperatures typical of those used to increase the shelf life of chilled

fish (c. 4°C). The main route for translocation of viable digeneans through use of bait and berley therefore

appears to be via live finfish or their molluscan first intermediate hosts. Live baits are mainly used by

recreational fishers, though because live finfish are not commercially available as bait, the quantity of live

bait that is translocated long distances is likely to be low. Similarly, live molluscs are generally not available

commercially, but are widely available frozen. Because of this, and taking into account the high prevalence

of these disease agents, the likelihood estimations for the occurrence of viable digeneans in these


Release assessment for infection of finfish and molluscs with digeneans

Commodity

type

Live

finfish

and

molluscs

Whole fresh

dead finfish

and molluscs

Frozen

whole finfish

and molluscs

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release



Marine and freshwater teleosts throughout Australia are already at risk of exposure to digenean parasites.

They are natural parasites of wild fishes and only certain species of digeneans cause disease, usually in

circumstances where they are introduced into new hosts (Mitchell et al. 2005). Movements of live and

_________________________________________________________________________________________________________________________________________________________________________________


127

www.digsfish.com

whole fresh dead fishes and molluscs via their use as bait could theoretically transmit these disease agents to

new regions, but even digeneans with low host specificity such as C. formosanus can be specific for at least

one of the hosts in their lifecycle, reducing their chances of exposure and establishment. Indeed, infection

and establishment via movements of infected fish or molluscs would occur only if live digeneans or their

infective stages were introduced into an area where all susceptible intermediate and final hosts were present.

When assessing the risk of establishment in parasites with complex life cycles, the movements of other hosts

need to be considered. For C. formosanus, its spread to many parts of the world has been related more to the

spread of snail intermediate hosts (e.g. Melanoides tuberculata), as it is most host specific for the first

intermediate host (Scholz and Salgado-Maldonado 2000, Mitchell et al. 2005) than it is for the second

intermediate or final hosts. However, since M. tuberculata has been introduced into the Northern Territory

and possibly other parts of northern Australia, and other native Thiarid snails may also act as suitable

intermediate hosts, C. formosanus is more likely to be able to establish in at least some regions of Australia

via translocation of second intermediate hosts in bait and berley, or even through migrations of its bird and

mammalian definitive hosts.

Because live baitfish are not commercially available (except from ornamental species sold from pet shops),

fishes used for live bait are usually caught by recreational fishers at or near the fishing location, hence the

quantities of these commodities that are translocated long distances are likely to be low. Only small

quantities of fresh unfrozen fish are generally available from commercial fishing co-operatives for local use

as bait, and the vast majority of bait and berley that is translocated throughout the country is frozen whole or

processed marine fish, in which digeneans will be unviable. It must also be recognized that movements of

piscivorous bird definitive hosts may be a faster, more effective and uncontrolled method of translocation of

digenean parasites in regions where other hosts have already been introduced and/or are already endemic.

Because of this, the additional risk of exposure and establishment of digeneans via use of bait is reduced, but

remains non-negligible, and the likelihood of exposure and establishment of digeneans in new fish or

mollusc populations is considered to be Low.


Wild fish of all size classes can become infected with digeneans, and establishment of some species, such as

C. formosanus can have negative impacts on the health of individual fish and their populations. Indeed, C.

formosanus is of considerable veterinary importance as a threat to native fish populations due to its ability to

establish heavy infections (up to 6000 metacercariae per fish) in a wide range of fishes from different

families (Scholz and Salgado-Maldonado 2000), which in Australia probably could include several species

of threatened native fishes. The potential threat to human health through consumption of undercooked fish

containing metacercariae of C. formosanus should also not be underestimated (Scholz and Salgado-

Maldonado 2000). Establishment of other species, such as C. complanatum, could also reduce the market

value of affected fishes through location of grossly visible metacercariae in the fillet. However, only a very

few digeneans have the potential to cause damage to confined fish in aquaculture and aquaria, and for these

species disruption of the lifecycle in the rearing system by removal of the molluscan first intermediate host is

usually an effective method of control which does not necessarily add significantly to production costs of

affected aquaculture industries (Mitchell et al. 2005). No digenean disease agents are listed by the OIE or

NACA and none are reportable diseases in any State (Table 3b). Hence the spread of these disease agents is

_________________________________________________________________________________________________________________________________________________________________________________


128

www.digsfish.com

unlikely to have adverse impacts on trade. Considering all of these factors, establishment of the majority of

native digeneans in new areas would have mild or no biological consequences, however spread of invasive

introduced species such as C. formosanus would likely incur moderate or even significant biological

consequences, which would not be amenable to control in wild fish populations, and would cause some

unwanted environmental effects. It is therefore estimated that the consequences of introduction of native

digeneans into different parts of the Australian environment via use of infected bait would likely be Low, but

for Centrocestus formosanus, the consequences would be Moderate.


The unrestricted risk associated with digeneans is determined by combining the likelihood of entry and


digeneans does not exceed the ALOP for any of the commodity types, except for Centrocestus formosanus,

which presents a low risk from movements of live freshwater finfish and molluscs, suggesting that additional

risk management is required for this disease agent in these commodities.

Risk estimate for infection of finfish and molluscs with digeneans

Commodity type Live

finfish

and

molluscs

Whole

fresh dead

finfish and

molluscs

Frozen

whole

finfish and

molluscs

Frozen

fillets

Frozen

fish heads

Frozen

guts/offal

Combined


and exposure

Low Very Low Negligible Negligible Negligible Negligible

Consequences of

establishment


Risk estimation Low risk Negligible

risk

Negligible

risk

Negligible

risk

Negligible

risk

Negligible

risk

_________________________________________________________________________________________________________________________________________________________________________________


129

www.digsfish.com

5.15 Infection of finfish and crustaceans with nematodes and cestodes

5.15.1 Aetiologic agent: Cestode and nematode helminths that utilise copepods as their first intermediate

host, and fish as second intermediate hosts, including Bothriocephalus acheilognathi, Ligula intestinalis and

Camallanus cotti.


5.15.3 Australias status: Reported in QLD, NSW, ACT, Victoria and WA, and these diseases are not

reportable in any State (Table 3b).

5.15.4 Epidemiology

Nematodes and cestodes are endoparasitic helminths that live in the gastrointestinal tract of fishes and other

vertebrates. Their lifecycle generally requires crustaceans as the first intermediate host with plankton eating

fishes as final hosts, or second intermediate hosts in some lifecycles where final hosts include larger fishes,

sharks, birds or mammals (Rohde 1984, Noga 1996). Under most circumstances, the multi host lifecycles of

these parasites reduce the risk of their translocation, because additional hosts need to occur in the receiving

environment in order to complete the life cycle. However, invasive nematode and cestode parasites have low

host specificity, because low host specificity is a pre-requisite for invasiveness (Bauer 1991). In Australia,

the introduced European carp (Cyprinus carpio) are a common host for the Asian fish tapeworm

Bothriocephalus acheilognathi, a cestode with low host specificity that has been introduced to many parts of

the world from its original distribution in Taiwan through movements of cyprinid and poeciliid fishes (Bauer

1991, Kennedy 1993, Font 2003). Although B. acheilognathi has a complex life cycle, its intermediate hosts

are cosmopolitan copepods which are an important food source for plankton-eating juvenile fishes (Bauer

1991). In eastern Australia B. acheilognathi has been reported in wild populations of the European carp,

Gambusia spp. and several native fish species, particularly elotrids (Hypseleotris spp.) (Dove et al. 1997,

Dove and Fletcher 2000), as well as in poeciliids at ornamental fish wholesalers (Evans and Lester 2001). In

Western Australia, another invasive cestode Ligula intestinalis has been reported in wild populations of both

native and introduced fishes, with prevalence of infection significantly higher in native fishes (Lymbery et al.

2010). Ligula intestinalis also requires ubiquitous copepods as the first intermediate host, and has low host

specificity for the second intermediate host (planktivorous fishes), with its final host being piscivorous birds

(Hassan 2008). Native fishes (including threatened species such as galaxiids) infected by L. intestinalis

plerocercoids have grossly distended abdomens, markedly reduced gonad mass and are weak and slow

moving, making them easy targets for predators (Chapman et al. 2006, Lymbery et al. 2010).

The Asian nematode Camallanus cotti is another non host specific endoparasitic helminth that has spread

from southeast Asia to many parts of the world with the trade in ornamental fishes (see Levsen and Berland

2002, Levsen and Jakobsen 2002). Examination of guppies imported into Korea showed 14.4% prevalence

(Kim et al. 2002a), while C. cotti has been detected at a prevalence of 48% in poeciliids sampled from

ornamental finfish wholesalers in Australia (Evans and Lester 2001). Camallanus cotti caused 30%

mortalities following its introduction into an ornamental fish farm in Korea, where it infected 71% of the

cultured fishes (Kim et al. 2002b). Camallanus cotti normally uses planktonic copepods as intermediate

_________________________________________________________________________________________________________________________________________________________________________________


130

www.digsfish.com

hosts, but if they are not present, it can infect directly, fish-to-fish (Levsen and Jakobsen 2002). After

guppies were introduced into Hawaii for mosquito control, C. cotti switched hosts into 5 native fish species,

including an eleotrid (Eleotris sandwicensis) (see Font and Tate 1994, Font 1998), and members of the

Family Elotrididae are used as bait or berley in Australia. Langdon (1988) reported C. cotti as being present

and causing disease in captive populations of poeciliids, but to date it remains unclear whether this parasite

has become established in wild populations of poeciliids or other freshwater fishes in Australia.


Nematodes and cestodes occur in a wide range of marine and freshwater fish and crustacean species

throughout Australia. Different species of cestodes and nematodes exist in various hosts in different parts of

the country, and the identity and distribution of many species is not known at this time. Nematodes and

cestodes are, therefore, highly likely to occur in teleosts and crustaceans used as bait. Native and introduced

species of freshwater finfish in several regions of Australia are also known to be hosts for introduced

invasive nematode and cestode parasites, particularly B. acheilognathi, L. intestinalis and C. cotti. Fishes

used as bait or berley that are known to be infected by these disease agents include goldfish, European carp,

elotrids, poeciliids, cichlids, retropinnids and galaxiids (Table 8), while other native fishes are also likely to

be susceptible. It is likely, therefore, that these invasive parasites would be present in several species of

freshwater fishes used as bait or berley.

Freshwater finfish are not generally sold as live bait (except ornamental species sold from pet shops, though

this is not intended for use as bait), or as frozen bait in retail outlets. The largest quantity of bait and berley

translocated throughout the country is frozen, including frozen whole and processed fish and crustacean

products. However, prolonged freezing at -20°C for more than 72 hours kills all larval and adult stages of

cestodes and nematodes (Jones and Gibson 1997). Nematodes and cestodes do not survive for very long

after the death of their host at normal environmental temperatures (B.K. Diggles, personal observations),

hence the presence of viable nematodes and cestodes in fresh dead finfish is unlikely, though their viability

may be extended for short periods by the low temperatures used to increase the shelf life of chilled fish (c.

4°C). The main route for translocation of viable cestodes and nematodes through use of bait and berley

therefore appears to be via live finfish or their crustacean first intermediate hosts. Live baits are mainly used

by recreational fishers, though because live finfish are not commercially available as bait, the quantity of live

bait that is translocated long distances is likely to be low. Similarly, live crustaceans are generally not

available commercially, but 11% of recreational fishers catch their own prawns (Kewagama Research 2007),

and large quantities of crustaceans are widely available frozen. Because of this, and taking into account the

high prevalence of these disease agents, the likelihood estimations for the occurrence of viable cestodes and

nematodes in these commodities are listed below.

_________________________________________________________________________________________________________________________________________________________________________________


131

www.digsfish.com

Release assessment for infection of finfish and crustaceans with nematodes and cestodes

Commodity

type

Live

finfish and

crustaceans

Whole fresh

dead finfish

and

crustaceans

Frozen

whole finfish

and

crustaceans

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release



Marine and freshwater teleosts throughout Australia are already at risk of exposure to nematode and cestode

parasites. They are natural parasites of wild fishes and only certain species of nematodes and cestodes cause

disease, usually in circumstances where they are introduced into new hosts. Movements of live and whole

fresh dead fishes and crustaceans via their use as bait could theoretically transmit these disease agents to new

regions, especially for invasive nematodes and cestodes with low host specificity such as B. acheilognathi,

and L. intestinalis with ubiquitous copepod first intermediate hosts. Infection and establishment of B.

acheilognathi and L. intestinalis via movements of infected fish or crustaceans would occur only if live

nematodes or cestodes or their infective stages were introduced into an area where the copepod intermediate

host and bird final hosts were present. However for C. cotti, copepod intermediate hosts are not necessary

and it can transmit directly from fish to fish (Levsen and Jakobsen 2002).

When assessing the risk of establishment in parasites with complex life cycles, the movements of other hosts

need to be considered. For B. acheilognathi, L. intestinalis, their copepod first intermediate hosts are

ubiquitous, and because of this these parasites have been spread to many parts of the world via movements of

fishes or natural movements of their avian final hosts. And as mentioned previously, C. cotti does not

necessarily require use of copepod intermediate hosts for successful transmission, although the fitness of the

parasite is increased if the intermediate host is used (Levsen and Jakobsen 2002). This suggests that B.

acheilognathi, L. intestinalis and C. cotti are likely to be able to establish in some regions of Australia via

translocation of their fish second intermediate hosts in bait and berley, or even with translocation of some

crustaceans used as bait, as well as through migrations of their bird definitive hosts in the case of B.

acheilognathi and L. intestinalis.

Because live baitfish and crustaceans are generally not commercially available as bait (and luse of live bait in

freshwater areas is prohibited in some jurisdictions), fishes and crustaceans used for live bait are usually

caught by recreational fishers at or near the fishing location, hence the quantities of these commodities that

are translocated long distances are likely to be low. Only small quantities of fresh unfrozen fish are

generally available from commercial fishing co-operatives for local use as bait, and the vast majority of bait

and berley that is translocated throughout the country is frozen whole or processed marine fish, in which

nematodes and cestodes will be unviable. It must also be recognized that movements of piscivorous bird

_________________________________________________________________________________________________________________________________________________________________________________


132

www.digsfish.com

definitive hosts may be a faster, more effective and uncontrolled method of translocation of some nematode

and cestode parasites in regions where other hosts have already been introduced and/or are already endemic.

Because of this, the additional risk of exposure and establishment of nematodes and cestodes via use of bait

or burley is reduced, but remains non-negligible, and the likelihood of exposure and establishment of

nematodes and cestodes in new fish populations via translocation is considered to be Low.


All size classes of fish can become infected with nematode and cestode parasites, but planktivorous juvenile

fish are particularly susceptible to infection. Establishment of some species, such as B. acheilognathi, L.

intestinalis and C. cotti can have negative impacts on the health of individual fish and their populations,

including populations of threatened native species (Dove et al. 1997, Dove and Fletcher 2000, Chapman et

al. 2006, Lymbery et al. 2010). Establishment of species such as L. intestinalis which have large, grossly

visible plerocercoids, can also reduce the market value of affected fishes through location of grossly visible

metacercariae in the fillet. However, only a very few nematodes and cestodes have the potential to cause

damage to confined fish in aquaculture and aquaria, and for these species disruption of the lifecycle in the

rearing system by removal of the crustacean first intermediate host is a potential method of control. No

nematode or cestode disease agents are listed by the OIE or NACA and none are reportable diseases in any

State (Table 3b). Hence the spread of these disease agents is unlikely to have adverse impacts on trade.

Considering all of these factors, establishment of the majority of native nematodes and cestodes in new areas

would have mild or no biological consequences, however spread of invasive introduced species such as B.

acheilognathi, L. intestinalis and C. cotti would likely incur moderate or even significant biological

consequences, which would not be amenable to control in wild fish populations, and would cause some

unwanted environmental effects. It is therefore estimated that the consequences of introduction of native

nematodes and cestodes into different parts of the Australian environment via use of infected bait would

likely be Low, but for Bothriocephalus acheilognathi, Ligula intestinalis and Camallanus cotti, the

consequences would be Moderate.


The unrestricted risk associated with nematodes and cestodes is determined by combining the likelihood of


estimate for native nematodes and cestodes does not exceed the ALOP for any of the commodity types,

except for introduced species such as Bothriocephalus acheilognathi, Ligula intestinalis and Camallanus

cotti, which present a low risk from movements of live freshwater finfish, suggesting that additional risk

management is required for these disease agents in these commodities.

_________________________________________________________________________________________________________________________________________________________________________________


133

www.digsfish.com

Risk estimate for infection of finfish and crustaceans with nematodes and cestodes

Commodity type Live

finfish and

crustaceans

Whole

fresh dead

finfish and

crustaceans

Frozen

whole

finfish and

crustaceans

Frozen

fillets

Frozen

fish heads

Frozen

guts/offal

Combined

likelihood of

release and

exposure


Consequences of

establishment



risk

Negligible

risk

Negligible

risk

Negligible

risk

Negligible

risk

_________________________________________________________________________________________________________________________________________________________________________________


134

www.digsfish.com

5.16 Infection of finfish with copepods

5.16.1 Aetiologic agent: Ectoparasitic copepods (Family Caligidae, Family Lernaeidae) that infect fishes,

including Caligus epidemicus / other sea lice, and Lernaea cyprinacea / Lernaea spp.


5.16.3 Australias status: Copepod infections have been reported from finfish in all States, and sea lice

infections are reportable in Tasmania (Table 3b).

5.16.4 Epidemiology

Parasitic copepods live on the body surfaces, gills and in the musculature of marine and freshwater fishes.

Their lifecycles are direct with fish being infected by planktonic copepodid larval stages that hatch from eggs

deposited by adult copepods (Kabata 1984, Hemaprasanth et al. 2011). In Australia, feral goldfish and

European carp are common hosts for Lernaea cyprinacea (see Langdon 1988, Rowland and Ingram 1991), a

species that has been spread to many areas throughout the world with movements of goldfish (Bauer 1991,

Kennedy 1993). Lernaea cyprinacea is a member of the Family Lernaeidae (anchor worms), a group of

copepods highly adapted to a parasitic lifestyle in which adult parasites develop an attachment organ that

lodges deep in the musculature of the fillet of a wide range of fishes, causing considerable damage to the

muscle and internal organs, significant morbidity and even mortalities (Bond 2004). In eastern Australia L.

cyprinacea has been reported in wild populations of goldfish and European carp, redfin perch, and also

several native fish species including Murray cod (Maccullochella peelii), golden perch (Macquaria

ambigua), silver perch (Bidyanus bidyanus), Australian smelt (Retropinna semoni), and the endangered trout

cod (Maccullochella macquariensis) (Ashburner 1978, Langdon 1988, Rowland and Ingham 1991, Bond

2004). The endangered Australian grayling (Prototroctes maraena) and vulnerable mountain galaxias

(Galaxias olidus) were also infected by L. cyprinacea (see Hall 1983, Bond 2004). In Western Australia, L.

cyprinacea was recorded from four native fish species (Galaxias occidentalis Edelia vittata, Bostockia

porosa, and Tandanus bostocki) and three introduced fish species (goldfish, Gambusia holbrooki and

Phalloceros caudimaculatus) at two localities in the Canning River, in the south west of Western Australia

(Hassan 2008, Hassan et al. 2008, Lymbery et al. 2010). The likely source of introduction of the parasite

was via introduced cyprinids, particularly goldfish and European carp (Hassan et al. 2008).

A large number of marine fishes harbour copepod ectoparasites from the Family Caligidae (sea lice). These

copepods encounter their host as copepodids then attach to the host fish via the specialized chalimus larvae

(Kabata 1984), which is sedentary until such time as the copepod moults to the pre adult and adult stages,

which are mobile and can be found attached to gills, skin or fins (MacKenzie et al. 1998). High numbers of

chalimus larvae can cause pathological changes at their attachment sites (Roubal 1994, MacKenzie et al.

1998), while high numbers of pre adult and adult caligids (particularly members of the genera

Lepeophthirius and Caligus) have been responsible for disease and significant mortalities in the culture of

salmonids in several overseas countries (Pike and Wadsworth 1999). In cases where cultured fish become

heavily infected, they become stressed, and death commonly occurs, ultimately due to osmoregulatory

failure or secondary bacterial infection (MacKenzie et al. 1998, Pike and Wadsworth 1999).

_________________________________________________________________________________________________________________________________________________________________________________


135

www.digsfish.com


Parasitic copepods occur on a wide range of marine and freshwater fishes throughout Australia. Different

species of copepods exist on various hosts in different parts of the country, and the identity and distribution

of many species is not known at this time. Caligids have been reported from a very wide range of host

species, some of which are used as bait or berley (Appendix 1), but in most circumstances the species

involved have not been associated with disease. However one species, Caligus epidemicus, has caused

mortality in wild fishes (Hewitt 1971), and it infects a broad range of hosts in Australia, including several

species of bream (Acanthopagrus spp.), at least 4 species of mullet, and perchlets (Ambassis marianus) (see

Hewitt 1971, Byrnes 1987, Roubal 1994, Hallett and Roubal 1995) all of which are sometimes used as bait

(Table 8). Other species used as bait or berley which are susceptible to C. epidemicus include tilapia

(Oreochromis mossambicus) and theraponids (Lin et al. 1996). Lernaea cyprinacea has also been recorded

from a wide range of freshwater fishes that are used as bait and/or berley, including Carassius auratus,

Cyprinus carpio, Perca fluviatilis and members of the Families Poeciliidae, Retropinnidae, Teraponidae, and

Galaxiidae (Ashburner 1978, Langdon 1988, Rowland and Ingham 1991, Bond 2004, Hassan et al. 2008),

while several other groups of freshwater fishes are also likely to be susceptible to infection.

Copepods are, therefore, highly likely to occur in freshwater and marine teleosts used as bait. Finfish are not

generally sold as live bait in retail outlets, but they are sold fresh and the largest quantity of bait and berley

translocated throughout the country is frozen, including frozen whole and processed fish products. However,

freezing quickly deactivates copepods and their infective stages (Jones and Gibson 1997). Some copepods

can survive on their host for short periods of time after its death, provided the host is kept in the water, but in

all other circumstances death of copepods occurs soon after their hosts are removed from the water (BK

Diggles, personal observations), hence the presence of viable copepods in fresh dead finfish is unlikely. The

main route for translocation of viable copepods through use of bait and berley therefore appears to be via live

finfish. Live baits are mainly used by recreational fishers, though because live finfish are not commercially

available (except ornamental species sold from pet shops, which are not intended for use as bait), the

quantity of live bait that is translocated long distances is likely to be low. Because of this, and taking into

account the high prevalence of these disease agents, the likelihood estimations for the occurrence of viable

copepods in these commodities are listed below.

Release assessment for infection of finfish with copepods

Commodity

type

Live

finfish

Whole fresh

dead finfish

Frozen

whole finfish

Frozen

fillets

Frozen fish

heads

Frozen

guts/offal

Likelihood of

release


_________________________________________________________________________________________________________________________________________________________________________________


136

www.digsfish.com


Marine and freshwater teleosts throughout Australia are already at risk of exposure to copepod parasites.

They naturally infect wild fishes and only certain species of copepods cause disease, usually in

circumstances where environmental conditions are favourable for their multiplication on the host.

Movements of live and whole fresh dead fishes via their use as bait could theoretically transmit these disease

agents to new regions, especially for copepods such as Lernaea cyprinacea in freshwater fishes and Caligus

epidemicus in estuarine and marine fishes. However, infection and establishment would occur only if

sufficient quantities of infective copepodid stages (i.e. an infective dose) were introduced into an area where

susceptible hosts were present. However, copepod infections can become established if susceptible hosts are

exposed even to only one viable copepodid larvae originating from one egg (B.K. Diggles, personal

observations), although several factors can influence infectivity. For example, the infectivity of copepodids

of C. epidemicus increased with increasing copepodid density and varied with the age of the copepodid,

peaking after 3 or 4 days post hatching at 26 or 19°C, respectively, then declining over time (Hallett and

Roubal 1995). Further, some hosts appeared refractory to infection, while other individuals of the same host

species were extremely susceptible to infection, resulting in an overdispersed distribution typical of that seen

in many parasite/host relationships (Hallett and Roubal 1995). In any case, the translocation of fish products

with viable copepods remains a potential route of introduction and infection of fish in new geographic areas.

Because live baitfish are not commercially available (except ornamental species sold from pet shops, which

are not intended for use as bait), fishes used for live bait are usually caught by recreational fishers at or near

the fishing location, hence the quantities of these commodities that are translocated long distances are likely

to be low. Only small quantities of fresh unfrozen fish are generally available from commercial fishing co-

operatives for local use as bait, and the vast majority of bait and berley that is translocated throughout the

country is frozen whole or processed marine fish, in which copepods will be unviable. Because of this, the

additional risk of exposure and establishment of copepods via use of bait is reduced, but remains non-

negligible, and the likelihood of exposure and establishment of copepods in new fish populations via

translocation is considered to be Low.


All size classes of fish can become infected with copepod parasites, and establishment of some species with

low host specificity, such Lernaea cyprinacea in freshwater fishes and Caligus epidemicus in estuarine and

marine fishes, can have negative impacts on the health of individual fish and their populations in the wild

(Hall 1983, Bond 2004, Hassan et al. 2008). Indeed, Bond (2004) demonstrated infection with L. cyprinacea

resulted in high mortality rates and reduced swimming ability in Galaxias olidus, and some native galaxiids

are classed as threatened or endangered, as are several other smaller fish species. There is evidence that the

pathological effects of L. cyprinacea infections are greater on smaller fish because the attachment organ of

the parasite penetrates deeply into the body, often causing damage to internal organs in smaller fish (Hassan

et al. 2008). Likewise, C. epidemicus has caused mortalities of wild fishes (Hewitt 1971) and is emerging as

an important disease agent in aquaculture of several fish species. For example, a single yellowfin bream held

in experimental seacages harboured over 6000 C. epidemicus (see Roubal 1994). Similarly, a single a

surgeonfish in the Philippines was recorded to have been infected by 5000 C. epidemicus (see Ho et al.

2004), while in Taiwan, heavy infections by C. epidemicus resulted in mass mortalities of cultured fish (Lin

_________________________________________________________________________________________________________________________________________________________________________________


137

www.digsfish.com

et al. 1996). Establishment of species such as Lernaea cyprinacea which produce grossly visible lesions on

infected fish, can also reduce the market value of affected fishes through the damage they inflict on the fillet.

Both Lernaea cyprinacea and Caligus epidemicus have the potential to cause damage to confined fish in

aquaculture and aquaria, and control of copepod parasites can be problematic and results in significant

increases in costs of production. No copepod parasites are listed by the OIE or NACA, but sea lice occur on

the reportable disease list for Tasmania (Table 3b). Hence the spread of these disease agents could have

adverse impacts on trade at a regional level. Considering all of these factors, establishment of the majority

of copepods in new areas would have mild or no biological consequences, however spread of C. epidemicus

and introduced species such as Lernaea cyprinacea could incur moderate biological consequences, which

would not be amenable to control in wild fish populations, and would cause some unwanted environmental

effects. It is therefore estimated that the consequences of introduction of the majority of copepods into

different parts of the Australian environment via use of infected bait would likely be Low, but for Lernaea

cyprinacea in freshwater fishes and Caligus epidemicus, the consequences would be Moderate.


The unrestricted risk associated with copepods is determined by combining the likelihood of entry and


copepods does not exceed the ALOP, except for Lernaea cyprinacea and Caligus epidemicus, which present

a low risk from movements of live finfish, suggesting that additional risk management is required for these

disease agents in these commodities.

Risk estimate for infection of finfish with copepods

Commodity type Live

finfish

Whole

fresh dead

finfish

Frozen

whole

finfish

Frozen

fillets

Frozen

fish heads

Frozen

guts/offal

Combined


and exposure


Consequences of

establishment



risk

Negligible

risk

Negligible

risk

Negligible

risk

Negligible

risk

_________________________________________________________________________________________________________________________________________________________________________________


138

www.digsfish.com

5.17 Infection of finfish and annelids with myxosporeans

5.17.1 Aetiologic agent: Myxosporean parasites (Class Myxosporea) of the genus Kudoa, and other

myxozoan parasites of finfish.


5.17.3 Australias status: Myxosporean infections have been reported from finfish in all States, and

Whirling disease caused by Myxobolus cerebralis is a reportable disease in all States (Table 3b).

5.17.4 Epidemiology

Myxosporeans are economically important histozoic and coelozoic endoparasites which have adversely

affected the culture of freshwater and marine fishes worldwide (Alvarez-Pellitero and Sitja-Bobadilla 1993,

Moran et al. 1999a). The higher taxonomy of the group has been controversial in the past but the link

between myxosporeans and cnidarians within basal metazoa has now been confirmed (Smothers et al. 1994,

Holland et al. 2010), and it appears that myxosporeans are highly specialised parasitic cnidarians. Further,

discovery that most myxosporeans have multihost lifecycles which utilise invertebrates (particularly

oligochaetes) as intermediate hosts (Markiw and Wolf 1983, Wolf and Markiw 1984, Kent et al. 1994),

revolutionised understanding of the epidemiology of the diseases these parasites cause in wild and cultured

fishes (Kent et al. 2001). For example, Kudoa thyrsites is a histozoic myxosporean parasite that infects a

wide range of marine fish (Lom and Dykova 1992). The presence of K. thyrsites spores in the fillet

musculature of infected fish is associated with 'milky flesh' and post-mortem myoliquefaction of the

musculature due to excretion of histolytic enzymes (Langdon 1991a, St-Hilaire et al. 1997). This parasite

occurs naturally in the flesh of several wild fishes in Australia, and also infects cultured fishes reared in sea

cages, including mahi mahi (Coryphaena hippurus) and Atlantic salmon (Salmo salar) (see Langdon 1991a,

Moran and Kent 1999, Moran et al. 1999a, Kent 2000). Many other myxosporean species exist in Australia

that can cause economically significant disease (Grossel et al. 2003) or marketing problems (Lester 1982) in

wild and cultured fishes in both freshwater and marine environments.

While there is evidence for direct fish to fish transmission for some marine myxosporeans (Diamant 1997,

Swearer and Robertson. 1999, Yasuda 2002), the lifecycles of the vast majority of marine myxosporeans are

unknown (Kent et al. 2001), and some freshwater myxosporeans, such as Myxobolus cerebralis, require

myxospores excreted by fish to be eaten by oligochaete worms before the actinospores released by the

oligochaete can infect new fish hosts (Markiw and Wolf 1983, Wolf and Markiw 1984). Attempts to transfer

K. thyrsites infection via feeding spores to Atlantic salmon failed to transmit infection, however Atlantic

salmon held in seacages in marine waters where K. thyrsites was enzootic became infected within 2 weeks

(Moran et al. 1999b). This suggests that fish in seacages may become infected indirectly through contact

with infective stages (actinospores) released by intermediate hosts, directly by eating presporogonic stages

excreted in other infected fishes (or via cannibalism), or even by obtaining presporogonic stages via blood

transferred by blood feeding vectors such as copepods or leeches (Moran et al. 1999b).


_________________________________________________________________________________________________________________________________________________________________________________


139

www.digsfish.com

Myxosporeans occur on a wide range of marine and freshwater fishes throughout Australia (Heiniger et al.

2011), including many species commonly used as bait or berley (Appendix 1). Some species appear

ubiquitous, including K. thyrsites which has been recorded worldwide (Moran et al. 1999a, Whipps et al.

2003). In Australia, K. thyrsites has been recorded from many species used as bait, including pilchards

(Sardinops sagax neopilchardus) (prevalence 67%), southern anchovy (Engraulis australis) (prevalence

12%), scaly mackerel (Sardinella lemuru) (prevalence 9% ), blue sprats (Spratelloides robustus) prevalence

3%, and cultured Salmo salar (see Langdon et al. 1992, Munday et al. 1998, O'Donoghue and Adlard 2000).

Other species which infect sedentary hosts may have highly restricted geographic distributions, although the

identity and distribution of many species is not known at this time. Some myxosporeans that occur in highly

mobile fishes may still have restricted distributions, perhaps depending on the distributions of intermediate

hosts. For example, Unicapsula seriolae which infects kingfish (Seriola lalandi) in SE QLD and northern

NSW (Lester 1982) and possibly in WA (Stevens and Savage 2010), does not occur in the cooler waters of

South Australia (Hutson et al. 1997) or New Zealand, even though adult kingfish can move across the

Tasman Sea (Diggles 2002). Myxosporean infective stages can also be spread via translocation of

invertebrate intermediate hosts, such as oligochaete worms (Lowers and Bartholomew 2003, Hallett et al.

2006), and annelids are used as bait in all parts of the country (Table 1) (Davies et al. 2008, 2010a).

Live baitfish are not commercially available (except ornamental species sold from pet shops, which are not

intended for use as bait), but they are sold fresh and the largest quantity of bait and berley translocated

throughout the country is frozen finfish. Live worms (polychaetes and oligochaetes) are widely retailed

throughout the country in specialist tackle shops, and these can originate from commercial farms which

supply live and freeze dried polychaetes to retailers around the country14. Live fishes and annelids used as

bait are likely to be effective routes of translocation of viable myxosporeans, but the majority of bait and

berley used throughout the country is frozen. Even so, myxosporean spores can tolerate freezing. For

example, spores of Myxobolus cerebralis can tolerate freezing to -20 °C for a week (Arsan and Bartholomew

2008), to 3 months (El-Matbouli and Hoffmann 1991), while Langdon et al. (1992) reported spore extrusion

in K. thyrsites spores from snap frozen fish. Myxosporean spores are also likely to survive in their host for

considerable periods of time after its death, especially at lower temperatures such as when fish are put on ice.

Taking into account the high prevalence of these disease agents, the likelihood estimations for the occurrence

of viable myxosporeans in these commodities are listed below.

Release assessment for infection of finfish and annelids with myxosporeans

Commodity

type

Live

finfish

and

annelids

Whole fresh

dead finfish

and annelids

Frozen

whole finfish

and annelids

Freeze

dried

annelids

Frozen fish

fillets/heads

Frozen

guts/offal

Likelihood of

release

High Moderate Low Low Low Low

14 http://www.aquabait.com.au/

_________________________________________________________________________________________________________________________________________________________________________________


140

www.digsfish.com


Marine and freshwater teleosts throughout Australia are already at risk of exposure to myxosporean

parasites. They are natural parasites of wild fishes and only certain species of myxosporeans cause disease,

usually in circumstances where transmission rates are increased when their hosts are confined at high

densities. Infection of fish is initiated by penetration of the epithelium of the skin or gills by sporoplasms

released by actinospores that contact the fish (Kent et al. 1994), or in myxosporeans with direct lifecycles, by

horizontal transmission via exposure of susceptible fish species to trophozoites and/or sporogenic stages

excreted by infected fish through the urine or faeces (Yasuda et al. 2002) or via the per-os route via

coprophagia or necrophagia (Padros et al. 2001). Movements of live, whole fresh dead or even frozen fishes

and annelids via their use as bait could theoretically transmit these disease agents to new regions, however

infection and establishment of myxosporeans in new hosts via movements of infected fish or annelids would

occur only if viable infective stages were introduced into an area where susceptible fishes and suitable

intermediate host were present. However for some species of myxosporeans, intermediate hosts may not be

necessary and they may be directly transmissible horizontally from fish to fish.

Live baitfish are not commercially available (except ornamental species sold from pet shops, which are not

intended for use as bait), so fishes used for live bait are usually caught by recreational fishers at or near the

fishing location, hence the quantities of these commodities that are translocated long distances are likely to

be low. However, live annelids are commercially available and are translocated widely from a small number

of commercial suppliers. Only small quantities of fresh unfrozen fish are available from commercial fishing

co-operatives for local use as bait, and the vast majority of bait and berley that is translocated throughout the

country is frozen whole or processed marine fish, in which myxosporean infective stages can survive for at

least short periods of time. Other potential routes of exposure include movements by vectors such as

piscivorous predators, as spores of some species of myxosporeans can survive passage through the guts of

birds and fishes and remain infective (El-Matbouli and Hoffmann 1991). Transfer of spores or other

infective stages via soil or other material lodged in fishing boots, waders or other angling equipment is

another likely source of spread of myxosporean parasites (Gates et al. 2008, 2009). Nevertheless, the risk of

exposure and establishment of myxosporeans via use of bait and berley remains non-negligible, and the

likelihood of exposure and establishment of myxosporeans in new fish populations via translocation is

considered to be Moderate.


All size classes of fish can become infected with myxozoan parasites, and establishment of some

myxosporeans can cause serious disease and mortalities in wild and cultured fish, particularly those species

that infect vital organs or neurological tissues (Markiw and Wolf 1983, Diamant 1997, Yasuda et al. 2002,

Grossel et al. 2003). Other myxosporean species that infect the muscle of the fillet may not lead directly to

death of their hosts, but can cause severe problems with the marketing of infected products due to post

mortem myoliquefaction (mushy flesh, e.g. Kudoa spp.) or myodegeneration of the infected musculature

during slow cooking (e.g. Unicapsula seriolae) (Lester 1982, Langdon 1991a, Moran et al. 1999a). No

myxosporean parasites are listed by the OIE or NACA, and only the exotic M. cerebralis is listed as a

reportable disease in all States (Table 3b). Hence the spread of these disease agents is unlikely to have

adverse impacts on trade. Many myxosporean disease agents are already widely disseminated in the

_________________________________________________________________________________________________________________________________________________________________________________


141

www.digsfish.com

Australian environment, however, since the myxosporean fauna of Australian fish and invertebrates is so

poorly known, there is a high chance that previously undescribed myxosporean pathogens remain to be

discovered. Considering all of these factors, establishment of the myxosporean parasites in new areas would

depend on the parasite species translocated, but pathogenic or myoliquefactive species could have moderate

biological consequences, which would not be amenable to control in aquaculture, but they would appear

unlikely to cause noticeable environmental effects. It is therefore estimated that the consequences of

introduction of myxosporeans into different parts of the Australian environment via use of infected bait

would likely be Low.


The unrestricted risk associated with myxosporeans is determined by combining the likelihood of entry and


myxosporeans exceeds the ALOP and presents a low risk from movements of live finfish and annelids, and

also fresh whole dead finfish and annelids, suggesting that additional risk management is required for these

disease agents in these commodities.

Risk estimate for infection of finfish and annelids with myxosporeans

Commodity type Live

finfish

and

annelids

Whole

fresh dead

finfish and

annelids

Frozen

whole

finfish and

annelids

Freeze

dried

annelids

Frozen fish

fillets/heads

Frozen

guts/offal

Combined

likelihood of

release and

exposure


Consequences of

establishment


Risk estimation Low risk Low risk Very low

risk

Very low

risk

Very low

risk

Very low

risk

_________________________________________________________________________________________________________________________________________________________________________________


142

www.digsfish.com

5.18 Viral infections of freshwater crayfishes

5.18.1 Aetiologic agent: Cherax destructor systemic parvo-like virus (CdSPV) , a parvovirus from C.

quadricarinatus (CqPV) and Cherax giardia-like virus (CGV).


5.18.3 Australias status: CdSPV reported in SA, and CGV reported in north QLD. These diseases are not

reportable in any State (Table 3b).

5.18.4 Epidemiology

Freshwater crayfish in Australia are known hosts to a range of disease agents, including several viruses

(Evans et al. 1998, Edgerton et al. 2002). The relationship between infection and disease remains obscure

for the majority of crayfish viruses, as many have been found to occur in otherwise apparently healthy

animals (Edgerton et al. 2002), suggesting disease may be preventable in aquaculture situations by proper

husbandry. However, three viruses have been reported to be associated with disease in cultured crayfish,

namely a Giardiavirus-like virus of Cherax quadricarinatus (CGV), which was reported to be associated

with disease in captive juvenile redclaw crayfish (C. quadricarinatus) in north Queensland (Edgerton et al.

1994, Edgerton 1996), Cherax destructor systemic parvo-like virus (CdSPV), which was observed in a

moribund adult yabby (C. destructor) in a farm in South Australia (Edgerton 1996, Edgerton et al. 1997), and

a parvovirus in C. quadricarinatus (CqPV) which was associated with a significant mortality event at an

aquaculture farm in north Queensland (Bowater et al. 2002).

CGV was first observed in C. quadricarinatus collected in a survey of crayfish collected from aquaculture

farms in northern Queensland, Australia (Edgerton et al. 1994). The virus was subsequently associated with

morbidity and mortalities in experimental juvenile C. quadricarinatus populations (Edgerton et al. 1994), but

does not appear to cause disease in adult C. quadricarinatus (see Edgerton et al. 2002). Edgerton and Owens

(1997) showed that C. quadricarinatus is first susceptible to infection by CGV immediately after moulting

into juvenile stage 3 (the first feeding stage), with prevalence of CGV increasing to 58% at 3 weeks post

stage 3, and 86% at 6 weeks post stage 3. CGV is probably transmitted per-os, possibly within faecal

strings attached to detrital particles, or associated with phytoplankton or zooplankton (Edgerton et al. 2002).

CdSPV was detected in a moribund C. destructor collected from a pond bank (Edgerton 1996, Edgerton et

al. 1997). This crayfish exhibited extensive necrosis in the gills, hepatopancreas and muscle, however the

involvement of CdSPV in these lesions was unclear as gill was the only tissue in which the cytopathic

lesions were common (Edgerton et al. 2002), though the virus was also associated with necrotic muscle.

CqPV was detected in diseased juvenile crayfish during an epizootic that resulted in 96% mortality in 2 farm

ponds over a period of 2 months, after which the disease spread to juvenile and adult crayfish in other ponds,

resulting in an eventual 50% loss in total farm production (Bowater et al. 2002). Diseased juvenile crayfish

were lethargic, disoriented and anorexic, and had soft shells, and the disease was reproduced experimentally

by inoculating healthy crayfish with a cell free extract prepared from diseased crayfish, confirming the

pathogenicity of CqPV (Bowater et al. 2002). Large numbers of viral inclusions were observed in tissues of

endodermal, ectodermal and mesodermal origin of CqPV infected crayfish (Bowater et al. 2002).

_________________________________________________________________________________________________________________________________________________________________________________


143

www.digsfish.com


CGV is very common in farmed redclaw in northern Queensland (Edgerton et al. 1996). CdSPV has been

observed in only one C. destructor from a farm in southeastern South Australia (Edgerton 1996, Edgerton et

al. 1997), but since no other studies have been done; the distribution and prevalence of CdSPV remains

unknown (Edgerton et al. 2002). It appears that CqPV has been reported only from one farm in north

Queensland (Bowater et al. 2002). In all cases the origins of each virus are unclear.

Freshwater crayfish are used by around 8% of recreational anglers (Table 1). Freshwater crayfish are not

generally sold as live or frozen bait in retail outlets, which generally leaves fishers to catch them themselves,

though a small number of crayfish farms do sell live cultured crayfish for bait. Because they are relatively

hardy baits that are easy to transport, it is possible for freshwater crayfish to be translocated large distances

by private recreational fishers. So, even though the quantity of crayfish that is translocated live is likely to

be low, use of live crayfish as bait would be highly likely to translocate these viruses. There appears to be no

information available for any of these viruses in relation to their resistance to freezing or physical and

chemical treatments, or whether they may be translocated by vectors such as birds. However, for the

purposes of this analysis it will be assumed that viruses from freshwater crayfish are similar to other viruses

of aquatic animals which can remain viable after freezing, but may experience reduced viability upon

thawing.

Taking into account the quantities and types of freshwater crayfish products used as bait or berley (Table 1),

and the high prevalence of some of these disease agents, the likelihood estimations for the occurrence of

viruses in these commodities are listed below.

Release assessment for viral infections of freshwater crayfishes

Commodity

type

Live

crayfish

Whole fresh

dead crayfish

Frozen

whole

crayfish

Frozen

crayfish

tails

Frozen

crayfish

heads

Likelihood of

release

High High Low Very low Low


Freshwater crayfish in some regions of Australia are already at risk of exposure to viruses that may occur in

wild populations. The quantities of freshwater crayfish sold as live bait is likely to be relatively small,

however private recreational anglers may still catch their own at the fishing location, and since these animals

are hardy and transport well, a small proportion of anglers may translocate live freshwater crayfish to use as

bait in different rivers and lakes in freshwater regions. Also, fresh or frozen freshwater crayfish are widely

distributed as food fish and hence some of this product could be diverted to use as bait or berley around the

_________________________________________________________________________________________________________________________________________________________________________________


144

www.digsfish.com

country. Assuming that freshwater crayfish viruses survive freezing, some virus is likely to remain viable

after thawing, this suggests that direct pathways exist for viruses carried in live, fresh dead and frozen

freshwater crayfish products used as bait or berley to enter freshwater environments, thus potentially

exposing wild crayfish to these viruses.

Nothing is known about the transmission, potential host range or viability of CdSPV. CGV is probably

transmitted per-os, possibly within faecal strings attached to detrital particles, or associated with

phytoplankton or zooplankton, though the infective dose required for transmission is unknown, as is its host

range (Edgerton et al. 2002). CqPV appears to be transmitted horizontally given its role in the epizootic and

experimental infection work reported by Bowater et al. (2002). Injection of 0.1 ml of cell free filtrate from

infected crayfish into healthy crayfish was sufficient to cause mortalities starting on day 17 post injection

and continuing to day 73, at which time the cumulative mortality of virus injected crayfish was 100%

(Bowater 2002), however the minimum infective dose required for transmission was not determined, nor

were trials done to determine if the virus could be transmitted by other routes such as immersion or per-os.

The host range of CqPV is also unknown. Without this information, it is difficult to determine whether an

infective dose could be transmitted via the water to susceptible hosts in the immediate vicinity of bait or

berley that was clinically or subclinically infected with CqPV. These viruses all were recorded from

aquacultured freshwater crayfish only, and some States have bans on use of aquacultured products as bait (T.

Hawkesford, personal communication). However, given that pathways still exist for translocation and spread

of crayfish viruses into the environment via use of privately caught bait, and the fact that freshwater crayfish

are commonly used as live bait, the risk of exposure and establishment is non-negligible, and the likelihood

of exposure and establishment of crayfish viruses in new crayfish populations is considered to be Low.


Because of their association with disease, it is possible that if CqPV, and also CGV and /or CdSPV were

translocated, they could represent a threat to the health of other crayfish (including rare and threatened native

freshwater crayfish, see Coughran and Leckie 2007, Coughran et al. 2009) or even other crustaceans.

However without information on the host range of each virus, it is very difficult to determine the likely

consequences if they were to be translocated. However, it is clear that CqPV at least, poses a serious threat

to aquaculture of C. quadricarinatus (see Bowater et al. 2002). No viruses of freshwater crayfish are listed

by the OIE or NACA, and none of them are listed as a reportable diseases in any State (Table 3b). Hence the

spread of these disease agents is unlikely to have adverse impacts on trade at this time. Given the paucity of

surveillance of diseases of wild freshwater crayfish, it is difficult to assess whether wild populations

(including rare and threatened native freshwater crayfish) are already threatened by disease agents such as

viruses. However the emergence of CqPV in the one farm in north Queensland confirms that the viral

diseases of Australian freshwater crayfish are so poorly known, there is a high chance that previously

undescribed viral pathogens remain to be discovered. Considering all of these factors, the consequences of

establishment of these viruses in new areas would depend on the identity of the virus that was translocated

and that of the host infected, but infection of rare and threatened native crayfish with a novel virus that

would not be amenable to control in wild populations could have significant biological and environmental

consequences. Similarly, introduction of a virus such as CqPV into new geographical areas could pose a

significant threat to crayfish farming in those regions. It is therefore estimated that the consequences of

_________________________________________________________________________________________________________________________________________________________________________________


145

www.digsfish.com

introduction of crayfish viruses into different parts of the Australian environment via use of infected bait

would likely be Moderate.


The unrestricted risk associated with freshwater crayfish viruses is determined by combining the likelihood


risk estimate for crayfish viruses exceeds the ALOP for live freshwater crayfish and whole dead freshwater

crayfish, suggesting that additional risk management for these disease agents is required for these

commodities.

Risk estimate for viral infections of freshwater crayfishes

Commodity type Live

crayfish

Whole fresh

dead crayfish

Frozen whole

crayfish

Frozen

crayfish tails

Frozen

crayfish heads

Combined likelihood

of release and

exposure

Low Low Very Low Very low Very Low

Consequences of

establishment

Moderate Moderate Moderate Moderate Moderate

Risk estimation Low risk Low risk Very low risk Very low risk Very low risk

_________________________________________________________________________________________________________________________________________________________________________________


146

www.digsfish.com

5.19 Infection of prawns with Gill Associated Virus (GAV)

5.19.1 Aetiologic agent: Gill Associated Virus (GAV), a positive sense ssRNA virus in the genus Okavirus

in the family Roniviridae of the order Nidovirales.


5.19.3 Australias status: Reported in QLD, NSW, NT and WA, and is a reportable disease in all States

except the ACT (Table 3b).

5.19.4 Epidemiology

Gill Associated Virus (GAV) and lymphoid organ virus (LOV) have been reported from diseased and

healthy Penaeus monodon from Australia (Cowley et al. 1999, 2004, Wijegoonwardane et al. 2009). Both

viruses are members of the Yellowhead disease viral complex which includes yellowhead virus (YHV),

which causes yellowhead disease (YHD) which is enzootic in P. monodon farms in Taiwan, Indonesia,

China, Malaysia and the Philippines (Limsuwan 1991, Lightner 1996, Flegel 1997), however GAV and LOV

are genetically distinct from YHV (Cowley et al. 2004). Yellowhead-like viruses (YHLV) appear to

naturally infect only penaeids (Munro and Owens 2007). GAV has been the primary cause of a yellowhead-

like disease and associated mortalities that have affected the prawn aquaculture industry in Australia since

1994 (Spann et al. 1995, 1997a). The virus is indistinguishable from YHV by TEM, infects a similar range

of tissues, and causes similar pathology, however for GAV infections mortality is sometimes preceded by

varying degrees of red colouration of the body and pink to yellow colouration of the gills, with no evidence

of pale body colouration or yellowing of the cephalothorax in GAV affected prawns (Spann et al. 1997a).

Prior to the identification of GAV, a virus with similar morphology (LOV) was observed to be common in

healthy P. monodon in Australia (Spann et al. 1995). LOV causes the formation of distinct foci of

hypertrophic cells (spheroids) in the lymphoid organ which otherwise remains structurally intact (Spann et

al. 1995). LOV infections are chronic and widespread in wild and farmed P. monodon on Australias east

coast, suggesting the virus is a variant of GAV (Cowley et al. 2000, Munro and Owens 2007) which does not

cause disease in uncompromised P. monodon.

Several prawn species farmed commercially in Australia, including P. esculentus, P. merguiensis and P.

japonicus, are susceptible to experimental GAV infection (Spann et al. 2000). However, some age/size and

species-related differences have also been noted in their susceptibility to disease. For example, P. monodon,

P. esculentus, P. merguiensis, and small P. japonicus displayed overt signs of disease and mortalities which

reached 82 to 100% within 23 d post-injection, however cumulative mortalities in P. esculentus and P.

merguiensis were significantly lower than for P. monodon of the same size class (Spann et al. 2000).

Medium sized P. japonicus also developed overt signs of disease but cumulative mortalities were not

significantly higher than uninfected controls, while adult P. japonicus did not display symptoms of disease

and there were no significant mortalities up to 23 d post-injection (Spann et al. 2000). The ability of GAV to

cause disease experimentally in a wide range of penaeids and some palaemonids suggests these viruses

represent a considerable threat to a range of species of cultured prawns. Contaminated water is the major

method of natural infection with horizontal transmission through the water from infected prawns and by

_________________________________________________________________________________________________________________________________________________________________________________


147

www.digsfish.com

cannibalism of weak or moribund prawns (Flegel et al. 1995, 1997, Lightner 1996a). Transport water, intake

water, nets and other equipment are likely sources of virus introduction, while in hatcheries vertical

transmission (via contamination of the egg surface) has been recorded in GAV, with virus associated with

eggs originating from either of the male or female parents (Cowley et al. 2002).


GAV and LOV infect several prawn species used as bait (Table 1), namely P. monodon, P. esculentus, P.

merguiensis and other penaeids. The distribution of these viruses appears restricted to the northern regions

of Australia, with GAV being reported in aquacultured prawns in QLD, NSW, NT and WA, but absent in

South Australia (Roberts et al. 2010). The virus is enzootic in wild prawns along the eastern seaboard at

prevalences as high as 95% (Munro and Owens 2007, Oanh et al. 2011), and may also be found in wild

prawns in other parts of the country, especially adjacent to prawn farming areas. Large quantities of prawns

are used as bait throughout Australia (Kewagama Research 2002, 2007). Live prawns are generally not


and some of these could be translocated live. However, the majority of the volume of prawns distributed as

bait and berley throughout the country is frozen whole prawns (mostly Metapenaeus spp.), while fresh and

frozen prawns (Penaeus spp.) are also widely distributed for human consumption, and around 8% of

recreational fishers divert these to use as bait or berley (Kewagama Research 2007). GAV infected tissues

stored at –70°C can maintain infectivity (Spann et al. 2000), though there appears to be no information on

how long GAV can remain viable in tissues stored at -18°C. It would be nevertheless expected (Durand et

al. 2000, Biosecurity Australia 2009) that a large percentage of GAV could survive and remain viable after at

least one freeze-thaw cycle during freezing, storage and transport within Australia. YHLV remain viable

outside the host in aerated seawater for up to 72 h (Flegel et al. 1995). While some prawn viruses remain

infectious following passage through a bird gut, YHLV virions do not (Vanpatten et al. 2004). Live or fresh,

whole, green (uncooked) penaeid and palaemonid prawns from areas where GAV occurs in wild and/or

cultured prawns could therefore contain viable GAV. Frozen whole uncooked prawns would also be

expected to contain viable GAV, especially if prawns were taken from affected aquaculture ponds. Highest

GAV concentrations occur in the cephalothorax, with Lu et al. (1995) finding gill and head soft tissues

contained 10 to 800-fold higher titers of YHLV than the other tissues and organs tested. Removal of the

head section would therefore significantly reduce (but would not completely eliminate) the levels of GAV in

translocated prawns. Taking into account the quantities of prawn products used as bait or berley (Table 1),

and the high prevalence of the disease agent, the likelihood estimations for the occurrence of GAV in these


Release assessment for infection of prawns with GAV

Commodity

type

Live

prawns

Whole fresh

dead prawns

Frozen

whole

prawns

Frozen

prawn

tails

Frozen

prawn

heads

Likelihood of

release

High High Low Very Low Low

_________________________________________________________________________________________________________________________________________________________________________________


148

www.digsfish.com


Prawns throughout northern Australia are already at risk of exposure to GAV. However, translocation of

GAV infected prawns from areas where GAV is enzootic via their use as bait could transport the virus to

new regions. Infection and establishment of GAV in new hosts would occur only if viable viral particles

were introduced into an area where susceptible prawns were present under suitable environmental conditions

for transmission. The virus has a wide host range, and several penaeids that form the basis of important

recreational and commercial fisheries in southern Australia (e.g. Melicertus latisulcatus, P. plejebus) may be

susceptible to infection with GAV if they were to come in contact with the virus. YHLV remain infective in

the water for up to 3 days (Flegel et al. 1995), and susceptible crustaceans can become infected with YHLV

via per-os exposure through cannibalism and also by water-borne transmission through cohabitation (Flegel

et al. 1995, 1997, Lightner 1996). Although it is known that YHLV can be transmitted horizontally by co-

habitation, the minimum infective dose for transmission of GAV via immersion does not seem to have been

quantified. Large viral copy numbers occur in haemolymph of prawns clinically infected with YHV (5 x 107

/µl = 5 x 1010/ml, see Ma et al. 2008) and GAV (1 x 109 /ng, see Oanh et al. 2011). Virulent strains of YHV

were successfully transmitted via the per-os route by feeding prawns with a dose of 5 x 105 viral copies/g

(Sritunyalucksana et al. 2010), and GAV was transmitted and caused nearly 100% mortality when a similar

dose was injected (Oanh et al. 2011), suggesting threat of infection is lower via the per-os route as YHV is

supposed to be around 106 times more virulent than GAV (Walker and Mohan 2009). Ma et al. (2009)

demonstrated that palaemonid prawns were also susceptible to YHV by per-os exposure to prawn muscle

with around 4 x 104 viral copies/g, and they could act as reservoirs of infection for at least 36 days. These

data suggest that GAV may be transmissible by the per-os route and cause disease, but probably only if

susceptible hosts consume clinically diseased prawns. Water temperature is likely to play an important role

in transmission of the disease, with little information available on transmission rates of GAV at lower water

temperatures typical of southern States.

Wild prawns such as P. monodon often carry sub-clinical GAV infections and would be unlikely to suffer

mortalities unless they were exposed to significant stressors (Munro and Owens 2007). Once infected,

surviving prawns may transmit the infection vertically, which would further enhance the likelihood of

establishment of new GAV infections in the environment. If the susceptible crustacean in the index case

died, it is well known that dead and moribund crustaceans can be a source of GAV transmission, especially

given the high viral titres in clinically diseases prawns, however predation of moribund crustaceans by non

susceptible species such as fish and crabs may be an important factor modulating transmission and spread of

GAV in some index cases. Taking these various factors into consideration, the risk of exposure and

establishment of GAV via use of bait and berley remains non-negligible, and the likelihood of exposure and

establishment of GAV in new prawn populations via translocation is considered to be Low.


GAV naturally infects wild populations of prawns throughout northern Australia at high prevalences, with no

known detrimental impacts on wild prawn fisheries at this time (Biosecurity Australia 2009). Therefore it

appears that establishment of the virus in new regions would have little if any effect on the viability of prawn

populations or the fisheries that rely on them. However, GAV is a recognized pathogen of cultured prawns,

and hence its introduction into new areas would likely have significant consequences for prawn aquaculture

_________________________________________________________________________________________________________________________________________________________________________________


149

www.digsfish.com

industries in the affected region. Although some adaptation to GAV may occur over time, the presence of

the virus would remain a significant obstacle to industry competitiveness and profitability. YHLV are listed

by the OIE or NACA, and GAV is listed as a reportable disease in all States except the ACT (under review

in the NT) (Table 3b). Hence the spread of GAV to new areas is likely to adversely impact on trade.

Considering all of these factors, establishment of GAV in new areas would have significant consequences for

prawn aquaculture, that would not be amenable to control, but it would appear unlikely to cause noticeable

environmental effects. It is therefore estimated that the consequences of introduction of GAV into different

parts of the Australian environment via use of infected bait would likely be Moderate.


The unrestricted risk associated with GAV is determined by combining the likelihood of entry and exposure

(from Table 5) with the consequences of establishment (Table 7). The unrestricted risk estimate for GAV

exceeds the ALOP for live prawns and whole fresh dead prawns, suggesting that additional risk management

for this disease agent is required in these commodities.

Risk estimate for infection of prawns with GAV

Commodity type Live

prawns

Whole fresh

dead prawns

Frozen whole

prawns

Frozen

prawn tails

Frozen prawn

heads

Combined likelihood

of release and

exposure

Low Low Very Low Very Low Very Low

Consequences of

establishment


Risk estimation Low risk Low risk Very low risk Very low

risk

Very low risk

_________________________________________________________________________________________________________________________________________________________________________________


150

www.digsfish.com

5.20 Infection of crustaceans with Hepatopancreatic Parvovirus (HPV)

5.20.1 Aetiologic agent: Hepatopancreatic parvoviruses (HPV) are single-stranded, non-enveloped DNA

viruses classified within the family Parvoviridae in the genus Densovirus (Bonami et al. 1995a). There are

various strains, including Penaeus merguiensis densovirus (PmergDNV), Penaeus monodon densovirus

(PmonDNV), Parvo-like virus of Penaeus japonicus (P-PJ), and Penaeus chinensis parvovirus (HPVchin)

(see LaFauce et al. 2007a, Tang et al. 2008).


5.20.3 Australias status: Reported in prawns, crabs and crayfish from QLD (Owens et al. 2010). This

disease is not reportable in any State (Table 3b).

5.20.4 Epidemiology

Hepatopancreatic parvoviruses have a cosmopolitan geographic distribution (Tang et al. 2008) and their

known host range encompasses postlarvae and juveniles of at least ten wild and farmed penaeid prawn

species worldwide, including P. stylirostris, P. vannamei, P. monodon, P. merguiensis, P. indicus, P.

semisulcatus, as well as the freshwater prawn Macrobrachium rosenbergii (see Anderson et al. 1990,

Gangnonngiw et al. 2009), blue swimmer crab Portunus pelagicus and mud crab Scylla serrata (see La

Fauce et al. 2007a, 2007b). Similar viruses have also been recorded from other crab species (Mari and

Bonami 1988). Strains isolated from penaeid prawns have been shown to be able to infect crayfish and

crickets Acheta domesticus under certain conditions, but virus replication did not occur suggesting a short

term carrier state was achieved (LaFauce and Owens 2007, 2008). Hepatopancreatic parvoviruses are often

found in healthy crustaceans and they are generally not recognized as causing severe disease (Lightner and

Redman 1985). However, HPV infection and in particular, multiple infections with HPV and other

pathogenic agents such as monodon baculovirus (MBV), has been correlated statistically with stunted growth

in P. monodon (see Flegel et al. 1999, 2004).

HPV was initially reported from Asia (Chong and Loh 1984, Lightner et al. 1985), and Australia (Paynter et

al. 1985, Roubal et al. 1989a, Spann et al. 1997b). There is evidence that HPV has been spread to many

parts of the world via the movement of live animals (Colorni et al. 1987, Turnbull et al. 1994, Manjanaik et

al. 2005). Gene sequencing has identified three main groups of HPV, from Korea, Thailand, and Australia

with a mean of 17% gene sequence divergence (Tang et al. 2008). New data suggest that HPV is an

emerging disease and that different strains of HPV are associated with different species and/or geographical

areas (LaFauce et al. 2007a, Tang et al. 2008).


HPV is known to naturally infect a wide range of crustaceans, including several species used as bait or berley

in Australia (Table 8), including Macrobrachium rosenbergii, Macrobrachium spp., P. monodon, P.

esculentus, P. merguiensis, Penaeus spp., Portunus pelagicus, Scylla serrata and Cherax quadricarinatus.

HPV infections are often asymptomatic and often occur at high prevalence. Large quantities of crustaceans

_________________________________________________________________________________________________________________________________________________________________________________


151

www.digsfish.com

are used as bait throughout Australia (Kewagama Research 2002, 2007). Live prawns, crabs or freshwater

crayfish are generally not commercially available, so the quantities translocated by the approximately 11% of

recreational fishers who catch their own live crustaceans is likely to be low. The majority of the volume of

crustaceans distributed as bait and berley throughout the country is frozen whole prawns (Metapenaeus spp.),

while fresh and frozen prawns (Penaeus spp.), crabs and freshwater crayfish are also widely distributed for

human consumption, and some of this is known to be diverted to use as bait or berley (Kewagama Research

2007). Infected fresh, whole, green (uncooked) crustaceans and frozen whole uncooked crustaceans would

also be expected to contain viable virus, as it is known that hepatopancreatic parvoviruses survive freezing (-

80°C) and remain infective in subsequent experimental trials (Catap et al. 2003). While survival and

viability of these viruses upon thawing from normal commercial freezing temperatures of -18°C does not

appear to be documented, it would be reasonable to expect that a large percentage of these viruses could

survive and remain viable after at least one freeze-thaw cycle during commercial freezing, storage and

transport within Australia.

HPV occurs in cells of the hepatopancreatic tubule epithelia and rarely in cells of the anterior midgut or

caecum epithelia (Lightner and Redman 1985). Hence removal of the head section containing the

hepatopancreas could result in a marked reduction in the viral load of infected prawns. Taking into account

the quantities of crustacean products used as bait or berley (Table 1), and the high prevalence of the disease

agent, the likelihood estimations for the occurrence of HPV in these commodities are listed below.

Release assessment for infection of prawns and crabs with HPV

Commodity

type

Live

crustaceans

Whole fresh

dead

crustaceans

Frozen

whole

crustaceans

Frozen

prawn

tails

Frozen

prawn

heads

Likelihood of

release

High High Low Very Low Low


Prawns throughout northern Australia are already at risk of exposure to HPV. However, translocation of

HPV infected prawns from areas where HPV is enzootic via their use as bait could transport the virus to new

regions. Infection and establishment of HPV in new hosts would occur only if viable viral particles were

introduced into an area where susceptible prawns were present under suitable environmental conditions for

transmission. The virus has a wide host range, and many crustaceans that form the basis of important

recreational and commercial fisheries in southern Australia (e.g. Melicertus latisulcatus, P. plejebus,

southern populations of Portunus pelagicus) may be susceptible to infection with HPV if they were to come

in contact with the virus. Susceptible crustaceans can become infected with HPV via per-os exposure, while

horizontal transmission of HPV due to co-habitation is considered unlikely, but possible (Paynter et al. 1985,

Flegel et al. 1995). Catap et al. (2003) successfully transmitted HPV to post larval P. monodon by feeding

_________________________________________________________________________________________________________________________________________________________________________________


152

www.digsfish.com

them once with 2 grams of infected frozen PL, while a dose of 106 virions orally and by injection were both

sufficient to kill redclaw crayfish in experimental challenges (LaFauce and Owens 2007). Susceptible

species of crustaceans feeding on clinically diseased prawns or crabs used as bait or berley may therefore

receive sufficient dose of the virus to become infected, however it is not known if HPV can be transmitted

via the per-os route if subclinically affected crustaceans were used. HPV may not be inactivated by passage

through the bird digestive system and as such, spread via mechanical vectors such as seagulls is possible

(Biosecurity Australia 2009). The single-stranded, non-enveloped virion structure and DNA-based genome

confers HPV a much greater resilience compared to some other lipid-containing enveloped viruses such as

WSSV and YHV (Biosecurity Australia 2009). Water temperature is likely to play an important role in

transmission of the disease, with little information available on transmission rates of HPV at lower water

temperatures typical of southern States.

Wild crustaceans commonly carry sub-clinical HPV infections, and are unlikely to suffer mortalities unless

they were exposed to significant stressors. If a clinically diseased crustacean was used as bait and the

susceptible crustacean in the index case became infected and died, it is possible that dead and moribund

crustaceans could be a source of HPV, however predation of moribund crustaceans by non susceptible

species such as fish may be an important factor modulating transmission and spread of HPV in some index

cases. Taking these various factors into consideration, the risk of exposure and establishment of HPV via

use of bait and berley remains non-negligible, and the likelihood of exposure and establishment of HPV in

new prawn populations via translocation is considered to be Low.


HPV naturally infects wild populations of crustaceans in many areas of northern Australia, with no known

detrimental impacts on wild crustacean fisheries (Biosecurity Australia 2009), and it appears that the

presence of HPV in healthy wild crustaceans is a normal occurrence in regions where this virus is endemic.

Therefore, establishment of the virus in new regions is unlikely to have any detectable effect on the viability

of crustacean populations or the fisheries that rely on them. HPV has been associated with disease in

cultured prawns, and hence its introduction into new areas would likely have some consequences for prawn

aquaculture industries in the affected region. HPV infection concurrent with stress tends to predispose

cultured crustaceans to infection with other (perhaps more virulent) pathogenic agents (Biosecurity Australia

2009), hence over time dual infection with other pathogenic agents may occur and result in mortality and

stunting (Flegel et al. 1999). Although HPV is not likely to cause a severe epidemic in larger cultured

prawns, it may still cause economic loss (Flegel et al. 1999) which would have a detrimental effect on the

profitability and hence viability of the prawn aquaculture industry in Australia. HPV is not listed by the OIE

or NACA, nor is it listed as a reportable disease in any State (Table 3b). Hence the spread of HPV to new

areas is not likely to adversely impact trade. Considering all of these factors, establishment of HPV in new

areas would have mild to moderate consequences for prawn aquaculture, that would probably be amenable to

control, but it would appear unlikely to cause noticeable environmental effects. It is therefore estimated that

the consequences of introduction of HPV into different parts of the Australian environment via use of

infected bait would likely be Low.

_________________________________________________________________________________________________________________________________________________________________________________


153

www.digsfish.com


The unrestricted risk associated with movements of crustaceans infected with HPV is determined by

combining the likelihood of entry and exposure (from Table 5) with the consequences of establishment

(Table 7). The unrestricted risk estimate for HPV does not exceed the ALOP for any of the commodity

types, suggesting that additional risk management is not required for this disease agent at this time.

Risk estimate for infection of prawns and crabs with HPV

Commodity type Live

crustaceans

Whole fresh

dead

crustaceans

Frozen whole

crustaceans

Frozen

prawn tails

Frozen prawn

heads

Combined likelihood

of release and

exposure

Low Low Low Very Low Low

Consequences of

establishment

Low Low Low Low Low


risk

Very low risk Very low risk Very low

risk

Very low risk

_________________________________________________________________________________________________________________________________________________________________________________


154

www.digsfish.com

5.21 Infectious Hypodermal and Haematopoietic Necrosis of prawns (IHHNV)

5.21.1 Aetiologic agent: Infectious Hypodermal and Haematopoietic Necrosis Virus (IHHNV) is single-

stranded, non-enveloped DNA virus classified as Penaeus stylirostris densovirus (PstDNV) in the genus

Brevidensovirus (Fauquet et al. 2005) within the family Parvoviridae (Bonami et al. 1990).


5.21.3 Australias status: Reported in QLD and the NT, and is a reportable disease in all States (Table 3b).

5.21.4 Epidemiology

The disease Infectious Hypodermal and Haematopoietic Necrosis was first detected in 1981 in juvenile P.

stylirostris imported into Hawaii from Costa Rica and Ecuador (Lightner et al. 1983a, 1983b). The most

pathogenic strains of the virus appear to have originated from Asia (Lightner 1999, Tang et al. 2003) and

were spread throughout the Americas via movements of live prawns as broodstock and PL (Lightner 1996,

Tang et al. 2003). IHHNV is now found at prevalences of 28% and 44–100% in wild South American P.

vannamei and L. stylirostris, respectively (Unzueta–Bustamante et al. 1998, Morales–Covarrubias et al.

1999, Nunan et al. 2000). IHHNV is considered to be endemic in wild P. monodon in South-East Asia and

Australia with reported prevalence as high as 94%, although there usually is no associated disease in this

species (Flegel et al. 2004, Krabsetsve et al. 2004). However, when assessing prevalences generated using

PCR-based surveys for IHHNV in P. monodon, the fact that certain virus-related sequences are integrated

into the genome of P. monodon in parts of Australasia and Africa must be considered and taken into account

(Tang and Lightner 2006, Tang et al. 2007, Rai et al. 2009b).

There are at least three strains of IHHNV (OIE 2010b). The Philippines strain, which appears to have been

the original source of infection of the Americas, is considered to be the most virulent, being associated with

mortality in farmed L. stylirostris and runt deformity syndrome in farmed P. vannamei and P. monodon (see

Primavera and Quinitio 2000). The South East Asian strain and Indian Ocean strain found in Madagascar

and Australia are considered to be less virulent to penaeids (Tang et al. 2003, Krabsetsve et al. 2004).

Australia appears to have both the latter strains in cultured populations of P. monodon (Biosecurity Australia

2008, 2009). IHHNV infection in Australasia has been reported in not only P. monodon, but a range of

cultured prawns including P. japonicus and P. vannamei (see Bell and Lightner 1984, Lightner et al. 1997),

as well as PL and juvenile Macrobrachium rosenbergii (see Hsieh et al. 2006). Experimentally infected

species include a wide range of other penaeids, including species such as P. semisulcatus and P. japonicus

that are both endemic to Australia and used as bait (Table 8) (Brock and Lightner 1990, Lightner 1996).

However, P. merguiensis appears to be refractory to infection (Brock and Lightner 1990, Lightner 1996). To

date, IHHNV has not been reported in other crustacean groups such as crabs or lobsters.

IHHNV is mainly a disease of P. stylirostris, which does not occur in Australia, but the virus occasionally

results in chronic runt deformity syndrome in P. monodon (Primavera and Quinitio 2000). However,

whether disease occurs depends on the stain of the virus involved, and infection with the strains of IHHNV

in P. monodon in Australia is not necessarily associated with disease (Krabsetsve et al. 2004). IHHNV

_________________________________________________________________________________________________________________________________________________________________________________


155

www.digsfish.com

infection can occur by horizontal transmission through the ingestion of dead infected prawns or by contact

with water containing infected animals (Lotz 1997). Vertical transmission occurs through the presence of

IHHNV in the ovaries of infected P. vannamei females (Motte et al. 2003). Thus, both horizontal and vertical

transmission may increase IHHNV prevalence in both wild and cultured prawns.


IHHNV is known to naturally infect a wide range of crustaceans, including several species used as bait or

berley in Australia (Table 8), such as Penaeus monodon, Penaeus esculentus, and Metapenaeus spp.

IHHNV infections are often asymptomatic and can occur at high prevalences. Large quantities of prawns are

used as bait throughout Australia (Kewagama Research 2002, 2007). Live prawns are generally not


and some of these could be translocated live. However, the majority of the volume of prawns distributed as

bait and berley throughout the country is frozen whole prawns (Metapenaeus spp.), while fresh and frozen

prawns (Penaeus spp.) are also widely distributed for human consumption, and around 8% of recreational

fishers divert these to use as bait or berley (Kewagama Research 2007). Both live and fresh, whole, green

(uncooked) prawns would be expected to contain viable virus.

Frozen whole uncooked prawns would also be expected to contain similar amounts of virus to fresh product,

as it is known that IHHNV is able to survive frozen storage between –20°C and –80°C and remain infectious

for over five years (OIE 2010b), while the Australian strain of IHHNV was found to remain infective after

15 years of frozen storage (Biosecurity Australia 2009). IHHNV can also survive several freeze–thaw cycles

with minimal loss of virulence (OIE 2010b). Therefore it is highly likely that a large percentage of these

viruses could survive and remain viable after at least one freeze-thaw cycle during commercial freezing,

storage and transport within Australia. IHHNV replicates in the cytoplasm of cells of ectodermal (epidermis,

gills, fore and hind gut, antennal gland and neurons) and mesodermal origin (haematopoietic tissue,

haemocytes, striated muscle, heart, lymphoid organ and connective tissues), however infection of the midgut

epithelium is rare (Lightner et al. 1983b). Because of this, removal of the head section alone would not

necessarily result in a marked reduction in the viral load of infected prawns, however removal of head and

shell would markedly reduce viral loads and head and shell wastes would be expected to contain high

concentrations of virus (Biosecurity Australia 2009). Taking into account the quantities of crustacean

products used as bait or berley (Table 1), and the prevalence of the disease agent, the likelihood estimations

for the occurrence of IHHNV in these commodities are listed below.

Release assessment for infection of prawns with IHHNV

Commodity

type

Live

Prawns

Whole fresh

dead prawns

Frozen

whole

prawns

Frozen

prawn

tails

Frozen

prawn

heads

Likelihood of

release

High High Moderate Moderate Moderate

_________________________________________________________________________________________________________________________________________________________________________________


156

www.digsfish.com


Prawns throughout northern Australia are already at risk of exposure to IHHNV. However, translocation of

IHHNV infected prawns from areas where IHHNV is enzootic via their use as bait could transport the virus

to new regions. Infection and establishment of IHHNV in new hosts would occur only if viable viral

particles were introduced into an area where susceptible prawns were present under suitable environmental

conditions for transmission. The virus has a wide host range, but only infects prawns and does not appear to

infect other crustaceans. Nevertheless, prawn species that form the basis of important recreational and

commercial fisheries in southern Australia (e.g. Melicertus latisulcatus, P. plejebus) may be susceptible to

infection with IHHNV if they were to come in contact with the virus.

Susceptible wild crustaceans can become infected with IHHNV via horizontal routes such as per-os

exposure, as well as horizontally via the water by co-habitation (Lightner et al. 1983a, 1983b, Lotz 1997).

Penaeus monodon is reported to require prolonged exposure to IHHNV before infection occurs, nevertheless

the high stocking densities used in intensive aquaculture provide suitable conditions for transmission of

IHHNV (Browdy et al. 1993). Susceptible species of prawns feeding on clinically diseased prawns used as

bait or berley may receive sufficient dose of the virus to become infected, however it appears unlikely the

IHHNV could be transmitted via the per-os route if subclinically affected crustaceans were used. IHHNV

has also been shown to maintain infectivity following passage through the gastrointestinal tract of seagulls

and chickens that have been fed infected prawns (Vanpatten et al. 2004), therefore indicating this agent can

be spread via birds and probably other mechanical vectors. Like HPV, the simple, single-stranded, non-

enveloped virion structure and DNA-based genome confers IHHNV a much greater environmental resilience

compared to some other lipid-containing enveloped viruses such as WSSV and YHV (Biosecurity Australia

2009).

Water temperature is likely to play an important role in transmission of the disease, with little information

available on transmission rates of IHHNV at lower water temperatures typical of southern States. The

replication rate of IHHNV at high water temperatures is significantly reduced in P. vannamei held at 32°C

compared to prawns held at 24°C. Prawns held at 32°C had approximately 102 lower viral load than prawns

held at 24°C (Montgomery Brock et al. 2007). Wild prawns commonly carry sub-clinical IHHNV

infections, and are unlikely to suffer mortalities unless they were exposed to significant stressors. If a

clinically diseased prawn was used as bait or berley and the susceptible prawn in the index case became

infected and died, it is possible that dead and moribund prawns could be a source of IHHNV, however

predation of moribund prawns by non susceptible species such as fish and crabs may be an important factor

modulating transmission and spread of IHHNV in some index cases. Taking these various factors into

consideration, the risk of exposure and establishment of IHHNV via use of bait and berley remains non-

negligible, and the likelihood of exposure and establishment of IHHNV in new prawn populations via

translocation is considered to be Low.


Although IHHNV is present in populations of wild P. monodon in Australia and other parts of the world,

there is no evidence yet available of it having a discernible impact on P. monodon in the wild. It appears that

the presence of IHHNV in apparently healthy wild P. monodon is a normal occurrence in regions where this

_________________________________________________________________________________________________________________________________________________________________________________


157

www.digsfish.com

virus is endemic (Flegel et al. 2004, Krabsetsve et al. 2004, Withyachumnarnkul et al. 2006). Only in areas

where the highly susceptible P. stylirostris occurs in the wild have adverse consequences of IHHNV

introduction into populations of wild prawns been observed, including a transient 50% reduction in landings

of P. stylirostris in the Gulf of California prawn fishery after introduction of IHHNV (Morales-Covarrubias

et al. 1999), though various other factors may have also contributed to the decline. IHHNV has certainly

been associated with disease in cultured prawns, and hence its introduction into new areas would likely have

some consequences for prawn aquaculture industries in the affected region. Although IHHNV is not likely

to cause a severe epizootic in larger cultured prawns, it may still cause economic loss which would have a

detrimental effect on the profitability and hence viability of the prawn aquaculture industry in Australia.

IHHNV is listed by the OIE and NACA, and it is listed as a reportable disease in all States (Table 3b).

Hence the spread of IHHNV to new areas is likely to adversely impact trade. Considering all of these

factors, establishment of IHHNV in new areas would have mild to moderate consequences for prawn

aquaculture, that would probably be amenable to control, but it would appear unlikely to cause noticeable

environmental effects in Australia. It is therefore estimated that the consequences of introduction of IHHNV

into different parts of the Australian environment via use of infected bait would likely be Low.


The unrestricted risk associated with transfer of IHHNV via infected prawns is determined by combining the

likelihood of entry and exposure (from Table 5) with the consequences of establishment (Table 7). The

unrestricted risk estimate for IHHNV does not exceed the ALOP for any of the commodity types, suggesting

that additional risk management for this disease agent is not required at this time.

Risk estimate for infection of prawns with IHHNV

Commodity type Live

Prawns

Whole fresh

dead prawns

Frozen whole

prawns

Frozen

prawn tails

Frozen prawn

heads

Combined likelihood

of release and

exposure

Low Low Low Very Low Low

Consequences of

establishment

Low Low Low Low Low


risk

Very low risk Very low risk Very low

risk

Very low risk

_________________________________________________________________________________________________________________________________________________________________________________


158

www.digsfish.com

5.22 Infection of prawns with Monodon Baculovirus (MBV)

5.22.1 Aetiologic agent: Penaeus monodon baculovirus (MBV). MBV is an occluded double-stranded

DNA enteric baculovirus that is tentatively classified as a species in the genus Nucleopolyhedrovirus.

Plebejus baculovirus and Bennettae baculovirus (Spann and Lester 1996) and several other similar viruses

exist, which are considered to be strains of MBV and they will be considered together with MBV.


5.22.3 Australias status: Reported in QLD, NSW, WA and SA, and is a reportable disease in WA and the

NT (Table 3b).

5.22.4 Epidemiology

MBV was first discovered during an epizootic of laboratory reared, adult P. monodon imported from Taiwan

into Mexico and maintained in quarantine (Lightner and Redman 1981, Lightner et al. 1983c). Subsequently

MBV was reported from numerous other regions, including Malaysia (Anderson et al. 1987), Indonesia

(Nash et al. 1988), and Australia (Lester et al. 1987, Doubrovsky et al. 1988, Spann and Lester 1996, 1997),

and over time it was shown to be widely distributed in cultured prawns in China, Philippines, Thailand, Sri

Lanka, Singapore, India, Israel, Japan, Kuwait, Oman, Italy, Kenya, Gambia and South Africa. MBV has

also been introduced into Tahiti, Hawaii, Brazil, Ecuador, Mexico, Puerto Rico and in some parts of the USA

with movements of cultured prawns. Various strains of the virus infect a variety of penaeid hosts, including

not only P. monodon, but also P. merguiensis, P. semisulcatus, P. kerathurus, P. vannamei, P. esculentus, P.

penicillatus, P. plebejus, Metapenaeus ensis, M. monoceros, M. elegans, M. bennettae and Melicertus

latisulcatus (Lightner et al. 1987, Lightner 1996, Spann and Lester 1996, Manivannan et al. 2004, Roberts et

al. 2010), and also Macrobrachium rosenbergii in Thailand (Gangnonngiw et al. 2010).

Affected prawns display lethargy, anorexia, darkened colouration, and heavy surface fouling. Acute MBV

causes disintegration of the hepatopancreatic tubules and midgut epithelia and consequently, dysfunction of

these organs often followed by secondary bacterial infections (Flegel 2006). MBV has been linked with high

mortalities (over 90%) in late postlarvae and juvenile cultured prawns in many countries, and is considered

by some observers to be partially responsible for the collapse of the prawn culture industry in Taiwan in the

late 1980s (Lin 1989). Usually late juvenile and adult P. monodon are more resistant to MBV than larval,

postlarval, and early juvenile stages prawns (Brock and Lightner 1990). Although good culture practices may

enhance survival of MBV infected stocks (Fegan et al. 1991, Flegel 1997), growth, crop value and

performance may be significantly reduced (Flegel et al. 2004, Flegel 2006) and MBV may predispose

infected prawns to infections by other pathogens, with corresponding higher mortality rates.

MBV infections are characterised by the presence of prominent single or, more often, multiple spherical

intranuclear occlusion bodies in hypertrophied epithelial cells of the hepatopancreas and midgut, or free

within lysed cell debris in the faeces (Lightner and Redman 1981, Flegel 2006). MBV produces a spherical

occlusion body in which virions are embedded in a crystalline protein matrix (Lightner and Redman 1981).

Crowding or environmental stress may increase the prevalence and intensity of MBV (Lightner et al. 1983c,

_________________________________________________________________________________________________________________________________________________________________________________


159

www.digsfish.com

Fegan et al. 1991, Lightner 1996, Flegel 1997). Transmission of MBV appears to be exclusively by the oral

route in which cannibalism and faecal-oral contamination are the principal mechanisms (Lightner et al.

1983c, Paynter et al. 1992, Lightner 1996, Spann and Lester 1996).

PCR based surveys showed that prevalence of MBV in wild P. monodon in the Philippines was 18-20% in

the dry season and 6 to 9% in the wet season in male and female prawns, respectively (De La Pena et al.

2008). In cultured P. monodon the prevalence of MBV can be much higher, for example up to 66.7 % in the

Philippines (see Natividad and Lightner 1992). The high prevalence observed in cultured prawns can be

attributed to culture conditions such as crowding stress and cannibalism that enhances stress-induced MBV

infection (De La Pena et al. 2008). In all samples that were tested by De La Pena et al. (2008), positive

results were obtained only after the nested PCR step. This suggests a low viral load in the samples and

consequently, wild populations of P. monodon in the Philippines can be considered as asymptomatic carriers

responsible for vertical and horizontal transmission of the virus in the hatchery phase (De La Pena et al.

2008). MBV-like viruses are only known to infect prawns.


MBV is known to naturally infect a wide range of prawns, including several species used as bait or berley in

Australia (Table 8), including Penaeus monodon, P. esculentus, P. semisulcatus, P. merguiensis, Melicertus

latisulcatus, Penaeus spp. and Metapenaeus spp.. Infections of wild prawns are usually asymptomatic

(Spann and Lester 1996) and can occur at high prevalences. For example, over 90% of Metapenaeus

bennettae were infected by MBV in Moreton Bay (Spann and Lester 1997), while a MBV-like virus was

found in 60% of wild M. latisulcatus in SA (Roberts et al. 2010). Some species of prawns may be refractory

to some strains of MBV. For example, P. monodon was reportedly refractory to experimental infection with

the Bennettae baculovirus strain of MBV, while P. esculentus, P. plebejus and M. macleayi caught in the

same trawls as the heavily infected M. bennettae, were apparently not infected when examined using

histopathology (Spann and Lester 1997).

Large quantities of prawns are used as bait throughout Australia (Kewagama Research 2002, 2007). Live

prawns are generally not available commercially, but 11% of recreational fishers catch their own prawns

(Kewagama Research 2007), and some of these could be translocated live. However, the majority of the

volume of prawns distributed as bait and berley throughout the country is frozen whole prawns

(Metapenaeus spp.), while fresh and frozen prawns (Penaeus spp.) are also widely distributed for human

consumption, and around 8% of recreational fishers divert these to use as bait or berley (Kewagama

Research 2007). Both live and fresh, whole, green (uncooked) prawns would be expected to contain viable

virus.

Frozen whole uncooked product would also be expected to contain viable virus, especially if affected prawns

were taken from affected aquaculture ponds, and MBV is known to retain virulence when thawed after

storage at -70°C for extended periods (Paynter et al. 1992, Spann et al. 1993, Spann and Lester 1996,

Manisseri et al. 1999). The ability of the virus to survive freezing may be due to the fact that MBV virions

encapsulated in crystalline occlusion bodies are well protected from inactivation by a range of chemical and

physical insults (Spann et al. 1993, Manisseri et al. 1999). While survival and viability of MBV upon

thawing from normal commercial freezing temperatures of -18°C is not documented in the literature, it

_________________________________________________________________________________________________________________________________________________________________________________


160

www.digsfish.com

would be reasonable to expect that a large percentage of these viruses could remain viable after at least one

freeze-thaw cycle during commercial freezing, storage and transport within Australia.

MBV infects the hepatopancreas and midgut almost exclusively (Johnson and Lightner 1988). Hence

removal of the head section containing the hepatopancreas could result in a marked reduction in the viral

load of infected prawns. Taking into account the quantities of prawn products used as bait or berley (Table

1), and the high prevalence of the disease agent, the likelihood estimations for the occurrence of MBV in

these commodities are listed below.

Release assessment for infection of prawns with MBV

Commodity

type

Live

Prawns

Whole fresh

dead prawns

Frozen

whole

prawns

Frozen

prawn

tails

Frozen

prawn

heads

Likelihood of

release

High High Moderate Very low Moderate


Prawns in many areas of Australia are already at risk of exposure to MBV. However, translocation of MBV

infected prawns from areas where these viruses are enzootic via their use as bait or berley could transport

MBV to new regions. Infection and establishment of MBV in new hosts would occur only if viable viral


conditions for transmission. The virus has a wide host range, but only infects prawns and does not infect

other crustaceans. The protection afforded to MBV by the crystalline occlusion body results in the virus

being very resistant to inactivation by chemical and physical means (Spann et al. 1993), meaning that it

could remain infective in the environment for an extended period. Susceptible crustaceans can become

infected with MBV via per-os exposure through cannibalism and faecal-oral contamination (Lightner et al.

1983c, Paynter et al. 1992, Lightner 1996, Spann and Lester 1996). Water temperature is likely to play an

important role in transmission of the disease, with little information available on transmission rates of MBV

at lower water temperatures typical of southern States.

The identity of all species of prawns susceptible to MBV is not fully known, though some species appear

refractory to infection by some strains of MBV (Spann and Lester 1996). Nevertheless, susceptible species

of prawns feeding on clinically diseased prawns used as bait or berley may receive sufficient dose of the

virus to become infected, however it appears unlikely the MBV can be transmitted via the per-os route if

sub-clinically affected crustaceans were used as bait (Spann and Lester 1996). If susceptible species of wild

penaeids in Australia received an infective dose of MBV from a clinically diseased prawn used as bait or

berley, and an index case occurred, the virus would be likely to persist in the population of susceptible

penaeids, as infected prawns would be unlikely to suffer disease and/or mortalities unless they were exposed

_________________________________________________________________________________________________________________________________________________________________________________


161

www.digsfish.com

to significant stressors. Nevertheless, predation of infected prawns by non susceptible species such as fish

and crabs may be an important factor modulating transmission and spread of MBV in some index cases.

Whether the virus could be transferred to progeny and become established is less clear, given that vertical

transmission has not been recorded for MBV. However the numerous reports of MBV introductions around

the world (Lightner 1996) suggests that there would be reasonable chances of establishment if it were

introduced. Taking these various factors into consideration, the risk of exposure and establishment of MBV

via use of bait and berley remains non-negligible, and the likelihood of exposure and establishment of MBV

in new prawn populations via translocation is considered to be Low.


Several strains of MBV are already known to be present in the Australian environment. Although MBV is

present in populations of wild penaeids in many parts of the world, there is no evidence of it having a

discernible impact in the wild. It appears that the presence of MBV in apparently healthy wild prawns is a

normal occurrence in regions where this virus is endemic. MBV has been associated with limited disease in

cultured prawns, but it is relatively easy to prevent infection in hatcheries by taking steps to eliminate faecal

contamination of spawned eggs and larvae by washing nauplii or eggs with formalin, iodophores and clean

sea water (Chen et al. 1990). Equipment can also be effectively disinfected using chlorine (Spann et al.

1993) and iodophores (Manissiri et al. 1999). Thus using simple hygiene practices, MBV can be controlled

and the consequences of establishment in farmed prawns can therefore be largely mitigated with minimal

detrimental effect on the profitability and hence viability of the prawn aquaculture industry. MBV is not

listed by the OIE and NACA, but is listed as a reportable disease in the NT and WA (Table 3b). Hence the

spread of MBV to new areas is likely to have minimal impacts on trade. Considering all of these factors,

establishment of MBV in new areas would have mild consequences for prawn aquaculture, that would be

amenable to control, and it would appear unlikely to cause noticeable environmental effects. It is therefore

estimated that the consequences of introduction of MBV into different parts of the Australian environment

via use of infected bait would likely be Very low.


The unrestricted risk associated with MBV is determined by combining the likelihood of entry and exposure

(from Table 5) with the consequences of establishment (Table 7). The unrestricted risk estimate for MBV

does not exceed the ALOP for any of the commodity types, suggesting that additional risk management for

this disease agent is not required at this time.

_________________________________________________________________________________________________________________________________________________________________________________


162

www.digsfish.com

Risk estimate for infection of prawns with MBV

Commodity type Live

Prawns

Whole fresh

dead prawns

Frozen whole

prawns

Frozen

prawn tails

Frozen prawn

heads

Combined likelihood

of release and

exposure

Low Low Low Very low Low

Consequences of

establishment

Very low Very low Very low Very low Very low


risk

Negligible risk Negligible risk Negligible

risk

Negligible risk

_________________________________________________________________________________________________________________________________________________________________________________


163

www.digsfish.com

5.23 Infection of prawns with Mourilyan virus (MoV)

5.23.1 Aetiologic agent: Mourilyan virus (MoV) is an enveloped, spherical to ovoid (85–100 nm diameter)

viral particle with a RNA genome that is genetically related to the Uukuniemi virus and other phleboviruses

of the Bunyaviridae (see Cowley et al. 2005).


5.23.3 Australias status: MOV infections have been reported from QLD, and this disease is not reportable

in any State (Table 3b).

5.23.4 Epidemiology

Mourilyan virus (MoV) was originally described from P. monodon from eastern Australia during studies of

GAV (Cowley et al. 2005), but has since been recorded in wild and cultured P. monodon, P. japonicus and

P. merguiensis from Australia, Fiji, Malaysia, Thailand and Vietnam (OIE 2007a). In P. monodon, MoV can

exist as a chronic or acute infection and affects tissues of ectodermal and mesodermal origin, with formation

of lymphoid organ spheroids (Cowley et al. 2005). MoV often occurs in mixed infections with GAV in P.

monodon along Australias east coast, and although MoV is not highly pathogenic in cultured P. monodon

(see Oanh et al. 2011), there is some evidence associating MoV with mortalities in cultured P. japonicus (see

Sellars et al. 2006, Cowley et al. 2009). It is thought that vertical transmission of these viruses by P.

monodon broodstock captured from these areas is the likely mechanism perpetuating their high prevalences

in the wild and in P. monodon farmed along eastern Australia (Cowley et al. 2009). Cowley et al. (2005)

found the prevalence of MoV in P. monodon broodstock captured from the east coast of Cape York

Peninsula, north Queensland and farmed and domesticated stocks generated from these broodstock to be

(98%) (60 of 61 prawns tested positive). In contrast, surveys of P. monodon taken from Weipa on the west

coast of Cape York Peninsula found MoV was detected at very low levels and a maximum of only 15%

prevalence using qPCR (see Cowley et al. 2009). Furthermore, as one of nine mantle samples from pearl

oysters collected at Weipa was also qPCR-positive for MoV, the significance of the low prevalence of very

low-level MoV infection in wild P. monodon from the area is not clear as it is possible that MoV was

detected in the absence of an infection having established (Cowley et al. 2009).

Minor nucleotide sequence variations (< 5%) occur between MoV isolates from Australia, Malaysia and

Thailand, indicating that strain variants exist in divergent populations of P. monodon (OIE 2007a). No

significant sequence variation has been detected between virus isolates infecting eastern Australian P.

monodon and P. japonicus, or among P. monodon sampled from various locations in north and eastern

Australia and in Fiji, suggesting a single genetic lineage might exists in these prawn populations (OIE

2007a). A study of captive breeding of P. japonicus demonstrated that broodstock reared from egg to

maturity in controlled environment tanks had significantly higher survival rates than sibling broodstock

sourced from farm ponds (Sellars et al. 2006). The low prevalence of MoV in the tank-reared stocks was

associated with high mean survival (±S.E.) (75.81±7.12%). Conversely, the high prevalence of MoV in the

pond-reared stocks was associated with low mean survival (11.29 ± 1.42%). The results indicated that P.

japonicus broodstock reared in controlled environment systems are less at risk from MoV, due to lower

_________________________________________________________________________________________________________________________________________________________________________________


164

www.digsfish.com

infection rates or a greater capacity to tolerate infection, compared to prawns sourced from farm ponds

(Sellars et al. 2006). Rajendran et al. (2006) experimentally transmitted MoV to healthy juvenile P.

japonicus using an inoculum prepared from diseased P. monodon by snap freezing tissues on dry ice after

collection and storing at −80°C until required. Horizontal transmission via injection has been shown and

infection is also likely via ingestion of infected tissue, whilst vertical transmission has not been reported, but

cannot be excluded (OIE 2007a). MoV has only been recorded from prawns and not other crustaceans.


MoV is known to naturally infect wild P. monodon in Queensland in asymptomatic infections that can occur

at prevalences near 100% in some regions (Cowley et al. 2009). The virus has also been recorded in cultured

P. monodon, and P. japonicus (see Sellars et al. 2006) and was also detected in P. merguiensis but a

productive infection state was not suspected (OIE 2007a). Large quantities of prawns are used as bait

throughout Australia (Kewagama Research 2002, 2007). Live prawns are generally not available

commercially, but 11% of recreational fishers catch their own prawns (Kewagama Research 2007), and some

of these could be translocated live. However, the majority of the volume of prawns distributed as bait and

berley throughout the country is frozen whole prawns (mainly Metapenaeus spp.), while fresh and frozen


fishers divert these to use as bait or berley (Kewagama Research 2007). Infected fresh, green (uncooked)

prawns would still be expected to contain viable virus, and frozen whole uncooked product would also

contain viable virus if infected prawns were taken from aquaculture ponds, as it is known that MoV survives

freezing (-80°C) and remained infective in experimental trials (Rajendran et al. 2006). While survival and

viability of MoV upon thawing from normal commercial freezing temperatures of -18°C is not documented,

it would be reasonable to expect that a large percentage of these viruses could survive and remain viable

after at least one freeze-thaw cycle during commercial freezing, storage and transport within Australia.

During disease outbreaks in cultured prawns along Australias east coast, MoV was widely distributed

throughout cephalothoracic tissues of mesodermal and ectodermal origin. Heavily infected tissues included

lymphoid organ spheroids and tubules, gill and cuticular epithelium, particularly in the foregut and

cephalothorax, organ connective tissues and glial, neurosecretory and giant cells in the segmental nerve

ganglia (Cowley et al. 2005). Hence removal of the head section containing the lymphoid organ, gills, and

foregut could result in a marked reduction in the viral load of infected prawns. Taking into account the

quantities of prawns used as bait or berley (Table 1), and the high prevalence of the disease agent, the

likelihood estimations for the occurrence of MoV in these commodities are listed below.

Release assessment for infection of prawns with MoV

Commodity

type

Live

Prawns

Whole fresh

dead prawns

Frozen

whole

prawns

Frozen

prawn

tails

Frozen

prawn

heads

Likelihood of

release

High High Moderate Very low Moderate

_________________________________________________________________________________________________________________________________________________________________________________


165

www.digsfish.com


Prawns in QLD are already at risk of exposure to MoV. However, translocation of MoV infected prawns

from areas where MoV is enzootic via their use as bait could transport the virus to new regions. Infection

and establishment of MoV in new hosts would occur only if viable viral particles were introduced into an

area where susceptible prawns were present under suitable environmental conditions for transmission. Little

is known about the natural host range and transmission of MoV, but to date it has only infected prawns and

does not appear to infect other crustaceans. Susceptible wild prawns are probably able to become infected

with MoV via per-os exposure (OIE 2007a), however to date this has not been reported experimentally.

Viral loads in healthy P. monodon infected with MoV were high, ranging from 2 x 105 to 2 x 109 RNA copies

/ng (Oanh et al. 2011). While injection of prawns with a mixed GAV/MoV inoculum containing 3 x 106

RNA copies /ng MoV resulted in an increase in MoV viral load to 2.5 x 108 RNA copies /ng after 9 days, the

death of the experimental prawns was related to increases in viral loads of GAV, and not MoV (Oanh et al.

2011).

It is possible, therefore, that P. monodon, P. japonicus, P. merguiensis and other prawn species that form the

basis of important recreational and commercial fisheries in other regions of Australia besides QLD (e.g.

Melicertus latisulcatus, P. plejebus) may be susceptible to infection with MoV if they were to come in

contact with the virus. Infected P. monodon can tolerate a high level of MoV infection without becoming

diseased (Oanh et al. 2011). However, water temperature is likely to play an important role in transmission

of the disease, with little information available on transmission rates of MoV at lower water temperatures

typical of southern States, and predation of moribund prawns by non susceptible species such as fish and

crabs may be an important factor modulating transmission and spread of MoV in some index cases. Taking

these various factors into consideration, the risk of exposure and establishment of MoV via use of bait and

berley remains non-negligible, and the likelihood of exposure and establishment of MoV in new prawn

populations via translocation is considered to be Low.


MoV is already known to be present in the Australian environment off the QLD coast. Although MoV is

present in populations of wild P. monodon, there is no evidence of it having a discernible impact in the wild.

Healthy P. monodon infected with MoV can carry very high viral loads (Oanh et al. 2011), hence it appears

that the presence of MoV in both wild and cultured P. monodon is not associated with disease, and that other

viruses such as GAV are more significant in instances where MoV has been associated with reduced survival

in P. japonicus cultured under certain conditions (Sellars et al. 2006). MoV is not listed by the OIE and

NACA, and is also not listed as a reportable disease in any States (Table 3b). Hence the spread of MoV to

new areas is not likely to impact trade. Considering all of these factors, establishment of MoV in new areas

would have mild consequences for prawn aquaculture, that would probably be amenable to control, and it

would appear unlikely to cause noticeable environmental effects. It is therefore estimated that the

consequences of introduction of MoV into different parts of the Australian environment via use of infected

bait would likely be Very low.

_________________________________________________________________________________________________________________________________________________________________________________


166

www.digsfish.com


The unrestricted risk associated with MoV is determined by combining the likelihood of entry and exposure

(from Table 5) with the consequences of establishment (Table 7). The unrestricted risk estimate for MoV

does/does not exceed the ALOP for any of the commodity types, suggesting that additional risk management

for this disease agent is not required at this time.

Risk estimate for infection of prawns with MoV

Commodity type Live

Prawns

Whole fresh

dead prawns

Frozen whole

prawns

Frozen

prawn tails

Frozen prawn

heads

Combined likelihood

of release and

exposure

Low Low Low Very low Low

Consequences of

establishment



risk


risk

Negligible risk

_________________________________________________________________________________________________________________________________________________________________________________


167

www.digsfish.com

5.24 Infection of prawns and crayfish with Spawner Isolated Mortality Virus (SMV)

5.24.1 Aetiologic agent: Spawner Isolated Mortality Virus (SMV) is a parvo-like DNA virus with

icosahedral virions 20 to 25 nm in diameter (Owens et al. 1998).


5.24.3 Australias status: SMV infections have been reported from QLD, and this disease is reportable only

in South Australia (Table 3b).

5.24.4 Epidemiology

In 1993, an epizootic occurred in captive Penaeus monodon spawners at a research facility in northern

Queensland, Australia. The spawners exhibited lethargy, failure to feed, redness of the carapace and

pleopods, and an increased mortality rate (Fraser and Owens 1996). Inoculation of healthy P. monodon with

cell free extracts of infected tissue filtered to 0.45 µm produced mortalities approaching 100%. Infected

prawns began to show signs of disease (becoming a dark red colour) by Day 6, produced red faeces by Day

10 and the first mortalities were observed by Day 13. Red faeces were a feature of this disease which had

not been previously reported (Fraser and Owens 1996). Investigations discovered a parvo-like virus with

icosahedral virions 20 to 25 nm in diameter inside affected cells, and the new virus was called spawner

isolated mortality virus (Owens et al. 1998).

Histopathology demonstrated pathological changes in the subcuticular epithelium and underlying muscle,

haematopoietic tissue, lymphoid organ, hepatopancreas and gut. There were extensive areas of haemocytic

infiltration and melanisation of the subcuticular epithelium with haemocytic replacement and necrosis of the

underlying muscle. In advanced cases there were also large areas of necrosis in the hepatopancreas with

pyknotic cells sloughing into the lumen (Fraser and Owens 1996).

In situ hybridisation of P. monodon demonstrated that SMV mainly infected the gastrointestinal tract (Owens

et al. 1998). Target organs included the hepatopancreas, midgut and to a lesser extent, the hindgut caecae.

In heavy infections, the virus would break through the lamina propria of the gut and become systemic,

localising in the lymphoid organ, gonads and heart, and permitting egress of the virus through the gut, voided

with the faeces (Owens et al. 1998). The disease could be transmitted to healthy prawns by intra-muscular

injection of filtrates from infected prawns, or by feeding infected prawn tissue (Fraser and Owens 1996).

Deaths began to occur within 14 days for prawns injected with filtrates and 30 days post infection in the

prawns exposed per-os via feed (Fraser and Owens 1996).

When the first outbreaks of mid-crop mortality syndrome (MCMS) occurred in Australian prawn farms in

1994, SMV was present (Owens et al. 1998). Further investigations found that MCMS was associated with

mixed infections, with GAV being mainly involved, together with SMV. Subsequent investigations focused

on determining the origins of SMV within the industry (Owens et al. 2003). Previous observations suggested

that SMV was vertically transmitted through hatcheries. Owens et al. (2003) surveyed 909 P. monodon and

P. merguiensis spawners from hatcheries and found 77 of the 909 examined (8.47%) tested positive for

_________________________________________________________________________________________________________________________________________________________________________________


168

www.digsfish.com

SMV. However, there was significant variation in the prevalence (0- 24%) over the years and between

seasons and species (Owens et al. 2003). Survival in hatchery tanks of PLs from SMV-positive spawners

was lower than those from SMV-negative spawners, supporting the hypothesis that SMV is vertically

transmitted from spawners to postlarvae in hatcheries and causes reduced survival of progeny (Owens et al.

2003). In later stages of growout in hatchery pools, nursery and grow-out ponds, however, progeny from

SMV-positive spawners sometimes had better survival rates than controls (Owens et al. 2003). SMV has

only been recorded from prawns and freshwater crayfish and not other crustaceans.


SMV is known to infect a number of hosts in Australia, including several species used as bait or berley, not

only P. monodon and P. merguiensis, but also P. esculentus, P. japonicus and Metapenaeus ensis (see

Owens et al. 1998). Owens and McElnae (2000) also demonstrated that freshwater crayfish (Cherax

quadricarinatus) displaying reduced tolerance of stress and reduced growth were suffering from mixed SMV

and other viral infections. Infections of SMV in wild caught broodstock occurred at prevalences of 24% in

P. monodon and 4% in P. merguiensis (Owens et al. 2003). Large quantities of prawns are used as bait


commercially, but 11% of recreational fishers catch their own prawns (Kewagama Research 2007), and some

of these could be translocated live. However, the majority of the volume of prawns distributed as bait and

berley throughout the country is frozen whole prawns (mainly Metapenaeus spp.), while fresh and frozen


fishers divert these to use as bait or berley (Kewagama Research 2007). Infected fresh, whole, green

(uncooked) prawns would still be expected to contain viable virus. Frozen whole prawns would also be

expected to contain viable virus, as it is known that SMV survives freezing (-20°C) and remained infective in

subsequent experimental trials (Owens et al. 1998). It is therefore likely that a high percentage of these

viruses could survive and remain viable after at least one freeze-thaw cycle during commercial freezing,

storage and transport within Australia.

SMV infects mainly the gastrointestinal tract of prawns (Owens et al. 1998), hence removal of the head

section containing the hepatopancreas, gills, lymphoid organ, and foregut would most likely result in a

marked reduction in the viral load of infected prawns. Taking into account the prevalence of the disease

agent and the quantities of prawns used as bait or berley (Table 1), the likelihood estimations for the

occurrence of SMV in these commodities are listed below.

Release assessment for infection of prawns with SMV

Commodity

type

Live

Prawns

Whole fresh

dead prawns

Frozen

whole

prawns

Frozen

prawn

tails

Frozen

prawn

heads

Likelihood of

release

High High Low Very low Low

_________________________________________________________________________________________________________________________________________________________________________________


169

www.digsfish.com


Prawns and crayfish throughout northern Australia are already at risk of exposure to SMV. However,

translocation of SMV infected prawns from areas where SMV is enzootic via their use as bait could transport

the virus to new regions. Infection and establishment of SMV in new hosts would occur only if viable viral


conditions for transmission. The virus has a host range that includes 4 species of prawns as well as a

freshwater crayfish. Other prawn species that form the basis of important recreational and commercial

fisheries in southern Australia (e.g. Melicertus latisulcatus, P. plejebus) as well as rare or threatened

freshwater crayfish may be susceptible to infection with SMV if they were to come in contact with the virus.

Transmission of SMV via feeding on infected tissue has been recorded (Fraser and Owens 1996) with deaths

beginning to occur within 30 days in prawns exposed per-os via feed, hence susceptible wild crustaceans

may become infected with SMV via horizontal routes such as per-os exposure, as well as horizontally via the

water. Susceptible species of prawns and freshwater crayfish feeding on clinically diseased prawns used as

bait or berley may receive sufficient dose of the virus to become infected, however it is not known whether

SMV could be transmitted via the per-os route if subclinically affected crustaceans were used as bait or

berley. It is possible, therefore, that P. monodon, P. merguiensis, P. japonicus and Metapenaeus spp. that

form the basis of important recreational and commercial fisheries in other regions of Australia besides QLD

(e.g. Melicertus latisulcatus, P. plejebus), as well as other species of freshwater crayfish (including

threatened species), may be susceptible to infection with MoV if they were to come in contact with the virus

If susceptible species of wild prawns or crayfish in Australia received an infective dose of SMV and an index

case occurred, the virus would be likely to persist in the population of susceptible crustaceans, as they would

be unlikely to suffer mortalities unless they were exposed to significant stressors. The virus would then be

likely to be transferred to progeny via vertical transmission and could become established. However, water

temperature is likely to play an important role in transmission of the disease, with little information available

on transmission rates of SMV at lower water temperatures typical of southern States. Predation of SMV

infected crustaceans by non susceptible species such as fish and crabs may be an important factor modulating

transmission and spread of SMV in some index cases. Taking these various factors into consideration, the

risk of exposure and establishment of SMV via use of bait and berley remains non-negligible, and the

likelihood of exposure and establishment of SMV in populations of prawns and crayfish via translocation is



SMV is already present in populations of wild prawns in some regions of Australia, and there is no evidence

of a discernible impact in the wild. It appears that the presence of SMV in healthy wild penaeids is a normal

occurrence in regions where this virus is endemic (Owens et al. 1998). However, SMV has been associated

with poor performance and reduced survival in cultured P. monodon (see Fraser and Owens 1996, Owens et

al. 2008), as well as mortalities of 60 to 100% in cultured freshwater crayfish (Owens and McElnae 2000),

hence its introduction into new areas would likely have some consequences for prawn aquaculture and

possibly significant adverse consequences for freshwater crayfish aquaculture in the affected region. SMV

has potential to have a detrimental effect on the profitability and hence viability of the prawn and freshwater

_________________________________________________________________________________________________________________________________________________________________________________


170

www.digsfish.com

crayfish aquaculture industries in Australia, as well as potentially negative ramifications for conservation of

threatened crayfish species. SMV is not listed by the OIE and NACA, but it is listed as a reportable disease

in SA (Table 3b). Hence the spread of SMV to new areas could impact trade to a limited extent.

Considering all of these factors, establishment of SMV in new areas would have mild (prawns) to moderate

(freshwater crayfish) consequences for prawn aquaculture, that would probably be amenable to control, but it

would appear unlikely to cause noticeable environmental effects to prawn populations, and uncertain effects

on wild populations of freshwater crayfish. It is therefore estimated that the consequences of introduction of

SMV into different parts of the Australian environment via use of infected bait would likely be Moderate.


The unrestricted risk associated with SMV is determined by combining the likelihood of entry and exposure

(from Table 5) with the consequences of establishment (Table 7). The unrestricted risk estimate for SMV

exceeds the ALOP for live prawns and whole fresh dead prawns, suggesting that additional risk management

for this disease agent is required for these commodities.

Risk estimate for infection of prawns with SMV

Commodity type Live

Prawns

Whole fresh

dead prawns

Frozen whole

prawns

Frozen

prawn tails

Frozen prawn

heads

Combined likelihood

of release and

exposure

Low Low Very low Very low Very low

Consequences of

establishment



risk

Very low risk

_________________________________________________________________________________________________________________________________________________________________________________


171

www.digsfish.com

5.25 White Tail Disease of freshwater giant prawns

5.25.1 Aetiologic agent: Macrobrachium rosenbergii Nodavirus (MrNV), a non-enveloped double stranded

RNA virus and its associated satellite extra small virus (XSV), a single stranded RNA virus (Bonami et al.

2005).


5.25.3 Australias status: MrNV infections have been reported from QLD, and this disease is reportable in

QLD, NSW and Victoria (Table 3b).

5.25.4 Epidemiology

Macrobrachium rosenbergii nodavirus (MrNV) and its associated extra small virus (XSV) cause white tail

disease (WTD) in freshwater giant prawns Macrobrachium rosenbergii (see Bonami and Widada 2003,

Bonami et al. 2005). The disease was first recognised in cultured post larvae of M. rosenbergii in Taiwan in

1992 which were affected by an epizootic disease characterised by white opaque areas in the abdominal

muscle (Tung et al. 1999). Then in 1999 a similar disease was reported in M. rosenbergii reared in

Guadeloupe, French West Indies (Arcier et al. 1999), followed by China (Qian et al. 2003), India (Vijayan et

al. 2003, 2005b, Sahul Hameed et al. 2004), Thailand (Yoganandhan et al. 2006) and in wild caught adult M.

rosenbergii in Australia (Owens et al. 2009).

The disease occurs mainly in postlarval M. rosenbergii, with adults usually not being affected (Sahul

Hameed et al. 2004, Ravi et al. 2010), however subclinically infected adult freshwater prawns can transmit

the virus to their progeny (Sudhakaran et al. 2007). In post-larvae, the clinical signs of WTD include

lethargy, anorexia and opaqueness of the abdominal muscle, giving rise to the name white tail disease. The

opaqueness gradually extends on both sides and leads to necrosis and degeneration of telson and uropods, in

severe cases, followed by heavy mortalities within 2-3 days reaching 99 to 100% within 10 days (Vijayan et

al. 2005b). Unlike idiopathic muscle necrosis, areas of necrotic muscle in WTD affected post larval M.

rosenbergii exhibit basophilic cytoplasmic inclusion bodies (Arcier et al. 1999, Tung et al. 1999) associated

with the presence of MrNV and XSV, but subclinically infected adults may not (Owens et al. 2009).

Transmission of these viruses occurs horizontally through the water, as well as via per-os exposure and

vertically (Sudahakaran et al. 2007, Owens et al. 2009) and transfer of the disease was documented to have

occurred with the movement of infected postlarval M. rosenbergii from Guadeloupe to Puerto Rico (OIE

2005). Indeed, the rapid emergence of the disease in regions of China, Bangladesh and India inhabited by

the western form of M. rosenbergii also suggests that it was introduced into these areas. The Australian

isolate appears to be slightly different, and being detected in wild caught broodstock, this suggests the

disease may be endemic to this country in the eastern form of M. rosenbergii (see Owens et al. 2009). Given

that the Australian isolate of MrNV is genetically most closely related to a Chinese isolate, this may suggest

the disease may have been introduced into Australia at some time in the past by human activities (Owens et

al. 2009).

_________________________________________________________________________________________________________________________________________________________________________________


172

www.digsfish.com


MrNV is known to naturally infect wild M. rosenbergii (Table 8) in north QLD in asymptomatic infections

(Owens et al. 2009). The virus is only known to naturally infect M. rosenbergii, although prawns, aquatic

insects and Artemia may act as carriers of the disease agent (Sudhakaran et al. 2006, 2007, 2008). Large

quantities of prawns are used as bait throughout Australia (Kewagama Research 2002, 2007), but these are

almost entirely penaeids and not M. rosenbergii. Live prawns are generally not available commercially, but

11% of recreational fishers catch their own prawns (Kewagama Research 2007), and it is known that M.

rosenbergii is a popular live bait used by recreational fishers in the northern parts of Australia (Ross 1995).

Since M. rosenbergii is a relatively hardy species (Ross 1995), it may be easily translocated live by fishers at

least short distances. Infected fresh, whole, green (uncooked) freshwater prawns would also still be expected

to contain viable virus, but there appears to be very limited published data available examining whether

MrNV and XSV can survive freezing and thawing and remain viable after transport within Australia.

MrNV and XSV occur in the gills, head muscle, stomach, intestine, heart, haemolymph, pleopods, ovaries

and tail muscle of experimentally injected adult freshwater prawns (Sauhl Hameed et al. 2004). Because of

this, removal of the head section alone would not necessarily result in a marked reduction in the viral load of

infected freshwater prawns. Taking into account the relatively small quantities of freshwater prawns used as

bait or berley, as well as the fact that the disease agent may be reasonably prevalent in wild populations of

freshwater prawns in at least some parts of the country (Owens et al. 2009), the likelihood estimations for the

occurrence of MrNV and XSV in these commodities are listed below.

Release assessment for white tail disease of giant freshwater prawns

Commodity

type

Live

freshwater

prawns

Whole fresh

dead

freshwater

prawns

Frozen

whole

freshwater

prawns

Frozen

freshwater

prawn

tails

Frozen

freshwater

prawn

heads

Likelihood of

release

High Moderate Low Low Low


Freshwater prawns throughout at least some parts of northern Australia are already at risk of exposure to

MrNV and XSV. However, translocation of infected prawns from areas where MrNV and XSV are enzootic

via their use as bait could transport the virus to new regions. Infection and establishment of these viruses in

new hosts would occur only if viable viral particles were introduced into an area where susceptible

freshwater prawns were present under suitable environmental conditions for transmission. The virus appears

to cause disease only in freshwater prawns and therefore the risks of other crustaceans becoming diseased

appears negligible, however penaeids and other crustaceans and insects have been identified as carriers.

_________________________________________________________________________________________________________________________________________________________________________________


173

www.digsfish.com

Both post larval, juvenile and adult M. rosenbergii can become infected with MrNV and XSV via horizontal

routes such as per-os exposure, as well as horizontally via the water and vertically (Sudahakaran et al. 2007,

Owens et al. 2009). Freshwater prawns feeding on clinically diseased M. rosenbergii used as bait or berley

have been shown to receive sufficient dose of the viruses to become infected (Owens et al. 2009), and while

these viruses do not generally cause disease in adult M. rosenbergii (unless they are stressed and/or in

confinement, see Owens et al. 2009), infected prawns are likely to survive and pass on the disease to their

progeny via vertical transmission (Sudahakaran et al. 2007). Since the minimum dose required for

successful transmission of these viruses is not yet known, it is difficult to determine whether MrNV and

XSV could be transmitted via the per-os route if subclinically affected crustaceans were used as bait.

However the viruses appear highly virulent for M. rosenbergii, especially post larval stages, and it is very

possible that the minimum infective dose is relatively low. These viruses have also been detected in carriers

such as aquatic insects and Artemia have been shown to be infective when fed to post larval M. rosenbergii

(see Sudhakaran et al. 2007), therefore indicating they can be spread via carriers or mechanical vectors. The

transmission and spread of these viruses in wild populations of M. rosenbergii after an index case occurs will

also likely be modulated by predation of moribund prawns by non susceptible species such as fish and crabs.

Taking these various factors into consideration, the risk of exposure and establishment of MrNV and XSV

via use of bait and berley remains non-negligible, and the likelihood of exposure and establishment of MrNV

and XSV in new prawn populations via translocation is considered to be Low.


Strains of MrNV and XSV are already known to be present in the Australian environment. Although MrNV

and XSV are present in populations of wild M. rosenbergii in many parts of the world, there is no evidence

to date of it having a discernible impact in wild populations. However, the fact that the virus can infect adult

freshwater prawns without causing disease, but can cause mass mortalities in their progeny, suggests that

adverse effects for wild populations of M. rosenbergii cannot be ruled out at this time. MrNV and XSV has

been associated with major disease outbreaks in cultured freshwater prawns, and hence its introduction into

new areas would likely have major consequences for freshwater prawn aquaculture in the affected region.

White tail disease is listed by the OIE and NACA, and it is listed as a reportable disease in QLD, NSW and

Victoria (Table 3b). Hence the spread of MrNV and XSV to new areas is likely to adversely impact trade.

Considering all of these factors, establishment of MrNV and XSV in new areas would have serious

consequences for freshwater prawn aquaculture, and it may also cause noticeable environmental effects. It is

therefore estimated that the consequences of introduction of MrNV and XSV into different parts of the

Australian environment via use of infected bait would likely be Moderate.


The unrestricted risk associated with WTD is determined by combining the likelihood of entry and exposure

(from Table 5) with the consequences of establishment (Table 7). The unrestricted risk estimate for WTD

exceeds the ALOP for live freshwater prawns and whole fresh dead freshwater prawns, suggesting that

additional risk management is required for these disease agents in these commodities.

_________________________________________________________________________________________________________________________________________________________________________________


174

www.digsfish.com

Risk estimate for white tail disease of giant freshwater prawns

Commodity type Live

freshwater

prawns

Whole fresh

dead

freshwater

prawns

Frozen whole

freshwater

prawns

Frozen

freshwater

prawn tails

Frozen

freshwater

prawn heads

Combined likelihood

of release and

exposure


Consequences of

establishment



risk

Very low risk

_________________________________________________________________________________________________________________________________________________________________________________


175

www.digsfish.com

5.26 Infection of crustaceans with rickettsia-like organisms (RLOs)

5.26.1 Aetiologic agent: Rickettsia –like and chlamydia-like organisms that infect crustaceans.


5.26.3 Australias status: Rickettsia –like and chlamydia-like organisms have been reported in all States,

and milky haemolymph disease of spiny lobsters is a reportable disease in QLD, NSW, Tasmania and SA

(Table 3b).

5.26.4 Epidemiology

Rickettsia –like organisms (RLOs) are intracellular prokaryotes that have been reported from a range of

crustaceans (Brock et al. 1986, Brock and Lightner 1990, Bower et al. 1994, 1996, Krol et al. 1991),

including several species known to be used as bait in Australia. They are often present in healthy

crustaceans and in mixed infections with viruses and other agents in diseased prawns (Anderson et al. 1987),

suggesting they are not usually primary pathogens. Because of this, most RLO infections are not considered

significant, however occasionally there are reports of serious disease outbreaks associated with RLOs in

cultured crustaceans including freshwater crayfish (Tan and Owens 2000), lobsters (OIE 2007b, Nunan et al.

2010) and penaeid prawns (Nunan et al. 2003a, 2003b). In Madagascar in 1999 a RLO was associated with

severe mortalities of farmed Penaeus monodon in grow-out ponds (Nunan et al. 2003a). In experimental

trials the RLO involved was able to infect P. vannamei only if it was injected, with infection unable to be

achieved by the oral route (Nunan et al. 2003b). The failure to transfer infection via the oral route is

significant as it suggests that this agent may only cause disease in grow out ponds when hosts are

compromised or stressed, or else it could suggest that a parasite or other aquatic species may be required to

complete the infection process (Nunan et al. 2003b).

In contrast, a “milky haemolymph disease” of spiny lobsters (Panulirus spp.) (MHD-SL) emerged recently

in Vietnam (OIE 2007b, Hung and Tuan 2009, Nunan et al. 2010). The aetiological agent involved with the

mortalities was a novel RLO which occurred in large numbers in haemolymph and muscle, causing large

scale mortalities and losses of production (OIE 2007b, Lightner et al. 2008). Onset of the disease was

reportedly very rapid. Affected lobsters became inactive and ceased feeding. Within another 3-5 days

affected lobsters were observed with milky haemolymph under swollen abdominal pleura of the exoskeleton

(visible on ventral side). Mortality occurred soon after clinical signs become apparent (Lightner et al. 2008).

Mortalities of up to 30% have been attributed to the disease (Tuan and Hung 2009). Haemolymph drawn

with a syringe from affected lobsters will not clot and ranges from slightly cloudy or turbid to milky white.

Dissection of affected lobsters shows the presence of milky coloured haemolymph in the haemocoel and

tissue spaces and white hypertrophied connective tissues (especially serosa and capsules) of all major organs

and tissues (Lightner et al. 2008).

Infection with the RLO Coxiella cheraxi and other RLOs were responsible for mortalities in cultured Cherax

quadricarinatus in north Queensland (Tan and Owens 2000, Edgerton et al. 2002) and Ecuador (Romero et

al. 2000). Very similar diseases, with similar gross and histopathological lesions, have also been reported in

_________________________________________________________________________________________________________________________________________________________________________________


176

www.digsfish.com

farmed Penaeus monodon and in captive European shore crab (Carcinus maenas) (see Lightner et al. 2008,

2009). Sequence information generated from 16 S rDNA amplified from the RLB from infected C. maenas,

P. monodon and Panilurus spp. show that the RLOs in each of these diseases are similar, but not necessarily

closely related (Nunan et al. 2003a, 2003b, Eddy et al. 2007, Nunan et al. 2010).


At this time MHD-SL has only been recorded from spiny lobsters in Vietnam. Hence it will not be

considered further here. However, other RLOs have been reported from a variety of crustaceans in Australia

(Edgerton and Prior 1999, Biosecurity Australia 2009), including species which are used as bait or berley,

such as penaeids and C. quadricarinatus. Large quantities of crustaceans, mainly prawns, are used as bait


commercially, but 11% of recreational fishers catch their own prawns (Kewagama Research 2007), some of

which are likely to be used live and thus could be translocated short distances by fishers. Infected fresh,

whole, green (uncooked) freshwater prawns would also still be expected to contain viable RLOs, and these

bacteria also survive freezing. For example, RLOs in Pandalus sp. survived freezing at -10°C and -15°C for

at least 10 days (Bower et al. 1994, 1996), and a RLO associated with disease in P. monodon survived

freezing at -70°C and remained infective by injection, but not via per-os exposure (Nunan et al. 2003b). It is

likely, therefore, that RLOs will be able to survive freezing and thawing and remain viable after transport

within Australia. Rickettsia like organisms can occur in systemic infections of a wide range of organs

including haemocytes (Romero et al. 2000) and tail muscle (Lightner et al. 2008), hence removal of the

cephalothorax of prawns or lobsters may not necessarily result in a marked reduction in the levels of these

infective agents.

Taking into account the relatively large quantities of crustaceans used as bait or berley, as well as the fact

that these agents may be reasonably prevalent in wild populations of crustaceans in some parts of the country

the likelihood estimations for the occurrence of RLOs in these commodities are listed below.

Release assessment for infection of crustaceans with RLOs

Commodity

type

Live

crustaceans

Whole fresh

dead

crustaceans

Frozen

whole

crustaceans

Frozen

crustacean

tails

Frozen

crustacean

heads

Likelihood of

release

High Moderate Low Low Low


Crustaceans throughout Australia are already at risk of exposure to RLOs that occur naturally in the

Australian environment. Translocation of infected crustaceans containing RLOs via their use as bait could

_________________________________________________________________________________________________________________________________________________________________________________


177

www.digsfish.com

transport these agents into new regions, but infection and establishment in new hosts would occur only if

viable RLOs were introduced into an area where susceptible hosts were present under suitable environmental

conditions for transmission. RLOs are thought to be transmitted horizontally by co-habitation or the per-os

routes (Lightner et al. 2008). Translocation of live crustaceans pose a high risk of introduction and

establishment of RLOs (Romero et al. 2000), but most translocated crustaceans that are used as bait in

Australia are frozen, and it appears that although some RLOs can survive freezing , their infectivity upon

thawing may be reduced. For example, Nunan et al. (2003b) found that upon thawing the a RLO stored at -

70°C, it remained infective only by injection, but not via per-os exposure, which would significantly reduce

the risk of disease transmission in the natural environment.

Wild crustaceans feeding on clinically diseased crustaceans used as bait or berley would likely have a high

chance of becoming infected by RLOs. However, most RLOs appears to cause disease only in crustaceans

reared in confinement, and given that some states do not permit sale of aquacultured products as bait, this

would significantly reduce the likelihood of crustaceans heavily infected with RLOs being used as bait.

Since the minimum dose required for successful transmission of these disease agents is not usually known, it

is difficult to determine whether RLOs could be transmitted via the per-os route if subclinically affected

crustaceans were used as bait. In any case, these disease agents do not generally cause disease unless their

hosts are stressed and/or held in confinement at high densities. The transmission and spread of these disease

agents in populations of wild crustaceans after an index case occurs will also likely be modulated by

predation of moribund crustaceans by non susceptible species such as fish. Taking these various factors into

consideration, the risk of exposure and establishment of RLOs in crustaceans via use of bait and berley

remains non-negligible, and the likelihood of exposure and establishment of RLOs in new crustacean

populations via translocation is considered to be Low.


Several types of RLOs that infect crustaceans are already known to be present in the Australian environment,

and there is not evidence to date that these agents have any discernible impact in wild populations.

However, some RLOs have become problematic in the aquaculture of various types of crustaceans, and

hence their introduction into new areas can have some consequences for crustacean aquaculture in the

affected region. However, these disease agents are relatively easy to control in aquaculture situations via

better husbandry practices and use of antibiotics (Lightner et al. 2008). Only MHD-SL is listed as a

reportable disease by the OIE and NACA (as well as in QLD, NSW and Victoria (Table 3b)), however

MHD-SL is exotic to Australia, and the spread of endemic RLOs to new areas is unlikely to impact trade.

Considering all of these factors, establishment of RLOs into new areas would have mild consequences for

crustacean aquaculture that would be amenable to control, and they are unlikely to cause any noticeable

environmental effects. It is therefore estimated that the consequences of introduction of RLOs into different

parts of the Australian environment via use of infected bait would likely be Very low.

_________________________________________________________________________________________________________________________________________________________________________________


178

www.digsfish.com


The unrestricted risk associated with crustacean RLOs is determined by combining the likelihood of entry


estimate for crustacean RLOs does not exceed the ALOP for any of the commodity types, suggesting that

additional risk management for these disease agents is not required at this time.

Risk estimate for infection of crustaceans with RLOs

Commodity type Live

crustaceans

Whole fresh

dead

crustaceans

Frozen whole

crustaceans

Frozen

crustacean

tails

Frozen

crustacean

heads

Combined likelihood

of release and

exposure


Consequences of

establishment



risk


risk

Negligible risk

_________________________________________________________________________________________________________________________________________________________________________________


179

www.digsfish.com

5.27 Infection of crustaceans with Hematodinium spp.

5.27.1 Aetiologic agent: Hematodinium australis and other parasitic dinoflagellates of the genus

Hematodinium (Family Syndiniceae).


5.27.3 Australias status: Hematodinium australis has been reported from crabs in Moreton Bay and the

Great Barrier Reef QLD (Hudson and Shields 1994), while Hematodinium-like agents have been reported

from crustaceans in Western Australia. The disease is not reportable in any State (Table 3b).

5.27.4 Epidemiology

Dinoflagellates in the genus Hematodinium are parasites of wild marine decapod crustaceans, particularly

crabs and lobsters (Stentiford and Shields 2005), however, recently they have also begun to be problematic

in cultured crustaceans such as mud crabs Scylla serrata (see Li et al. 2008) and even palaemonid prawns

(Xu et al. 2010). The genus Hematodinium was first described by Chatton and Poisson (1931), who found

Hematodinium perezi in the haemolymph of Carcinus maenas and Liocarcinus depurator in France. Several

studies have since described major Hematodinium spp. epizootics that have damaged fisheries for a wide

range of species of decapod crustaceans in many countries (Stentiford and Shields 2005). Several crab

species in Australia are infected with Hematodinium australis, including Portunus pelagicus, Scylla serrata

and Trapezia areolate (Shields 1992, Hudson and Shields 1994, Hudson and Adlard 1994, 1996). However,

to date prevalences of H. australis in the wild in Australia have been low, and there has been no indication of

significant disease at a population level at this time (Biosecurity Australia 2009). Hematodinium sp. have

been recorded from a broad range of decapod crustacean hosts, suggesting that these parasites are host

generalists (Stentiford and Shields 2005). Indeed, amphipods may act as alternate or reservoir hosts for

Hematodinium (Hudson and Shields 1994, Shields 1994).

Hematodinium australis was differentiated from the type species, H. perezi Chatton and Poisson 1931 on the

basis of size of the vegetative stage (trophont), the presence of rounded plasmodial stages and the southern

hemisphere location of H. australis (see Hudson and Adlard 1994). Molecular studies by Hudson and

Adlard (1996) later supported the separation of H. australis from other known forms of Hematodinium. The

highly pathogenic Hematodinium sp. parasites from Nephrops norvegicus (see Field et al. 1992) and

Chionoecetes bairdi (see Meyers et al. 1987) are likely to be new species, but until there is comparative work

with the type species, it will remain difficult to place them within the genus (Stentiford and Shields 2005).

Crabs and lobsters affected by Hematodinium sp. undergo dramatic pathological alterations to their organs,

tissues and haemolymph and eventually die (Meyers et al. 1987, Field et al. 1992, Stentiford and Shields

2005). Infections in N. norvegicus are associated with moribund lobsters displaying an abnormal dull orange

colouration, with 'watery' muscles, low haemolymph pressure and milky-white body fluids (Field et al.

1992). Other species exhibit similar signs of opaque discolouration of the carapace, milky white body fluids

and haemolymph that does not clot, and a “chalky” or “cooked” appearance of the flesh, with the external

_________________________________________________________________________________________________________________________________________________________________________________


180

www.digsfish.com

signs of infection accompanied by several physiological and biochemical disruptions to the muscles and

other organs which substantially alter the metabolism of infected hosts (Stentiford and Shields 2005).

The lifecycle consists of at least 3 phases (Stentiford and Shields 2005): a multinucleate plasmodial stage, a

vegetative phase (trophont, produced via merogony) and an asexual reproductive phase (sporont produced

via sporogony). In the Syndinida, sporogony leads to the formation of 2 dissimilar forms of biflagellate

dinospores (‘swarmers’) that arise from different parent infections and ensure dispersal and new infection.

Virtually all of the Syndinida are parasitic in the haemocoels of invertebrate hosts. They occur primarily as

plasmodial forms that divide and grow until they undergo sporogony to produce a motile spore stage. The

plasmodial stage has no chloroplasts and obtains nutrition via osmotrophy during the trophic phase, where

lipid and polysaccharide inclusions suggest active feeding at the expense of the host. Sporogenesis is simple

with multiplication of the nuclei, plasmodial and cytoplasmic divisions occurring to produce sporocysts,

from which the biflagellate zoospores are produced and liberated. Mortality rate of infected crabs is often

100% (Meyers et al. 1996) and death of the host usually follows sporulation (Stentiford and Shields 2005).

No resting cyst stages of the life cycle have been reported to date, though their presence cannot be ruled out

at this time (Stentiford and Shields 2005).

In Callinectes sapidus off Florida, USA, Hematodinium sp. infections reached a peak prevalence of 30 %

(Newman and Johnson 1975). Messick (1994) subsequently reported an epizootic of Hematodinium that

affected 70 to 100% of the juvenile Callinectes sapidus in the seaside bays of Maryland and Virginia in 1991

and 1992. In France, Hematodinium perezi infections are associated with winter crab mortalities, with peak

prevalences in velvet crab Necora puber observed to be as high as 87 %, resulting in a catastrophic 96%

decline in the local fishery (Wilhelm and Boulo 1988, Wilhelm and Mialhe 1996). Infections of

commercially fished populations of tanner and snow crabs in the Bering Sea and southeast Alaskan waters

with Hematodinium sp. were reported with peak prevalences approaching 100 % (Meyers et al. 1990, 1996,

Eaton et al. 1991). Evidence from the snow crab (Chionoecetes opilio) fishery in Newfoundland, Canada,

shows the prevalence of Hematodinium sp. has increased steadily from 0.037% to 4.25% over a 10 year

period (Pestal et al. 2003), affecting over 9% of males and 25% of females in an epizootic occurring in

Conception Bay in 2000 (Shields et al. 2005). Prevalence of Hematodinium sp. in wild Nephrops norvegicus

was up to 70% (Field et al. 1992). Outbreaks of disease due to Hematodinium sp. in wild populations of

crustaceans tend to occur in areas with entrained water masses such as lagoons, embayments or fjords with

shallow sills (Meyers et al. 1987, 1990, Eaton et al. 1991, Field et al. 1992, Wilhelm and Miahle 1996,

Messick 1994). Clearly, given suitable conditions, at least some variants of Hematodinium sp. represent a

significant threat to wild populations of decapod crustaceans.


Several crustaceans that occur along the east coast of Australia which can be used as bait and/or berley are

known to harbour infections of Hematodinium australis, including Scylla serrata and Portunus pelagicus

(see Shields 1992, Hudson and Adlard 1994). Hematodinium-like agents have also been reported from P.

pelagicus in Western Australia15. Large quantities of crustaceans are used as bait throughout Australia

(Kewagama Research 2002, 2007), but these are mainly penaeid prawns, and H. australis has not been

15 http://asvp.inov8design.info/wp-content/uploads/2010/07/de-2008-26.pdf

_________________________________________________________________________________________________________________________________________________________________________________


181

www.digsfish.com

recorded from penaeids in Australia to date. The quantities of crabs used as bait in Australia is likely to be

relatively small (used by around 2.1% of fishers, Kewagama Research 2007), and the prevalences of H.

australis in crab populations is very low, up to 4% in P. pelagicus and 1.5% of S. serrata in Moreton Bay

(Hudson and Shields 1994, Hudson and Lester 1994). Infected crabs can, however, harbour very high

intensity Hematodinium infections, for example 1 x 106 Hematodinium cells/ml in the haemolymph (Hudson

and Shields 1994). Also, fresh or frozen crabs are widely distributed as food fish and hence some product

could be diverted to use as bait or berley around the country.

Frozen crustaceans would not be expected to contain viable Hematodinium sp. parasites as freezing

inactivates most types of vegetative stages of protozoan parasites of similar size, due to disruption of their

cell walls (Jones and Gibson 1997). However, Hematodinium spp. are algae, and since cyst stages of many

algal species are highly resistant to freezing, it is possible that if a resting cyst stage occurs in the lifecycle of

Hematodinium spp, this could also be resistant to freezing (Robinson et al. 2005). Taking into account the

relatively small quantities of crabs used as bait or berley, as well as the low prevalence of these agents in

wild populations of crustaceans in some parts of the country, the likelihood estimations for the occurrence of

Hematodinium spp. in these commodities are listed below.

Release assessment for infection of crustaceans with Hematodinium spp.

Commodity

type

Live

crustaceans

Whole fresh

dead

crustaceans

Frozen

whole

crustaceans

Frozen

crustacean

tails

Frozen

crustacean

heads

Likelihood of

release

Moderate Moderate Extremely

Low

Extremely

Low

Extremely

Low


Crustaceans throughout Australia are already at risk of exposure to H. australis that occurs naturally in the

Australian environment. Translocation of infected crustaceans containing H. australis via their use as bait

could nevertheless transport this disease agent into new regions, but infection and establishment in new hosts

would occur only if viable H. australis was introduced into an area where susceptible hosts were present

under suitable environmental conditions for transmission. Inoculation experiments have shown that all

stages of the Hematodinium lifecycle, including filamentous trophonts, vegetative amoeboid trophonts,

microspores and macrospores, are capable of establishing new infections (Meyers et al. 1987, Eaton et al.

1991, Hudson and Shields 1994). Meyers et al. (1996) found potential evidence for sexual transmission of

Hematodinium in Chionoecetes bairdi with parasites present in the seminal fluids of the vas deferens in a

few males, but the importance of this potential route of infection needs further work. Infection by co

habitation through horizontal transmission of dinospores is possible (Stentiford and Shields 2005, Frischer et

al. 2006), as is infection via the per-os route through cannibalism. Walker et al. (2009) demonstrated how

blue crabs (Callinectes sapidus) became infected after eating a small portion of heavily infected conspecifics,

_________________________________________________________________________________________________________________________________________________________________________________


182

www.digsfish.com

with 50% mortality occurring after 4 days, however Hudson and Shields (1994) did not observe per-os

transmission in P. pelagicus or S. serrata when they were fed tissue from P. pelagicus that was infected with

H. australis. It appears likely that some of the experimental conditions (e.g. water temperature) available to

Hudson and Shields (1994) was unfavourable for natural transmission of the parasite at the time of their

experiments, especially given that infection was transmitted when the same hosts were injected with 0.1 ml

of haemolymph containing 0.2 - 1 x 106 Hematodinium cells/ml, with the LD50 by inoculation being

approximately 1 x 105 Hematodinium cells (Hudson and Shields 1994).

Based on the information available from overseas studies, wild crabs and lobsters feeding on clinically

diseased crustaceans used as bait or berley would likely have a significant chance of becoming infected by

Hematodinium spp. via the per-os route, or even by horizontal transmission (Frischer et al. 2006). Since the

minimum dose required for successful transmission of Hematodinium spp. is not well known, it is difficult to

determine whether the disease could be transmitted via the per-os route if subclinically affected crustaceans

were used as bait. In any case, Hematodinium sp. infections are highly pathogenic, with sub-clinical disease

naturally progressing to clinical disease with most infected crabs dying within 4 to 40 days, depending on

factors such as water temperatures (Stentiford and Shields 2005, Frischer et al. 2006, Walker et al. 2009).

Because susceptible crustaceans such as crabs and lobsters are predominantly scavengers, the transmission

and spread of these disease agents in populations of wild crustaceans has been observed, particularly in

entrained water masses such as lagoons, embayments or fjords with shallow sills flows (Stentiford and

Shields 2005). This suggests that Hematodinium spp. could be transmitted and become established after an

index case occurs, although these events would likely be modulated to a certain extent by predation of

moribund crustaceans by non susceptible species such as fish. Taking these various factors into

consideration, the risk of exposure and establishment of Hematodinium spp. in crustacean populations via

use of bait and berley remains non-negligible, and the likelihood of exposure and establishment of

Hematodinium spp. in new crustacean populations via translocation is considered to be Low.


A species of Hematodinium is already known to be present in the Australian environment at low prevalence,

and there is no evidence to date that it has any discernible impact on wild populations. However, in other

regions of the world, Hematodinium spp. infections have had significant detrimental impacts on fisheries and

populations of crabs and lobsters (Wilhelm and Boulo 1988, Wilhelm and Mialhe 1996). More recently,

Hematodinium spp. have become problematic in the aquaculture of various types of crustaceans, and hence

their introduction into new areas may result in significant mortalities and ongoing financial losses to

aquaculturists (Li et al. 2008, Xu et al. 2010), especially as there are no methods of control available,

although some individual crabs appear refractory to infection (Stentiford and Shields 2005). Hematodinium

spp. is not listed as a reportable disease by the OIE or NACA, and is also not listed as reportable in any State

(Table 3b), so at this time, the spread of Hematodinium spp. to new areas is unlikely to adversely impact

trade. Considering all of these factors, establishment of Hematodinium spp. into new areas would have

significant consequences for aquaculture of susceptible crustaceans (particularly crabs and lobsters)

potentially causing disease that would not be readily amenable to control, and its introduction into new

regions could also cause significant biological consequences and environmental effects, as well as potentially

significant adverse economic effects to crustacean fisheries. It is therefore estimated that the consequences

_________________________________________________________________________________________________________________________________________________________________________________


183

www.digsfish.com

of introduction of Hematodinium spp. into different parts of the Australian environment via use of infected

bait would likely be Moderate.


The unrestricted risk associated with Hematodinium spp. is determined by combining the likelihood of entry


estimate for Hematodinium spp. exceeds the ALOP for live crustaceans and whole fresh dead crustaceans,

suggesting that additional risk management for this disease agent is required in these commodities.

Risk estimate for infection of crustaceans with Hematodinium spp.

Commodity type Live

crustaceans

Whole fresh

dead

crustaceans

Frozen whole

crustaceans

Frozen

crustacean

tails

Frozen

crustacean

heads

Combined likelihood

of release and

exposure


Consequences of

establishment


Risk estimation Low Risk Low Risk Negligible risk Negligible

risk

Negligible risk

_________________________________________________________________________________________________________________________________________________________________________________


184

www.digsfish.com

5.28 Infection of crustaceans with Microsporidians

5.28.1 Aetiologic agent: Thelohania spp., and other parasites of the Phylum Microsporea that infect

crustaceans.


5.28.3 Australias status: Microsporidian infections have been reported from crustaceans in all States, and

microsporidosis of crustaceans is reportable in WA (Table 3b).

5.28.4 Epidemiology

Microsporidians are obligate intracellular parasites known to infect a wide variety of eukaryotic hosts,

including arthropods, fish, and mammals (Lom and Dykova 1992). Microsporidians are common parasites

of crustaceans, with a large number of genera being reported from crustacean hosts, including Agmasoma,

Ameson, Enterocytozoon, Enterospora, Flabelliforma, Glugoides, Indosporus, Nadelspora, Nosema,

Ordospora, Pleistophora, Thelohania, Vavraia, Tuzetia and others. Microsporidosis generally occurs at low

prevalence in wild crustaceans all over the world, and occasionally they are reported in cultured crustaceans

as well (Anderson et al. 1989, Flegel et al. 1992, Hudson et al. 2001, Vidal–Martinez et al. 2002, Tourtip et

al. 2009). Crustaceans infected by microsporidians often have opaque musculature and are unmarketable,

and the condition has been associated with significant disease in some instances (Flegel et al. 1992, Lightner

1996, Hudson et al. 2001). The life-cycles of many microsporidians affecting crustaceans may be indirect

(Breed and Olson 1977, Flegel et al. 1992, Herbert 1988, Edgerton et al. 2002), which means that the

chances of transmission of the disease in some crustacean populations may be reduced as it may depend on

the presence of intermediate hosts (such as copepods or insects). However there may be some exceptions to

this (Langdon and Thorne 1992, Hudson et al. 2001), and these examples of direct transmission via per-os

routes may be due to consumption of presporogenic stages (Langdon and Thorne 1992). Because the life

cycles of microsporidians that infect crustaceans are so poorly understood, the possibility remains that life

cycles may differ even between closely related species (Edgerton et al. 2002).


In Australia, microsporidian infections have been recorded throughout the country in a wide variety of

crustacean species used as bait or berley (O’Donoghue and Adlard 2000), including penaeid prawns,

(Penaeus monodon, Penaeus esculentus, Penaeus semisulcatus, Penaeus merguiensis, Melicertus

latisulcatus, Penaeus spp.), freshwater prawns (Macrobrachium spp.), freshwater crayfish (Cherax

destructor, Cherax quadricarinatus, Cherax tenuimanus, C. cainii, Cherax spp.), crabs (Portunus pelagicus)

and lobsters (Panulirus cygnus, Panulirus spp.) (Table 8). Several species of microsporidians have been

identified in Australia, including Vavraia parastacida (see Langdon 1991b, Langdon and Thorne 1992),

Thelohania spp. (see Herbert 1988, Shields 1992, Jones and Lawrence 2001), Thelohania montirivulorum

(see Moody et al. 2003a), T. parastaci (see Jones and Lawrence 2001, Moody et al. 2003c), Vairimorpha

cheracis (see Moody et al. 2003b) and others. Prevalence of infection of wild prawns can be quite low, with

Ameson spp. occurring in 0.1% of prawns from northern Australia (Owens and Glazebrook 1988) and at

_________________________________________________________________________________________________________________________________________________________________________________


185

www.digsfish.com

similarly low prevalences in P. pelagicus in Moreton Bay (Shields and Wood 1993) , while prevalence of

microsporidosis in crayfish can be higher, with Herbert (1988) finding Thelohania spp. at a prevalence of

7.8% in C. quadricarinatus. These parasites appear to occur in a range of environments throughout the

country, however the extent of the distributions of the various species of microsporidian parasites of

crustaceans remains largely unknown.

Large quantities of crustaceans, mainly prawns, are used as bait throughout Australia (Kewagama Research

2002, 2007). Live prawns are generally not available commercially, but 11% of recreational fishers catch

their own prawns (Kewagama Research 2007), some of which are likely to be used live and thus could be

translocated short distances by fishers. In some crustaceans heavily infected with microsporidians, the

animals do not feed, suggesting that they would be less likely to enter traps, but their locomotion may also be

impaired by the infection, suggesting that they would be more likely to be collected by fishers using

equipment such as dipnets (Moodie et al 2003b). Infected fresh, whole, green (uncooked) crustaceans would

also still be expected to contain viable microsporidians, even crayfish or prawns with heads removed, as

microsporidians commonly infect the tail muscle. Also, fresh or frozen crayfish are widely distributed as

food fish and hence some product could be diverted to use as bait or berley around the country, however

since heavy microsporidian infections are grossly visible, infected crustaceans would be removed from sale

via normal quality control processes. The likelihood of release will depend on the ability of infective stages

to remain viable under the conditions of use of their hosts as bait or berley, and it appears that

microsporidian spores can remain viable in the natural environment for months to years. For example,

spores of Loma salmonae remained viable when stored in freshwater or seawater at 4°C for up to 95 days

(Shaw et al. 2000), and spores of Glugea stephani remained viable after 17 months at 5°C (Amigo et al.

1996). The viability of microsporidian spores after freezing varies widely, depending on the species studied.

Amigo et al. (1996) found 3.6% of spores of G. stephani ( a fish parasite) remained infective after being

frozen at -19°C for 24 hours, while Overstreet and Whatley (1975) found that some spores of Ameson

michaelis from a crab survived 67 days freezing at -22°C. Taking into account the large quantities of

crustacean products used as bait or berley (Table 1), and also the fact that microsporidian infections have

been recorded from a wide range of crustaceans used as bait throughout the country, though mostly at low

prevalences, the likelihood estimations for the occurrence of viable microsporidian parasites in these


Release assessment for infection of crustaceans with microsporidians

Commodity

type

Live

crustaceans

Whole fresh

dead

crustaceans

Frozen

whole

crustaceans

Frozen

crustacean

tails

Frozen

crustacean

heads

Likelihood of

release

High High Moderate Moderate Moderate

_________________________________________________________________________________________________________________________________________________________________________________


186

www.digsfish.com


Crustaceans throughout marine and freshwater environments in Australia are likely to be already at risk of

natural exposure to infective stages of microsporidians. However, large quantities of fresh or frozen whole

or processed crustaceans are offered for sale as bait or berley via wholesale outlets such as commercial

fishing co-operatives, or packaged for resale at retail outlets such as fishing tackle shops, service stations and

supermarkets. Crustaceans heavily infected with microsporidians often show grossly visible signs of disease

such as white discolouration of the flesh, which would be identified and rejected during normal quality

control procedures used for fresh or frozen food fish, largely eliminating the pathway of exposure via

diversion of food fish for use as bait or berley. However, subclinical infections cannot be detected visually,

and even heavily infected crustaceans with gross signs of disease would still be harvested and considered

acceptable for use as bait. This suggests that direct pathways exist for translocated crustacean products

infected by microsporidians to enter both freshwater and marine environments, thus potentially exposing

potentially susceptible wild crustaceans to viable infective stages of novel microsporidians. However,

infection and establishment would occur only if sufficient quantities of infective stages (i.e. an infective

dose) were introduced into an area where susceptible hosts were present under conditions suitable for

transmission.

Some microsporidians of crustaceans are transmitted directly (Langdon and Thorne 1992), but the minimum

infective dose of infective stages required for successful transmission has not been determined for the

majority of species, and this also probably will vary depending on the identity of the host and its immune

status. Infection can be achieved by the per-os route for at least some crustacean microsporidians if

susceptible crustaceans ingest presporogenic stages, though susceptibility may vary between hosts (Langdon

and Thorne 1992). When the natural course of infection is considered, it is clear that infection can be

theoretically achieved after exposure to a dose as small as a single viable spore or presporogenic stage,

though the dose required to cause host mortality will depend on many factors. Lightly infected crustaceans

may contain thousands of spores and presporogenic stages, while one heavily infected crustacean may

contain literally millions of spores and presporogenic stages (Lom and Dykova 1992).

Given that pathways exist for translocation and spread of viable microsporidians into the environment via

use of bait, and acknowledging that some microsporidians can infect a range of susceptible hosts (but others

may not), and the infective doses required for transmission may be very small in comparison to the parasite

burden carried by a single infected crustacean, the risk of exposure and establishment is non-negligible, and

the likelihood of exposure and establishment of microsporidians in new crustacean populations is considered

to be Moderate.


Crustaceans of all size classes can become infected with microsporidians. In susceptible species,

disfigurement and reduction of market value of affected crustaceans can result in economic losses, while

mortalities of wild and cultured crustaceans have been recorded in Australia and other parts of the world

where microsporidian outbreaks have occurred. This suggests that some microsporidians have the potential

to cause damage to wild crustacean populations (including rare and threatened native freshwater crayfish, see

Coughran and Leckie 2007, Coughran et al. 2009) as well as aquacultured crustaceans. While the full range

_________________________________________________________________________________________________________________________________________________________________________________


187

www.digsfish.com

of microsporidian parasites and their susceptible hosts in Australia remain to be determined, microsporidians

have already been recorded from a wide range of crustaceans in both freshwater and marine environments,

and translocations of microsporidians via movements of live crustaceans have already been documented

(Langdon 1991b, Jones and Lawrence 2001). No microsporidian diseases of crustaceans are listed by the

OIE or NACA, but microsporidosis is a reportable disease in WA (Table 3b). Hence the spread of these

disease agents could have minor adverse effects on trade. Considering all of these factors, establishment of

microsporidians in new areas would likely have moderate biological consequences, which may not be

amenable to control in wild or cultured populations, and could also cause some unwanted environmental

effects. It is estimated that the consequences of introduction of microsporidians into crustacean populations

in different parts of the Australian environment via use of infected bait would likely be Low.


The unrestricted risk associated with microsporidian infections of crustaceans is determined by combining

the likelihood of entry and exposure (from Table 5) with the consequences of establishment (Table 7). The

unrestricted risk estimate for microsporidians exceeds the ALOP for live crustaceans and whole fresh dead

crustaceans, suggesting that additional risk management is required for these disease agents in these

commodities.

Risk estimate for infection of crustaceans with microsporidians

Commodity type Live

crustaceans

Whole fresh

dead

crustaceans

Frozen whole

crustaceans

Frozen

crustacean

tails

Frozen

crustacean

heads

Combined likelihood

of release and

exposure

Moderate Moderate Low Low Low

Consequences of

establishment

Low Low Low Low Low


risk

Very low risk

_________________________________________________________________________________________________________________________________________________________________________________


188

www.digsfish.com

5.29 Infection of crabs with Sacculina spp. and other rhizocephalan barnacles

5.29.1 Aetiologic agent: Sacculina granifera and other parasitic barnacles in the Family Sacculinidae.


5.29.3 Australias status: Sacculinid infections have been reported from crabs in all States, and infection

with Sacculina spp. was reportable in SA under the Fisheries Act (Table 3b).

5.29.4 Epidemiology

Rhizocephalan barnacles of the genera Sacculina Loxothylacus and Heterosaccus and several others (Family

Sacculinidae) are parasitic castrators of several species of crabs in various parts of the world (Boschma 1955,

Murphy and Goggin 2000, Walker 2001, Glenner et al. 2008). In Australia, Sacculina granifera has a

marked effect on the gonad development, growth and behaviour of its host Portunus pelagicus (see Phillips

and Cannon 1978, Bishop and Cannon 1979, Weng 1987). The barnacle infects both male and female crabs

mainly as juveniles, but P. pelagicus were susceptible to infection by the parasite at any size/age (Weng

1987, Shields and Wood 1993). Infected P. pelagicus are frequently castrated by the parasite, but in some

cases, castration was incomplete and the reproductive potential of the infected host was not zero (Shields and

Wood 1993). Although male crabs were completely castrated (Shields 1992), infected female crabs were

capable of mating and, in a few cases, infected female crabs produced egg clutches with reduced numbers of

viable eggs (Shields and Wood 1993).

Sacculina granifera modifies the behaviour of its host, inducing migration of infected male and female crabs

to the spawning areas usually frequented only by berried uninfected female crabs (Bishop and Cannon 1979).

This is thought to increase the chance of infection of both adult female crabs as well as crab recruits by

retaining larval S. granifera in the same water masses as those of larval P. pelagicus (Phillips and Cannon

1978, Bishop and Cannon 1979). Rhizocephalan barnacles are obligate endoparasites and infection of the

host is horizontal and direct via planktonic naupliar stages that exit the externa and moult into the larval

settlement stage (cypris larvae). Details of the various modes of infection vary between parasite species,

however a generalized lifecycle involves female nauplii moulting into female cypris larvae that settle on the

host and either penetrate the host or carapace directly (akentrogonid species), or penetrate after moulting into

other infective instars such as kentrogon and vermigon stages (Hoeg 1990, Walker 2001). Once inside the

host, the female infective stage circulates in the haemolymph and begins to develop the characteristic “root”

system inside host organs (Hoeg 1990). In time a virgin externa erupts out of the crabs abdomen, and male

cypris larvae settle on the externa and moult to the trichogon instar to enable fertilization (Hoeg 1990,

Walker 2001).

The moult condition of P. pelagicus is significantly affected by the presence of S. granifera, with

development of the externae of the parasite inhibiting moulting activity (Weng 1987, Shields and Wood

1993, Sumpton et al. 1994). Since moulting and mating are intimately linked in P. pelagicus (Females mate

with males immediately after moulting), the presence of S. granifera in females therefore results in eventual

reproductive failure once stored sperm supply is exhausted (Shields and Wood 1993, Sumpton et al. 1994).

_________________________________________________________________________________________________________________________________________________________________________________


189

www.digsfish.com

Prevalence of S. granifera in Moreton Bay varied with season, ranging from over 40% of males and 20% of

females in spring and early summer, to less than 5% and 0% in males and females, respectively, in winter

(Shields and Woods 1993). Parasite externae were also most common in late spring/ early summer, with

internae prevalence peaking in June (Shields and Woods 1993). Studies that examine for the presence of

externae only tend to underestimate the prevalence of the parasite, such as Weng (1987) who recorded S.

granifera prevalences of only 6% in Moreton Bay and 1.2% in the Gulf of Carpentaria.


In Australia, infections by rhizocephalan parasites have been recorded in several species of crabs including

P. pelagicus in QLD (Phillips and Cannon 1978, Bishop and Cannon 1979, Weng 1987, Shields 1992,

Shields and Wood 1993), and Charybdis spp. in QLD, NT and WA (Walker and Lester 1998). These

parasites appear to occur in a range of environments throughout the country, however the extent of the

distributions of the various species of rhizocephalan parasites of crustaceans remains largely unknown.

Large quantities of crustaceans are used as bait throughout Australia (Kewagama Research 2002, 2007), but

these are mainly penaeid prawns, and rhizocephalan parasites infect crabs. The quantities of crabs used as

bait in Australia is likely to be relatively small (used by around 2.1% of fishers, Kewagama Research 2007),

however prevalence of infection of wild crabs can be high, approaching or exceeding 30% in some locations

and at certain times of year (Shields and Wood 1993). Also, fresh or frozen crabs are widely distributed as

food fish and hence some product could be diverted to use as bait or berley around the country. Fresh dead

crabs are also likely to contain some viable infective stages at certain times of year, though the length of time

these parasites remain alive in dead crabs does not appear to have been published. Frozen crabs would not

be expected to contain viable rhizocephalan parasites as freezing inactivates most types of metazoan

parasites due to disruption of their cell walls (Jones and Gibson 1997). Taking into account the relatively

small quantities of crabs used as bait or berley, as well as the sometimes high prevalence of these agents in

wild populations of crabs in some parts of the country, the likelihood estimations for the occurrence of

rhizocephalan parasites in these commodities are listed below.

Release assessment for infection of crabs with Sacculina spp. and other rhizocephalan barnacles

Commodity

type

Live crabs Whole fresh

dead crabs

Frozen

whole crabs

Likelihood of

release

High Very low Negligible


Crabs throughout many parts of Australia are already at risk of exposure to rhizocephalan parasites that

occur naturally in the Australian environment. Translocation of infected crabs containing rhizocephalan

_________________________________________________________________________________________________________________________________________________________________________________


190

www.digsfish.com

parasites via their use as bait or berley could nevertheless transport these disease agents into new regions, but

infection and establishment in new hosts would occur only if viable infective stages were introduced into an

area where susceptible hosts were present under suitable environmental conditions for transmission.

Infection by cohabitation through horizontal transmission of larval cyprid infective stages is possible, as this

is how the disease is transmitted naturally in the wild. Infection and establishment of rhizocephalan parasites

in new hosts requires the presence of both male and female cyprid infective stages in sufficient

concentrations in the water over a long period of time for them to encounter and infect a new host and allow

fertilization of the virgin externa by male cyprids. The concentrations of larvae and time scales required for

successful transmission are not fully known, and would likely vary significantly depending on environmental

conditions. However, if an index case occurred, the chances of establishment of the parasite within the host

population would be greatly increased, although these events could be modulated to a certain extent by

predation of infected crustaceans by non susceptible species such as fish. Taking these various factors into

consideration, the risk of exposure and establishment of rhizocephalan parasites in new crab populations via

use of bait and berley remains non-negligible, and the likelihood of exposure and establishment of

rhizocephalan parasites in new crab populations via translocation is considered to be Low.


Various species of rhizocephalan parasites are already known to be present in the Australian environment in

some populations of crabs, often at reasonably high prevalence, but they appear absent in others, such as the

P. pelagicus fishery in SA (M. Deveney, personal communication). The presence of Sacculina granifera in

P. pelagicus in Moreton Bay is likely to have a significant impact on the recruitment available to the fishery

(Shields and Wood 2003), and infected crabs may also be considered less marketable due to the presence of

the visible externa. On the other hand, infections due to rhizocephalan parasites are unlikely to be

problematic in the aquaculture of crabs provided broodstock are free from the infection, although it would be

detrimental to soft shell crab production based on capture of wild crabs, as the parasite prevents moulting.

Rhizocephalan parasites are not listed as reportable diseases by the OIE or NACA, however, infection by

Sacculina spp. is reportable in SA (Table 3b), so at this time, the spread of rhizocephalan parasites to new

areas is unlikely to adversely impact trade. Considering all of these factors, establishment of rhizocephalan

parasites into new areas could have significant consequences for wild crab fisheries, but limited adverse

consequences for aquaculture, and its introduction into new regions could also cause significant biological

consequences and environmental effects, as well as potentially significant adverse economic effects to crab

fisheries. It is therefore estimated that the consequences of introduction of rhizocephalan parasites into

different parts of the Australian environment via use of infected bait would likely be Moderate.


The unrestricted risk associated with rhizocephalan parasites is determined by combining the likelihood of


estimate for rhizocephalan parasites exceeds the ALOP only for live crabs, suggesting that additional risk

management for these disease agents is required for these commodities.

_________________________________________________________________________________________________________________________________________________________________________________


191

www.digsfish.com

Risk estimate for infection of crabs with Sacculina spp. and other rhizocephalan barnacles

Commodity type Live crabs Whole fresh dead crabs Frozen whole crabs

Combined likelihood

of release and

exposure

Low Very low Negligible

Consequences of

establishment

Moderate Moderate Moderate

Risk estimation Low risk Very low risk Negligible risk

_________________________________________________________________________________________________________________________________________________________________________________


192

www.digsfish.com

5.30 Viral Ganglioneuritis of Abalone (AVG)

5.30.1 Aetiologic agent: Abalone herpes-like virus (AbHV-1) that is the second member of the genus

Haliotivirus (Family Malacoherpesviridae), which infects abalone Haliotis laevigata and Haliotis rubra (see

Savin et al. 2010).


5.30.3 Australias status: AVG infections have been reported from Victoria and Tasmania, and the disease

is reportable in all States except the ACT (Table 3b).

5.30.4 Epidemiology

In December 2005 a disease outbreak in greenlip abalone (Haliotis laevigata), blacklip abalone (H. rubra)

and hybrid abalone (Haliotis laevigata × H. rubra) due to a novel herpes-like virus occurred in three abalone

aquaculture facilities, two landbased farms in western Victoria and one sea based farm in central Victoria

(Victoria DPI 2006, Hooper et al. 2007, Hills 2007, Corbeil et al. 2010). The virus was associated with

inflammation and necrosis of neural ganglia and the ganglioneuritis was associated with sudden high levels

of mortalities up to 90% within 14 days of onset (Hooper et al. 2007). All sizes of abalone were affected and

exhibited signs including swollen mouths and prolapse of the radula, and loss of righting reflex (Hooper et

al. 2007). Electron microscope and genetic studies confirmed the disease agent causing abalone

ganglioneuritis (AVG) was a neurotrophic herpes-like virus (Tan et al. 2008, Corbeil et al. 2010, Savin et al.

2010) closely related to herpes-like viruses responsible for mortalities in abalone in Taiwan (Chang et al.

2005, Corbeil et al. 2010, Savin et al. 2010). The sea based farm in Westernport Bay central Victoria had

received stock from one of the landbased farms, noted increased mortality rates due to AVG, and voluntarily

destocked and decontaminated the facility (Hills 2007, Hooper et al. 2007). However a 4th farm in

Westernport Bay, located 640 meters away from the 3rd farm, became infected in late April 2006 and was

depopulated and decontaminated by early May 2006 (Victoria DPI 2006, Hills 2007). Both landbased farms

pumped seawater into the facility, through their tanks then back out into the ocean via settling ponds. The

landbased farms did not immediately destock and in early May 2006, diseased abalone with AVG were

detected in wild populations of abalone on reefs adjacent to one of the affected facilities at Port Fairy

(Victoria DPI 2006). Since then AVG has spread from this point easterly and westerly along the Victorian

coast, at up to 5 - 10 km/month (Hills 2007), significantly impacting wild abalone populations and

substantially reducing commercial catches and recruitment in the wild fishery (Mayfield et al. 2011).

In mid 2008, wild caught abalone sampled from a commercial processing facility in Tasmania that was

recording low levels of mortality were positive for AbHV-1 by PCR at a prevalence of 39% (Crane et al.

2009, Corbeil et al. 2010). Further research found that the disease agent occurs naturally at very low

prevalences (3 out of 1625 abalone = 0.18% prevalence) in subclinical infections of wild populations of

abalone in Tasmania (Corbeil et al. 2010). The virus was detected again at another abalone processing plant

_________________________________________________________________________________________________________________________________________________________________________________


193

www.digsfish.com

in Tasmania in 2009, followed by more recent outbreaks of AVG in processing facilities in December 2010

and a landbased abalone aquaculture facility in Bicheno on Tasmanias east coast in January 201116.


It appears that AbHV-1 is endemic in wild abalone populations in Tasmanian waters at very low prevalences

and another strain of the virus also occurs in the coastal waters of western Victoria at moderate prevalences

(Crane et al. 2009, Corbeil et al. 2010, MJ Crane, personal communication). The disease agent is only

known to infect abalone at this time, and it has not been reported from any other regions of Australia.

Abalone processing waste (abalone guts) were sold commercially and used as bait and berley by less than

1% of recreational fishers in Australia in 2006, mainly in Tasmania, NSW and QLD (Kewagama Research

2007). However, since the outbreak of AVG in Victoria, use of abalone processing waste as bait and berley

has been halted in all states (e.g. Victoria DPI 2010). However, live, fresh and frozen abalone are widely

distributed as seafood and hence some abalone products could be illegally diverted to use as bait or berley

around the country.

Crane et al. (2009) froze the viral inocula at -20°C, -80°C and in liquid nitrogen (-196°C) for periods of up to

21 months to examine the stability of the virus in storage. Their data suggested that AbHV-1 is quite stable

at -20°C and can remain viable after freezing with minimal loss of infectivity after 6 months, with reduced

infectivity (but still mortality) after 21 months. There was minimal loss of infectivity after 21 months at -

80°C and -196°C (Crane et al. 2009). AbHV-1 is naturally transmitted horizontally via the water or by

mucus trails (Crane et al. 2009) and infection of susceptible abalone causes pathological changes in the

neural ganglia, mainly those in the cerebral and buccal regions, but also in nerve bundles and pleuropedal

ganglia within the foot muscle (Hooper et al. 2007). Because of this, removal of the viscera from abalone

would not necessarily result in a marked reduction in the viral load of clinically diseased abalone. Taking

into account the small quantities of abalone used as bait or berley, as well as the fact that the disease agent

may be reasonably prevalent in wild populations of abalone in Victoria, the likelihood estimations for the

occurrence of AbHV-1 in these commodities are listed below.

Release assessment for AVG

Commodity type Live

abalone

Whole fresh

dead abalone

Frozen

whole

abalone

Frozen

abalone

meat

Frozen

abalone

viscera

Likelihood of release High High Moderate Moderate Moderate

16 http://www.dpiw.tas.gov.au/inter.nsf/WebPages/SCAN-75F423?open

_________________________________________________________________________________________________________________________________________________________________________________


194

www.digsfish.com


Abalone in Tasmania and parts of Victoria are already at risk of exposure to AbHV-1. However,

translocation of infected abalone from areas where AbHV-1 is enzootic via their use as bait could transport

the virus to new regions. Infection and establishment of this virus in new hosts would occur only if

sufficient viable viral particles were introduced into an area where susceptible abalone were present under

suitable environmental conditions for transmission. The virus appears to cause disease only in abalone, and

therefore the risks of other molluscs becoming diseased appears low, however there is little information

available regarding the host range of AbHV-1 and it is not known at this time whether other gastropod

molluscs are susceptible to infection or act as carriers.

Abalone of all sizes can become infected with AbHV-1 horizontally via the water. Crane et al. (2009) used

viral homogenates from clinically diseased abalone in a dilution series to find the LD50 by injection into the

pedal muscle, and found the LD50 to be 10-6.39 of the stock solution, suggesting that AbHV-1 is highly

virulent for abalone. When AbHV-1 was added to the water, the LD50 increased to around 10-2 of the stock

solution and after a 3 to 8 day prepatent period, death occurred over a period of 7 to 16 days (Crane et al.

2009). AbHV-1 was also transmissable horizontally by co-habitation with infected abalone with 100%

mortality observed within 8 days (McColl et al. 2007, Crane et al. 2009). Abalone in areas where infected

abalone products were being used as bait or burley could be exposed to viral particles, however the

concentration of viral particles required to initiate transmission horizontally via the water is much higher

(around a hundred fold dilution of stock solutions made from infected abalone) than the very small quantities

of virus needed for transmission via inoculation (Crane et al. 2009). However if portions of clinically

diseased abalone (perhaps a subclinically diseased individual taken from a processing plant and sold live into

a restaurant where the infection progressed to clinical disease and the abalone in the tank died and were not

fit for human consumption) were diverted for use as bait or berley, and were used in areas frequented by wild

abalone, a large amount of virus could be introduced into the water. It is not known whether exposure to low

concentrations of viral particles predisposes abalone to subclinical infection, but if infected bait or berley

were used in a small body of water (e.g. a rock pool), concentrations of virus may be sufficient for an index

case to occur. Given the population structure of these molluscs (and the precedents in the wild fishery in

Victoria), transmission and further spread from the index case would be possible, and the disease could

become established in the population, most likely in subclinically infected abalone. Taking these various

factors into consideration, the risk of exposure and establishment of AbHV-1 via use of bait and berley

remains non-negligible, and the likelihood of exposure and establishment of AbHV-1 in new abalone

populations via translocation is considered to be Moderate.


Although AbHV-1 is already present in populations of wild abalone in some regions of Australia, other

regions appear free of infection at this time. There is evidence that AbHV-1 can cause major disease

outbreaks and significant impacts on populations of both wild and cultured abalone. AGV is listed by the

OIE and NACA, and it is also listed as a reportable disease in all States except the ACT (Table 3b). Hence

the spread of AbHV-1 to new areas is likely to adversely impact trade. Considering all of these factors,

establishment of AbHV-1 in new areas would have serious biological consequences for abalone aquaculture

and cause significant economic harm together with irreversible environmental effects for wild abalone

_________________________________________________________________________________________________________________________________________________________________________________


195

www.digsfish.com

fisheries. It is therefore estimated that the consequences of introduction of AbHV-1 into different parts of

the Australian environment via use of infected bait would likely be High.


The unrestricted risk associated with AVG is determined by combining the likelihood of entry and exposure

(from Table 5) with the consequences of establishment (Table 7). The unrestricted risk estimate for AVG

exceeds the ALOP for all commodity types, suggesting that additional risk management for this disease

agent is required in these commodities.

Risk estimate for infection AVG

Commodity type Live abalone Whole fresh

dead abalone

Frozen whole

abalone

Frozen abalone

meat

Combined likelihood

of release and

exposure

Moderate Moderate Low Low

Consequences of

establishment

High High High High

Risk estimation High risk High risk Moderate risk Moderate risk

_________________________________________________________________________________________________________________________________________________________________________________


196

www.digsfish.com

5.31 Infection of oysters with Bonamia spp., and/or unidentified microcells

5.31.1 Aetiologic agent: Haplosporidian microcells of the genus Bonamia, including B. roughleyi, B.

exitiosa, and unidentified Bonamia species and/or other unidentified microcells.


5.31.3 Australias status: Bonamia and/or microcell infections have been reported in NSW, Victoria,

Tasmania, SA and WA, and the disease is reportable in all States (Table 3b).

5.31.4 Epidemiology

Microcell parasites of the genus Bonamia are currently classified within the haplosporidia (Hine et al 2009),

and several species within the genus occur in many regions around the world in wild and cultured oysters.

Bonamia microcells are very small (2-3 microns in diameter) and infect oyster haemocytes, epithelial cells

and connective tissues. The first described species of Bonamia, namely B. ostreae, caused epizootic disease

in populations of Ostrea edulis in France in 1979 (Pichot et al. 1980). Further research indicated that the

parasite was probably introduced into France and Spain through importation of O. edulis seed from the USA,

where undescribed microcell diseases had affected native flat oyster (Ostrea conchaphila) populations as

early as the 1960’s (Elston et al. 1986, Cigarria and Elston 1997). Between 1986 and 1992 mortalities of

over 90% of dredge oysters (Ostrea chilensis) were recorded in a wild fishery in Foveaux Strait in New

Zealand (Dinamani et al. 1987, Doonan et al. 1994, Cranfield et al. 2005). The deaths were found to be due

to a new parasite, Bonamia exitiosa (Hine et al. 2001, Berthe and Hine 2004). There is evidence that this

parasite was present in O. chilensis as far back as at least the 1960’s (Hine 1996a, Cranfield et al. 2005). The

increasing frequency and severity of epizootics in this fishery in recent years (another epizootic occurred

between 1999 and 2004, see Cranfield et al. 2005) are possibly related to environmental stressors such as

modification of benthic substrate by dredges, which may have increased the susceptibility of oysters to

bonamiosis (Cranfield et al. 2005). In light of the more recent detection of Bonamia exitiosa and B. exitiosa-

like microcells in a wide variety of locations around the world, including flat oysters in Spain (Abollo et al.

2008), Chile (Lohrmann et al. 2009), Tunisia (Hill et al. 2010) and Cornwall, England (OIE 2011), as well as

moribund Crassostrea ariakensis in North Carolina (Burreson et al. 2004, Bishop et al. 2006), the possibility

of translocation of the parasite to and/or from NZ and Australia (through routes such as oyster hull fouling

on shipping and ballast water, see Howard 1994, Bishop et al. 2006) cannot be discounted.

There may be two or more species of Bonamia in Australian waters. The first species discovered was

originally described as Mikrocytos roughleyi by Farley et al. (1988) and had been associated with “winter

mortality” in Sydney rock oysters (Saccostrea glomerata) as far back as the 1920’s (Roughley 1926).

However there are fundamental differences between this parasite and the ultrastructure of the only other

member of the genus, Mikrocytos mackini (the cause of Denman Island disease in Crassostrea gigas in

Canada, see Farley et al. 1988, Bower et al. 1997). Subsequent studies using DNA technology indicated that

M. roughleyi is in fact another species of Bonamia, namely Bonamia roughleyi (Cochennec-Laureau et al.

2003). Bonamia roughleyi is particularly interesting as unlike other Bonamia species, it appears to cause

disease in winter, not summer, but genetically it appears virtually identical to B. exitiosa (see Hill et al.

_________________________________________________________________________________________________________________________________________________________________________________


197

www.digsfish.com

2010). Another Bonamia was associated with mortalities of flat oysters Ostrea angasi in south eastern

Australia in early 1991 (Hine 1996a). Populations of O. angasi in NSW, Tasmania and WA are also infected

with this Bonamia sp. (see Hine 1996a, Heasman et al. 2003, Jones and Creeper 2006, Corbeil et al. 2009),

which is morphologically indistinguishable from B. exitiosa, as well as being very closely related on a

molecular basis (see Corbeil et al. 2006, Hill et al. 2010), though it exhibits slight differences in tissue

trophism and other aspects of infection (Corbeil et al. 2009). These differences could be related to the host,

however, rather than the parasite, due to the fact that the parasite could have been introduced when large

numbers of live NZ dredge oysters were moved from New Zealand to Australia in the early 1900’s to

replenish stocks of O. angasi which suffered from massive mortalities in the late 1800’s (Hine 1996a). The

fact that Roughley (1926) recorded mortalities in S. glomerata in NSW in the 1920’s, which was after the

introduction of the flat oysters from NZ into NSW waters, suggests winter mortality could well be a case of a

new non-equilibrium host-parasite relationship caused by introduction of B. exitiosa from New Zealand.

Indeed, it is possible that B. roughleyi and Bonamia sp. in Australia are both actually B. exitiosa (see Hill et

al. 2010). Bonamia-like microcell parasites were also visualised in Crassostrea gigas in SA (Diggles 2003),

but the identity of this parasite remains to be resolved. Clearly, further epidemiological and ultrastructural

study and multi-gene analysis will be needed to clarify the interrelationships between the various Bonamia

spp. in Australian waters (Lohrmann et al. 2009, Hill et al. 2010).


Bonamiasis has been detected in association with overt mortality in experimental stocks of Ostrea angasi in

Victoria, and in farmed O. angasi in WA, and with little or no mortality in Tasmania, WA, and NSW

(Corbeil et al. 2009). Bonamia-like parasites were also reported in C. gigas in SA with no mortality (Diggles

2003). Winter mortality due to B. roughleyi occurs on an irregular basis in NSW both spatially and

temporally from Port Stephens to the NSW/Victoria border (Adlard and Lester 1995). Using histology, the

mean prevalence of low intensity Bonamia sp. infections in healthy wild flat oysters O. angasi at 5 sites in

southern NSW was 26% (range 12.8%-44.1%) (Heasman et al. 2003), however histology is about half as

sensitive for detecting Bonamia infections compared to molecular diagnostic techniques (Diggles et al.

2003), and the actual prevalence of Bonamia sp. in wild populations of flat oysters in NSW therefore may be

closer to 50%. Overall prevalence of a Bonamia-like microcell in a histological survey of healthy C. gigas

from SA was 16%, ranging from 3.3% up to 66.4% at different sites (Diggles 2003), while Bonamia sp. in

healthy Tasmanian O. angasi is usually encountered in histological surveys at prevalences of 10% or less17.

A Bonamia roughleyi-like parasite observed in pearl oysters in WA is probably uninucleate stages of the

haplosporidian Minchinia occulta (see Bearham et al. 2009).

Bonamia spp. are only known to infect oysters. Molluscs are widely used as bait throughout Australia, with

“other shellfish” (not including abalone) being used by an average of 33-38% of recreational anglers in all

states except the NT (Kewagama Research 2002, 2007), with highest use being in SA (84-88% of anglers)

and Victoria (55-65%) of anglers (Kewagama Research 2002, 2007). However, the majority of the molluscs

commercially available are likely to be other bivalves such as live or frozen pipis (Plebidonax spp.) and

venerid clams (Katelysia spp., Anadara spp.). Live oysters are occasionally collected at the fishing site and

used as bait and/or berley by recreational fishers (B. Diggles, personal observations), but the quantities

17 http://www.fish.wa.gov.au/docs/pub/FHSlideofQuarter/200603.php?0408

_________________________________________________________________________________________________________________________________________________________________________________


198

www.digsfish.com

translocated by recreational fishers is likely to be extremely small. However, live, fresh and frozen cultured

flat and rock oysters are widely distributed as seafood and hence some oyster products could be diverted to

use as bait or berley around the country.

Bonamia spp. do not survive freezing, but can survive in dead oysters for at least 24 hours after the death of

the host, and in the water column for at least 4 days under appropriate conditions (B.K. Diggles and P.M.

Hine, personal observations, Diggles and Hine 2002). Taking into account the small quantities of oysters

used as bait or berley, as well as the fact that Bonamia spp. are reasonably prevalent in wild populations of

oysters in some areas of the country, the likelihood estimations for the occurrence of Bonamia spp. in these


Release assessment for infection of oysters with Bonamia spp., and/or unidentified microcells

Commodity type Live

oysters

Whole fresh

dead oysters

Frozen

whole

oysters

Frozen

oyster

meat

Oyster

shells

Likelihood of release High Moderate Negligible Negligible Negligible


Oysters in several regions of Australia are already at risk of exposure to Bonamia, however, translocation of

infected oysters from areas where Bonamia is enzootic via their use as bait or berley could transport the

parasite to new regions. Infection and establishment of Bonamia in new hosts would occur only if sufficient

viable infective particles were introduced into an area where susceptible oysters were present under suitable

environmental conditions for transmission. Bonamia causes disease only in flat and cupped oysters, and

therefore the risks of other molluscs becoming diseased appears low. Unlike other haplosporidians, the life

cycle of Bonamia exitiosa-like parasites is direct and horizontal. Spores have been detected in only one

Bonamia species, B. perspora from North Carolina, USA (Carnegie et al. 2006), and in all other Bonamia

species spores have not been observed and the microcell is known to act as the infective stage, with the

disease being transmitted by cohabitation (Diggles and Hine 2002, Hine et al. 2002b) or inoculation. Around

40% of microcell infective particles survive 48 hours in seawater at 18°C and 50% survive at least 4 days in

seawater at 4°C (Diggles and Hine 2002). Exposure to 103 - 105 B. exitiosa via the water causes mortalities

of 25 to 40% of O. chilensis within 18 weeks, while the 18-week LD50 for B. exitiosa was experimentally

determined to be approximately 2 x 105 microcells per oyster (Diggles and Hine 2002). This is slightly

higher than Hervio et al. (1995) found for Bonamia ostreae in Ostrea edulis (LD50 8 x 104 microcells per

oyster). Heavily infected O. chilensis with terminal Bonamia infections contain on average around 5 x 108

microcells (Diggles and Hine 2002, Cranfield et al. 2005), and this large difference between the infective

dose and parasite burdens in moribund oysters suggests the latter pose a serious risk to any uninfected

oysters that may occur nearby.

_________________________________________________________________________________________________________________________________________________________________________________


199

www.digsfish.com

Bivalves are efficient filter feeders and therefore are also efficient particle collectors, and can collect spores

and infective stages of not only bivalve disease agents (Barber and Ford 1992, Ford et al. 2009), but also

viruses and bacteria. Because of this, oysters are particularly susceptible to infection by protozoan infective

stages delivered via the water, and this is one of the reasons why movements of infected oysters are highly

likely to result in exposure and establishment of oyster pathogens in new areas. If infected live oysters or

even fresh dead oysters stored at low temperatures (e.g. on ice) for short periods of time were diverted for

use as bait or berley, and were used in areas frequented by wild oysters, a large number of infective particles

could be introduced into the water. It is not known whether exposure to low concentrations of infective

particles (< 103 /oyster) predisposes oysters to subclinical infection, but if infected bait or berley were used,

particularly in a small body of water (e.g. a rock pool), concentrations of infective stages may be sufficient

for an index case to occur. If the index case became diseased, transmission and further spread from the index

case would be possible because bivalves are such efficient particle collectors, and these disease agent can be

highly pathogenic under suitable conditions. Because of these reasons, translocated Bonamia is very likely

to become established in new oyster populations, as has been demonstrated several times in other parts of the

world (e.g. Howard 1994, Bishop et al. 2006). Taking these various factors into consideration, the risk of

exposure and establishment of Bonamia via use of bait and berley remains non-negligible, and the likelihood

of exposure and establishment of Bonamia in new oyster populations via translocation is considered to be

Moderate.


Although Bonamia is already present in populations of wild and cultured oysters in some regions of

Australia, it is possible that other regions remain free of infection at this time. Also, it is currently unclear

whether there are 2 or more species of Bonamia in Australia at this time, and the likely impacts of

introduction and spread of new species or strains of Bonamia into areas where other strains are already

endemic are unclear. There is certainly evidence that Bonamia can cause major disease outbreaks and

significant impacts on populations of both wild and cultured oysters. Bonamiosis is listed by the OIE and

NACA, and it is also listed as a reportable disease in all States (Table 3b). Hence the spread of Bonamia to

new areas is likely to adversely impact trade. Considering all of these factors, establishment of Bonamia in

new areas would have significant biological consequences for oyster aquaculture and could cause significant

economic harm together with significant environmental effects for ecosystems (where oysters act as

ecosystem engineers providing habitat and important bentho-pelagic coupling services) and wild oyster

fisheries that would be irreversible. It is therefore estimated that the consequences of introduction of

Bonamia into different parts of the Australian environment via use of infected bait would likely be

Moderate.


The unrestricted risk associated with Bonamia spp. is determined by combining the likelihood of entry and


Bonamia spp. exceeds the ALOP for live oysters and whole fresh dead oysters, suggesting that additional

risk management is required for this disease agent in these commodities.

_________________________________________________________________________________________________________________________________________________________________________________


200

www.digsfish.com

Risk estimate for infection of oysters with Bonamia spp., and/or unidentified microcells

Commodity type Live

oysters

Whole

fresh dead

oysters

Frozen

whole

oysters

Frozen

oyster meat

Oyster shells

Combined likelihood

of release and

exposure

Moderate Low Negligible Negligible Negligible

Consequences of

establishment


Risk estimation Moderate

risk

Low risk Negligible

risk

Negligible

risk

Negligible

risk

_________________________________________________________________________________________________________________________________________________________________________________


201

www.digsfish.com

5.32 Infection of molluscs with Haplosporidians

5.32.1 Aetiologic agent: Infection of molluscs by members of the genera Haplosporidium, Minchinia,

Urosporidium and other parasites (excluding Bonamia spp.) within the Phylum Haplosporidia.


5.32.3 Australias status: Haplosporidosis of molluscs has been reported in WA, and the disease is

reportable in all States except QLD and NSW (Table 3b).

5.32.4 Epidemiology

The Phylum Haplosporidia is composed of histozoic and coelozoic parasites that infect a wide variety of

freshwater and marine invertebrates worldwide. Infection of molluscs by haplosporidian parasites has

resulted in economically and ecologically significant mass mortalities in many parts of the world. For

example, in the USA, the haplosporidian parasite Haplosporidium nelsoni causes MSX disease which has

resulted in massive epizootics of eastern oysters (Crassostrea virginica) along the east coast of the United

States (Andrews 1968, 1996, Haskin and Ford 1982, Burreson et al. 2000). This parasite was probably

translocated to that region from Japan through imports of live seed oysters (Friedman 1996, Burreson et al.

2000, Kamaishi and Yoshinaga 2002). Reports of H. nelsoni from Crassostrea gigas in France (Renault et

al. 2000) provide further evidence this parasite has been moved with translocation of infected oysters, despite

the fact H. nelsoni is thought to have an indirect lifecycle that requires an intermediate host (Barber and Ford

1992, Ford et al. 2001).

In New Zealand, a new haplospridian parasite emerged in cultured abalone (Haliotis iris) resulting in

mortalities of up to 90% in affected raceways (Diggles et al. 2002, Hine et al. 2002a). The New Zealand

abalone parasite (NZAP) contained rickettsiales-like prokaryotes in its cytoplasm (Hine et al. 2002) and

molecular and ultrastructural analysis suggests that it falls at the base of the Phylum Haplosporidia (Reece et

al. 2004, Hine et al. 2009). The inability to transmit infection horizontally (Diggles et al. 2002) suggests that

an intermediate host is required for completion of the lifecycle of the NZAP. The lack of subsequent reports

of the NZAP in other abalone culture facilities or wild abalone, even after national abalone disease surveys

may suggest that it is either extremely rare, and/or that abalone may not be a normal host of the NZAP (B.K.

Diggles and P.M. Hine, personal observations).

In Australia, haplosporidians of the genus Haplosporidium and Minchinia have been associated with

sporadic but heavy mortalities in wild rock oysters (Saccostrea cuccullata) and hatchery reared pearl oysters

(Pinctada maxima) in Western Australia (Hine and Thorne 1998, 2000, 2002, Jones and Creeper 2006). The

parasite in pearl oysters was identified as Haplosporidium hinei and is considered to represent a serious risk

to the pearl industry (Bearham et al. 2008b, 2009). The parasite in the rock oysters has been associated with

mortalities of up to 80% and was identified as Minchinia occulta (see Bearham et al. 2007, 2008a, 2008c).

Mixed infections of M. occulta and H. hinei have also been recorded in P. maxima (see Bearham et al. 2009).

The uninucleate and multinucleate vegetative stages of M. occulta in S. cuccullata occur in the connective

tissues of the gills, mantle and around digestive diverticulae, multinucleate plasmodia with up to 25 nuclei

_________________________________________________________________________________________________________________________________________________________________________________


202

www.digsfish.com

occurred connective tissue adjacent to the digestive tract, while sporulation was confined to the connective

tissues around, but not within, the digestive tubules (Hine and Thorne 2002, Bearham et al. 2008c). Infected

oysters were in poor condition due to the absence of gonad tissue overlying the digestive tissue (Hine and

Thorne 2002).


Haplosporidosis has been detected in S. cuccullata, but not S. glomerata, near Exmouth in WA at

prevalences up to 28% at Varanus Island using histology (Hine and Thorne 2002). Few oysters were lightly

infected, with the disease appearing progressive and fatal (Hine and Thorne 2002). However, histology

appears to be only half as sensitive as detecting infections of M. occulta compared to molecular diagnostic

techniques (see Bearham et al. 2008a), suggesting the prevalence of the parasite in wild populations of S.

cuccullata may exceed 50% at times, though at other times it is undetectable in oysters from the same

locations where heavy mortalities occurred in the past (Bearham et al. 2008c). Of the various species of

molluscs used as bait in Australia, to date, haplosporidians have only been recorded in rock oysters.

However, disease surveillance of mollusc populations in Australia is mainly passive, and many outbreaks of

disease in wild populations of molluscs go unobserved or unreported (Cranfield et al. 2005). The emergence

of haplosporidian diseases in species such as S. cuccullata and P. maxima and occurrence of Bonamia-like

parasites (which could also be uninucleate stages of haplosporidians such as M. occulta, see Bearham et al.

2007, 2008a, 2008b, 2009) in species such as C. gigas in SA (Diggles 2003) and molluscs in other regions of

Australia demonstrate that the full range of haplosporidian infections that are present in the various species

of molluscs used as bait remains to be determined.

Molluscs are widely used as bait throughout Australia, with “other shellfish” (not including abalone) being

used by an average of 33-38% of recreational anglers in all states except the NT (Kewagama Research 2002,

2007), with highest use being in SA (84-88% of anglers) and Victoria (55-65%) of anglers (Kewagama

Research 2002, 2007). Only 11–13% of recreational fishers in WA used “other shellfish” for bait

(Kewagama Research 2002, 2007). The majority of the molluscs commercially available are likely to be

other bivalves such as live or frozen pipis (Plebidonax spp.) and venerid clams (Katelysia spp., Anadara

spp.), from which haplosporidians have not been detected to date. Live rock oysters are occasionally

collected at the fishing site and used as bait and/or berley by recreational fishers (B. Diggles, personal

observations), but the quantities translocated by recreational fishers is likely to be extremely small.

However, live, fresh and frozen cultured rock oysters are widely distributed as seafood and hence some

oyster products could be diverted to use as bait or berley around the country. There is currently no culture of

S. cuccullata in WA from the regions where haplosporidians are known to occur.

Vegetative stages of haplosporidian parasites are unlikely to survive freezing, however the tolerance of spore

stages to freezing appears to be unknown. It is likely that vegetative stages can survive in dead oysters for

some unknown period of time that would be extended as storage temperature decreased. Taking into account

the small quantities of oysters used as bait or berley, as well as the fact that haplosporidian parasites are

sporadically prevalent in wild populations of oysters in some areas of the country, the likelihood estimations

for the occurrence of haplosporidian parasites in these commodities are listed below.

_________________________________________________________________________________________________________________________________________________________________________________


203

www.digsfish.com

Release assessment for infection of molluscs with haplosporidians

Commodity type Live

molluscs

Whole fresh

dead

molluscs

Frozen

whole

molluscs

Frozen

mollusc

meat

Mollusc

shells

Likelihood of release Moderate Very low Negligible Negligible Negligible


Oysters in the northwest region of Australia are already at risk of exposure to haplosporidians, however,

translocation of infected oysters from areas where Haplosporidium and Minchinia are enzootic via their use

as bait or berley could transport these parasites to new regions. Infection and establishment of these

parasites in new hosts would occur only if sufficient viable infective particles were introduced into areas

where susceptible intermediate hosts and oyster final hosts were present under suitable environmental

conditions for transmission. Minchinia occulta is only known to infect S. cuccullata at this time, however as

haplosporidians appear to require an intermediate host to complete their lifecycle (Haskin and Andrews

1988, Ford et al. 2001, Diggles et al. 2002), in absence of knowledge of the identity of the intermediate

host(s) it is not possible at this time to determine whether other invertebrates may be susceptible to disease.

The fact that S. cuccullata, but not S. glomerata were infected (Hine and Thorne 2002) suggests that these

parasites may have high host specificity, which may reduce the chances of exposure and establishment.

However Hine and Thorne (2002) pointed out that S. glomerata was sampled only from areas where S.

cuccullata were uninfected, hence it remains possible that S. glomerata is also susceptible to M. occulta.

The inability to transmit haplosporidians directly by cohabitation or injection of spores suggests they have an

indirect lifecycle requiring an alternate host. The earliest vegetative stages of Haplosporidium nelsoni are

found in the epithelia of the gills and palps, suggesting that the infective stage is waterborne and can be

easily spread (Haskin and Andrews 1988). Neither the infective stage nor the mode of transmission has ever

been identified (Powell et al. 1999, Sunila et al. 2000), although it is known that the infective stage for H.

nelsoni can pass through a 150 µm filter , but not a 1 µm filter followed by UV irradiation (Ford et al. 2001).

At certain times of the year in areas where haplosporidians are endemic, large numbers of spores occur in the

water column and these can be concentrated within the digestive tract of filter feeding bivalves, and therefore

bivalve movements can transfer spores to new areas (Barber and Ford 1992). Because an intermediate host

is presumably needed in order to complete the lifecycle of haplosporidians, the exposure pathway required

for transmission remains unknown, as does important information such as the minimum infective dose

required for an index case to occur. However, if an index case occurred, these disease agents are highly

pathogenic and it would be likely that the infected bivalve would become diseased, after which transmission

and further spread from the index case would be possible. Because of this, the current restricted distribution

of these parasites may be due to the fact that their intermediate hosts may also be restricted in distribution.

However, the fact that other species of Haplosporidians (e.g. Haplosporidium nelsoni) have been

translocated and established infections in new regions (Friedman 1996, Burreson et al. 2000, Renault et al.

2000), suggests that some of the intermediate hosts may be widespread and/or ubiquitous (e.g. planktonic

copepods), or that these parasites may have lower host specificity for the intermediate host. Taking these

various factors into consideration, the risk of exposure and establishment of haplosporidians via use of bait

_________________________________________________________________________________________________________________________________________________________________________________


204

www.digsfish.com

and berley remains non-negligible, and the likelihood of exposure and establishment of haplosporidians in

new oyster populations via translocation is considered to be Low.


Although haplosporidian parasites are already present in populations of wild molluscs in some regions of

Australia, other regions appear free of infection at this time. There is evidence that haplosporidians can

cause major disease outbreaks and significant impacts on populations of both wild and cultured molluscs in

Australia and overseas. Haplosporidosis is no longer listed by the OIE and NACA, but these disease agents

remain listed as a reportable disease in all States except QLD and NSW (Table 3b). Hence the spread of

haplosporidian parasites to new areas is likely to adversely impact trade. Considering all of these factors,

establishment of haplosporidians in new areas would potentially have significant biological consequences

and could cause significant economic harm for mollusc aquaculture industries, together with potentially

significant and irreversible environmental effects for ecosystems (where oysters act as ecosystem engineers

providing habitat and important bentho-pelagic coupling services) and wild mollusc fisheries. It is therefore

estimated that the consequences of introduction of haplosporidians into different parts of the Australian



The unrestricted risk associated with haplosporidians is determined by combining the likelihood of entry and


haplosporidians exceeds the ALOP for live molluscs, suggesting that additional risk management is requires

for these disease agents in these commodities.

Risk estimate for infection of molluscs with haplosporidians

Commodity type Live

molluscs

Whole

fresh dead

molluscs

Frozen

whole

molluscs

Frozen

mollusc

meat

Mollusc

shells

Combined likelihood

of release and

exposure

Low Very low Negligible Negligible Negligible

Consequences of

establishment


Risk estimation Low risk Very low

risk

Negligible

risk

Negligible

risk

Negligible

risk

_________________________________________________________________________________________________________________________________________________________________________________


205

www.digsfish.com

5.33 Infection of oysters and annelids with Marteilia sydneyi (QX disease)

5.33.1 Aetiologic agent: Marteilia sydneyi, a parasite (Phylum Paramyxea) that infects oysters Saccostrea

glomerata, and some polychaete annelids, including Nephtys australiensis and Perinereis nuntia.


5.33.3 Australias status: Marteilia sydneyi infections have been reported in QLD, NSW and WA, and the

disease is reportable in all States except QLD and the ACT (Table 3b).

5.33.4 Epidemiology

Members of the Phylum Paramyxea are parasites of marine invertebrates that are characterised by the

formation of spores via internal cleavage of sporangia within plasmodia (Desportes and Perkins 1990).

Paramyxean parasites of the genus Marteilia have caused significant disease and economic impacts on oyster

culture in several regions of the world (Berthe et al. 2004). Marteilia refringens devastated the flat oyster

(Ostrea edulis) industry in France beginning in the late 1970’s (Grizel et al. 1974, Grizel 1985), and this

parasite also infects other bivalves including mussels (Robeldo and Figueras 1995, Longshaw et al. 2001),

and razor clams (Lopez Flores et al. 2008). Marteilia sydneyi is responsible for QX disease that has caused

massive losses (up to 98% mortality) in wild and cultured Sydney rock oyster Saccostrea glomerata along

the east coast of Australia from Great Sandy Straits in QLD to the NSW/Victoria border since the early

1970’s (Wolf 1972, Perkins and Wolf 1976, Lester 1989, Roubal et al. 1989b, Adlard and Ernst 1995).

QX disease is due to massive infection of the digestive gland with M. sydneyi (Kleeman et al 2002a, 2002b).

Oysters appear to be exposed to the infective stage for only a short period, usually after heavy rainfall in the

summer months, and under suitable conditions for disease development oyster deaths increase in late

summer and autumn (Lester 1989, Roubal et al. 1989b, Wesche 1995). The parasite remains present in

healthy oyster populations at low prevalences in most estuaries (Adlard and Nolan 2008), and in estuaries

where QX disease occurs, outbreaks do not occur every year (Butt et al. 2006). This is probably because the

onset of QX disease is related to immunosuppression of the host (Peters and Raftos 2003, Butt and Raftos

2007), due to reduced salinity (Butt et al. 2006, Green and Barnes 2010) and as yet unidentified water born

contaminants carried in runoff (Butt and Raftos 2007). Oysters that carry low levels of M. sydneyi infections

can shed the parasite and make a full recovery (Roubal et al. 1989b).

The lifecycle of Marteilia spp. is indirect (Roubal et al. 1989b) and requires at least one alternate host.

Marteilia refringens in France has a lifecycle that includes the planktonic copepod Paracartia grani, females

of which can become infected through contact with spore stages (see Audemard et al. 2001, 2002, 2004).

Marteilia sydneyi also has an indirect lifecycle which appears to include polychaete worms of the Family

Nephtyidae as one alternate host (see Adlard and Nolan 2008, Cribb 2010). The polychaete Nephtys

australiensis harboured developmental stages of M. sydneyi, grows to 85 mm and is described as being more

common in muddy rather than sandy sediments (Rainer and Hutchings 1977). The increased virulence of M.

sydneyi in degraded estuaries compared to historical times may therefore be due to a combination of

increased immunosuppression of the host due to declining water quality together with increased abundance

_________________________________________________________________________________________________________________________________________________________________________________


206

www.digsfish.com

of polychaete intermediate hosts that are favored by sedimentation, eutrophication and other anthropogenic

changes derived from catchment development (B.K. Diggles, personal observations, Green et al. 2011).


Marteilia sydneyi has been detected in association with mortality in wild and cultured S. glomerata in QLD

and NSW, as well as at low prevalence in the absence of disease in WA (Hine and Thorne 2000). The

parasite is also known to infect polychaetes within the Family Nephtyidae, and many other species of filter

feeding invertebrates in estuaries are likely to temporarily accumulate Marteilia spores in their digestive tract

(see Audemard et al. 2002), and hence could act as mechanical vectors.



2007), with highest use being in SA (84-88% of anglers) and Victoria (55-65%) of anglers (Kewagama

Research 2002, 2007). However, the majority of the molluscs commercially available are likely to be other

bivalves such as live or frozen pipis (Plebidonax spp.) and venerid clams (Katelysia spp., Anadara spp.)

which are unlikely to harbour M. sydneyi (though the required epidemiological surveys needed to rule this

out have not been done). Live oysters are occasionally collected at the fishing site and used as bait and/or

berley by recreational fishers (B.K. Diggles, personal observations), but the quantities translocated by

recreational fishers is likely to be extremely small. However, live, fresh and frozen oysters are widely

distributed as seafood and hence some oyster products could be diverted to use as bait or berley around the

country. Marine annelids (or “worms”, mainly polychaetes) are widely used as bait by recreational fishers in

every state except the NT (Kewagama Research 2007), with highest use occurring in QLD (40-42% of

anglers), NSW (32-38% of anglers) and Victoria (27-33% of anglers) (Kewagama Research 2002, 2007). A

reasonably large proportion of these annelids are sold live, though a larger proportion is likely to be frozen or

freeze dried (B.K. Diggles, personal observations).

Spores of M. sydneyi survive freezing, with 20% survival after 7 days and 5.8% of spores viable after 220

days at -20°C (Wesche et al. 1999). Spores can survive in the water for up to 35 days at 15 °C and 34 ppt

salinity (Wesche et al. 1999), but do not survive passage through the gut of fish or birds (Wesche et al.

1999). It is not known whether spores or infective stages of M. sydneyi can survive freeze drying. Taking

into account the small quantities of oysters and the large quantities of polychaetes used as bait or berley, as

well as the fact that Marteilia sydneyi is highly prevalent in wild populations of oysters in some areas of the

country, the likelihood estimations for the occurrence of M. sydneyi in these commodities are listed below.

Release assessment for infection of oysters and annelids with Marteilia sydneyi

Commodity type Live

oysters

Whole

fresh dead

oysters

Frozen

whole

oysters

Live

annelids

Fresh

dead

annelids

Frozen

annelids

Freeze

dried

annelids

Likelihood of

release

High Moderate Low High Moderate Low Low

_________________________________________________________________________________________________________________________________________________________________________________


207

www.digsfish.com


Oysters in the eastern and northwestern regions of Australia are already at risk of exposure to M. sydneyi,

however, translocation of infected oysters or annelids from areas where M. sydneyi is enzootic via their use

as bait or berley could transport these parasites to new regions. Infection and establishment of these

parasites in new hosts would occur only if sufficient viable infective particles were introduced into areas

where susceptible intermediate hosts and oyster final hosts were present under suitable environmental

conditions for transmission. At this time Marteilia sydneyi is only known to infect S. glomerata and

polychaetes of the Family Nephtyidae (see Adlard and Nolan 2008), however these polychaetes are

ubiquitous in the Australian environment (Rainer and Hutchings 1977), meaning that the indirect lifecycle of

M. sydneyi may not be a barrier to its wider dissemination through movements of oysters. Polychaetes are

known natural hosts for other species of paramyxeans, such as Paramyxoides nephtys in Nephtys caeca (see

Larsson and Koie 2005), and therefore it is possible that other (currently undescribed) paramyxeans may also

parasitize polychaete annelids in Australia.

The earliest vegetative stages of M. sydneyi are found in the epithelia of the gills and palps, suggesting that

the infective stage is waterborne and can be easily spread (Kleeman et al. 2002a). Neither the infective stage

nor the mode of transmission have been identified to date, however at certain times of year in areas where M.

sydneyi is endemic, large numbers of infective stages and spores must occur in the water column and

sediments (Roubal et al. 1989b) and these can be concentrated within the digestive tract of filter feeding

bivalves, which are efficient particle collectors, as well as many other species of filter feeding invertebrates

(see Audemard et al. 2002), which could act as mechanical vectors. Thus movements of polychaete alternate

hosts as well as other invertebrate mechanical vectors could translocate infective stages of M. sydneyi to new

locations which may contain susceptible oyster hosts.

Because the lifecycle of M. sydneyi has not yet been completed experimentally, the exact exposure pathway

required for transmission remains unknown. Hence important information, such as the minimum infective

dose required for an index case to occur, is not available. Oysters that carry low levels of M. sydneyi

infections can shed the parasite and make a full recovery (Roubal et al. 1989b), hence ambient environmental

conditions as well as the immune status of the host will both influence whether the disease agent would

establish once an index case occurred. In any case, the current restricted distribution of these parasites does

not seem to be due to restricted distribution of the likely intermediate hosts (Family Nephtyidae), as these

annelids are ubiquitous in the Australian environment, being more common in muddy rather than sandy

sediments (Rainer and Hutchings 1977). Taking these various factors into consideration, the risk of exposure

and establishment of M. sydneyi via use of bait and berley remains non-negligible, and the likelihood of

exposure and establishment of M. sydneyi in new oyster populations via translocation is considered to be

Low.


Although M. sydneyi is already present in populations of wild molluscs in some regions of Australia, other

regions appear free of infection at this time. The intermediate hosts for M. sydneyi are widespread and

therefore the indirect lifecycle may not restrict dispersal of the parasite. There is evidence that M. sydneyi

can cause major disease outbreaks and significant impacts on populations of both wild and cultured molluscs

_________________________________________________________________________________________________________________________________________________________________________________


208

www.digsfish.com

in Australia. Marteilia sydneyi is no longer listed by the OIE and NACA, but these disease agents remain

listed as a reportable disease in all States except QLD and the ACT (Table 3b). Hence the spread of M.

sydneyi to new areas is likely to adversely impact trade. Considering all of these factors, establishment of M.

sydneyi in new areas would potentially have significant biological consequences and could cause significant

economic harm for mollusc aquaculture industries, together with potentially significant and irreversible

environmental effects for ecosystems (where oysters act as ecosystem engineers providing habitat and

important bentho-pelagic coupling services) and wild mollusc fisheries. It is therefore estimated that the

consequences of introduction of M. sydneyi into different parts of the Australian environment via use of

infected bait would likely be Moderate.


The unrestricted risk associated with M. sydneyi is determined by combining the likelihood of entry and


M. sydneyi exceeds the ALOP for live oysters and annelids, and whole fresh oysters and annelids, suggesting

that additional risk management is required for this disease agent in these commodities.

Risk estimate for infection of oysters and annelids with Marteilia sydneyi

Commodity type Live

oysters

Whole

fresh dead

oysters

Frozen

whole

oysters

Live

annelids

Fresh

dead

annelids

Frozen

annelids

Freeze

dried

annelids

Combined

likelihood of

release and

exposure

Low Low Very low Low Low Very low Very low

Consequences of

establishment

Moderate Moderate Moderate Moderate Moderate Moderate Moderate

Risk estimation Low risk Low risk Very low

risk

Low risk Low risk Very low

risk

Very low

risk

_________________________________________________________________________________________________________________________________________________________________________________


209

www.digsfish.com

5.34 Infection of molluscs with Perkinsus olseni

5.34.1 Aetiologic agent: Perkinsus olseni and other parasitic protozoa within the Family Perkinsidae.


5.34.3 Australias status: Perkinsus olseni infections have been reported from all states except Tasmania,

and the disease is reportable in all States except the ACT (Table 3b).

5.34.4 Epidemiology

Members of the genus Perkinsus within the Family Perkinsidae are closely related to dinoflagellates (Reece

et al. 1997) and these obligate protistan parasites are known to infect a wide range of marine molluscs in

many regions of the world (Villalba et al. 2004). The life cycle of Perkinsus spp. involves vegetative

proliferation within the host by trophozoites that undergo successive bipartitioning. Other stages that have

been observed include hypnospores, zoosporangia and zoospores, the latter which are probably natural

dispersal and infective stages (Goggin et al. 1989). When host tissues infected by Perkinsus spp. are

incubated in fluid thioglycollate medium (FTM), the trophozoites enlarge and develop a thick cell wall,

becoming easy to visualise after staining with lugols iodine (Ray 1966). When these enlarged hypnospore

stages are transferred into seawater, they form zoosporangia and production of hundreds to thousands of

zoospores occurs within the original cell wall (Villalba et al. 2004). The biflagellated zoospores 3-5 µm in

size leave the zoosporangium through discharge tubes and enter the water to reinfect new hosts via the gills,

palps and digestive tract (Villalba et al. 2004). Infection of susceptible molluscs can occur horizontally

through the water by cohabitation via contact with zoospores, but trophozoites and hypnospores have also

been shown experimentally to cause infection (Goggin et al. 1989), and the disease can be transmitted via

vectors such as ectoparasitic snails (White et al. 1987).

The first described species of Perkinsus was Perkinsus marinus, the agent responsible for “Dermo disease”

that was associated with significant mortality events in oyster (Crassostrea virginica) populations along the

eastern and southern coast of the United States since the late 1940’s (Ray 1996, Andrews 1996). Distribution

of P. marinus in oysters in these regions and also in Mexico is variable depending on environmental factors,

with persistence of the parasite being favoured by high water temperatures (>20°C) and high salinity (>15

ppt) (Chu and Greene 1989). The second Perkinsus species described was Perkinsus olseni, which was

originally identified from blacklip abalone Haliotis rubra near Port Lincoln in Spencer Gulf, SA (Lester and

Davis 1981). P. olseni was subsequently associated with severe mortalities in greenlip abalone Haliotis

laevigata around 140 km away in the western side of Gulf St Vincent (Lester 1986, O’Donoghue et al. 1991;

Goggin and Lester 1995) in SA, and more recently the same parasite has been associated with significant

mortality events in abalone along the central and southern coast of NSW (Lester and Hayward 2005). The

presence of P. olseni in infected abalone of all sizes was characterized by the presence of macroscopic

necrotic nodules (0.5-8.0 mm in diameter) in the adductor muscles and mantle (O’Donoghue et al. 1991),

and the disease process is facilitated by high water temperatures >20°C (Lester 1986, Lester and Hayward

2005). Perkinsus-like parasites have also been reported from 30 out of 84 species of molluscs examined

from the Great Barrier Reef (Goggin and Lester 1987), from pearl oysters Pinctada maxima from Torres

_________________________________________________________________________________________________________________________________________________________________________________


210

www.digsfish.com

Strait (Norton et al. 1993) as well as several species of molluscs from WA (Hine and Thorne 2000). To date

the Perkinsus –like parasites from Australian molluscs have all been identified as P. olseni (see Murrell et al.

2002, Lester and Hayward 2005). Perkinsus olseni has also been recorded in many other regions worldwide,

including in cockles (Austrovenus stutchburyi) in the North Island of New Zealand, where its distribution is

probably limited by temperature (Hine and Diggles 2002b). Perkinsus olseni has also been associated with

mass mortalities of the Manila clam Tapes philippinarum in South Korea and has been detected in clams

from Japan, China , Vietnam, Europe and Uraguay (Villalba et al. 2004, Park et al. 2005). Four other species

of Perkinsus are currently recognised, including, including P. qugwadi, P. chesapeaki, P. andrewsi and P.

mediterraneus, all of which are parasites of molluscs (Villalba et al. 2004).


In Australia Perkinsus spp. have been detected in two areas of SA from abalone H. laevigata, H. rubra, H.

cyclobates, H. scalaris, cockles Barbatia pistachia, Katelysia rhytiphora, razor shells Pinna bicolor, and

scallops Chlamys bifrons, in NSW from H. rubra, H. laevigata and H. roei , in QLD from pearl oysters

Pinctada maxima and 23 families of bivalves from the Great Barrier Reef, in Victoria from O. angasi, and

from north west WA in Pinctada albina, rock oysters S. glomerata and S. cuccullata, hammer shells

(Malleus meridianus), and razor shells (Isognomon isognomon, Pinna bicolour and P. deltoides) (Lester and

Davis 1981, Goggin and Lester 1987, 1995, Norton et al. 1993, Hine and Thorne 2000, Lester and Hayward

2005). Greenlip abalone (H. laevigata) appear to be particularly susceptible to P. olseni infection, while

blacklip abalone (H. rubra) appear to be relatively resistant to infection (O’Donoghue et al. 1991, Lester and

Hayward 2005).

Perkinsus olseni infects many species of molluscs used as bait in Australia, particularly abalone and venerid

clams (or cockles, Anadara spp., Katelysia spp.), but this parasite has not yet been reported from pipis

(Plebidonax spp.). However, disease surveillance of mollusc populations in Australia is mainly passive,

hence the full range of mollusc species that can carry Perkinsus infections remains to be determined.



2007), with highest use being in SA (84-88% of anglers) and Victoria (55-65% of anglers) (Kewagama

Research 2002, 2007). The majority of the molluscs commercially available are likely to be bivalves such as

live or frozen pipis (Plebidonax spp.) and venerid clams (Katelysia spp., Anadara spp.). Live pipis and

venerid clams are commonly collected at the fishing site and used as bait and/or berley by recreational

fishers in many parts of the country (B. Diggles, personal observations), and the quantities of these shellfish

that are translocated small distances (< 100 km) by recreational fishers is likely to be significant. Fresh and

frozen pipis and venerid clams are also widely distributed as seafood and bait, hence some of these could

also be diverted to use as bait or berley around the country.

Vegetative stages of P. olseni are extremely resistant and can survive in dead hosts at 4°C and 0°C for at

least 24 hours with high (22-27%) survival, while they also survive freezing (-60°C) in both dried abalone

tissue (19% survival after 28 days) and in abalone tissue stored in seawater (37% survival after 197 days)

(Goggin et al. 1990). Zoospores can survive in seawater at temperatures of 20-25°C for up to 28 days (Chu

and Greene 1989). It is likely that vegetative stages can survive in dead molluscs at normal environmental

temperatures (15-25°C) for the 2-4 days it takes for hypnospores to form and complete zoosporogenesis (Chu

_________________________________________________________________________________________________________________________________________________________________________________


211

www.digsfish.com

and Greene 1989), as shown by natural transmission of Perkinsus spp. after the death of its host (Villalba et

al. 2004). Taking into account the large quantities of susceptible molluscs used as bait or berley, as well as

the fact that P. olseni is sporadically prevalent in wild populations of molluscs in several areas of the

country, the likelihood estimations for the occurrence of P. olseni in these commodities are listed below.

Release assessment for infection of molluscs with Perkinsus olseni

Commodity type Live

molluscs

Whole fresh

dead molluscs

Frozen whole

molluscs

Frozen

mollusc meat

Mollusc

shells

Combined likelihood of

release and exposure

High High Moderate Moderate Negligible


Molluscs in several regions of Australia are already at risk of exposure to P. olseni, however, translocation of

infected molluscs from areas where P. olseni is enzootic via their use as bait or berley could transport these

parasites to new regions. Infection and establishment of these parasites in new hosts would occur only if

sufficient viable infective particles were introduced into areas where susceptible hosts were present under

suitable environmental conditions for transmission. At this time P. olseni is known to infect a wide variety

of molluscs (Goggin et al. 1989), and it is known that all life stages of the parasite are potentially infective,

however as for P. marinus, it is not known which stage is most effective or what is the principal stage for

transmitting the disease in the natural environment (Villalba et al. 2004). Viable P. marinus cells are

released from live infected oysters through diapedesis and in faeces (Bushek et al. 2002), but host death does

not prevent transmission since hypnospore formation would allow further transmission either through the

hypnospores themselves, or by their development into zoosporangia giving rise to infective zoospores

(Villalba et al. 2004). Under natural circumstances, susceptible molluscs need to be in close proximity to

diseased molluscs for horizontal transmission to occur, possibly due to the fact that a relatively high numbers

of infective stages (zoospores, and/or trophozoites and/or hypnospores) are required to initiate infection

(around 1 x 105 infective stages/oyster for P. marinus, see Andrews 1996). However, each zoosporangia can

liberate up to 2000 zoospores (Andrews 1996), and thus dead molluscs can potentially liberate large numbers

of infective stages, which can then be concentrated within the digestive tract of filter feeding bivalves as well

as many other species of filter feeding invertebrates which could act as mechanical vectors. The infective

stages of Perkinsus spp. survive passage through the gut of scavenger teleosts (Hoese 1964) and the origin of

the parasite does not affect its ability to infect molluscs from different localities (Goggin et al. 1989).

Environmental conditions as well as host immune status will play important roles in transmission and

establishment of P. olseni in an index case. Water temperatures above 20°C appear to be required to

increase the chances of infection, and transmission dos not appear to occur in water of low salinity (<10 ppt).

Further, molluscs that carry low levels of Perkinsus spp. infection can shed the parasite during the winter

months and possibly eliminate the infection (Goggin and Lester 1995). Taking these various factors into

_________________________________________________________________________________________________________________________________________________________________________________


212

www.digsfish.com

consideration, the risk of exposure and establishment of P. olseni via use of bait and berley remains non-

negligible, and the likelihood of exposure and establishment of P. olseni in new mollusc populations via

translocation is considered to be High.


Although P. olseni is already present in populations of wild molluscs in some regions of Australia, the

distribution of this parasite is often sporadic at smaller scales, and several regions may remain free from

infection at this time, including Tasmania where, like the South Island of NZ, water temperatures may be too

cold for P. olseni at this time. There is evidence that P. olseni can cause significant disease outbreaks as well

as causing sub-lethal disease (reduction of fecundity, growth, and condition) which has significant impacts at

the population level in both wild and cultured molluscs. Perkinsus olseni is listed by the OIE and NACA as

a reportable disease, and remains listed as a reportable disease in all States except the ACT (Table 3b).

Hence the spread of P. olseni to new areas is likely to adversely impact trade. Considering all of these

factors, establishment of P. olseni in new areas would potentially have significant biological consequences

and could cause significant economic harm for mollusc aquaculture industries, together with potentially

significant and irreversible environmental effects for ecosystems and wild mollusc fisheries. It is therefore

estimated that the consequences of introduction of P. olseni into different parts of the Australian



The unrestricted risk associated with Perkinsus olseni is determined by combining the likelihood of entry and


Perkinsus olseni exceeds the ALOP for live molluscs, whole fresh dead molluscs and whole frozen molluscs

and mollusc meat, suggesting that additional risk management for this disease agent is required for these

commodities.

Risk estimate for infection of molluscs with Perkinsus olseni

Commodity type Live molluscs Whole fresh

dead molluscs

Frozen whole

molluscs

Frozen

mollusc meat

Mollusc

shells

Combined likelihood

of release and

exposure

High High Moderate Moderate Negligible

Consequences of

establishment


Risk estimation Moderate Risk Moderate Risk Moderate Risk Moderate Risk Negligible

risk

_________________________________________________________________________________________________________________________________________________________________________________


213

www.digsfish.com

5.35 Infection of molluscs with spionid mudworms

5.35.1 Aetiologic agent: Spionid polychaetes (including the genera Boccardia and Polydora) that infect the

shells of molluscs.


5.35.3 Australias status: Mudworm infections have been reported from all States, and certain species of

these agents are reportable in SA (Table 3b).

5.35.4 Epidemiology

Spionid polychaetes are predominantly free living organisms that are found in soft muddy estuarine

sediments worldwide, however several species are commonly associated with molluscs, using the shells of

bivalve molluscs and abalone as settlement substrates (Read 2010, Walker 2011) as they feed on suspended

or resuspended particles or plankton (Dauer et al. 1981). Most spionids infesting molluscs are

ectocommensal, the planktonic larvae settling on the outer shell, however some larvae can settle inside the

mantle cavity and/or on the edge of the shell lip, with the growing worm establishing a cover of mucus and

debris while enlarging a burrow on the inner surface of the shell valve in the extrapallial space, accumulating

sediment and detritus inside their burrow as the mollusc covers it with nacre, resulting in shell blistering

(thus the name “mudworm”) (Read 2010). At least 37 species of spionids have been recorded in Australia to

date, with at least 12 species of Polydora and 10 species of Dipolydora occurring on the east coast alone (see

Walker 2009, 2011). The main species that are usually reported to be problematic in mollusc aquaculture

include Polydora websteri, P. haswelli and P. hoplura, Boccardia knoxi and B. chilensis) (see Nell 2001,

Lleonart et al. 2003a, 2003b). Low intensity infections are innocuous and usually confined to the shell,

however some species may cause unsightly mud blisters in the shell and abscesses in the adductor muscle if

the blister contacts the tissue (Whitelegge 1890). It is notable that many of the early museum specimens

originally identified as P. websteri from Australian oysters were, upon re-examination, other species such as

various Dipolydora spp., while some specimens identified as P. polybranchia by Haswell (1885) and

Whitelegge (1890) from S. glomerata were re-identified as Boccardia polybranchia and P. wellingtonensis

(see Walker 2009, 2011).

Prevalence and intensity of mudworm infestation vary considerably with local water quality and growing

height of sedentary species such as oysters. Light mudworm infection rarely causes mortalities and infected

oysters can usually be marketed, however mud blisters may interfere with shucking and reduce the

commercial value of oysters to be served on the half-shell (Nell 2001). Prevalence and intensity of infection

increases in the vicinity of muddy substrates (Whitelegge 1890), and infections can be reduced by off bottom

bivalve culture techniques (at least 0.5 m above the mud substratum), preferably at heights that dry out the

mollusc for at minimum 2 hours in each tidal cycle (Nell 2001).

While trans-Tasman exports of live oysters from New Zealand were commonplace during the late nineteenth

century, there is no evidence that mudworms were problematic in New Zealand at that time (Read 2010).

The earliest reports of mudworm in New Zealand only date from the early 1970s and only from northern

_________________________________________________________________________________________________________________________________________________________________________________


214

www.digsfish.com

New Zealand, whereas a century earlier at least one of these pest worms had become widespread along

eastern Australian coasts (Nell 2001, Ogburn et al. 2007, Read 2010). This suggests that “mudworm

disease” of wild and cultured sub tidal oysters on Australias east coast from around 1870 onwards (Roughley

1939) may not have been due to introduction of exotic mudworms from New Zealand (as hypothesized by

some authors, see Ogburn et al. 2007). Instead, proliferation of native mudworm species may have occurred

due to increased eutrophication and organic enrichment of Australian estuaries, a process that began to be

noticeable after floods from 1870 onwards due to catchment clearing and development (McCulloch et al.

2003). Mudworms are very abundant in muddy tidal flats compared to clean sandy ones, as mudworm

settlement is stimulated by high microbial counts associated with muddy sediments (Sebesvari et al. 2006).

This suggests that organic enrichment/eutrophication and sedimentation (the first anthropogenic changes that

tend to occur in estuaries following on from any extensive development in the catchment, see Paterson et al.

2003) will promote increased abundance of mudworms (Nell 2001). In effect, Haswell (1885) noted this by

stating “some local circumstances, such as muddiness of the water produced by increasing traffic, tend to

decrease the vital powers of the oysters and thus favour the inroads of the parasites”. The subsequent

disappearance of sub tidal oyster beds throughout much of the east coast (Ogburn et al. 2007) is more likely

due to spatfall failure (spat set does not occur on dirty surfaces covered in sediment trapped by the algae

generated by organic enrichment), as well as QX disease in recent years (B.K. Diggles, unpublished data).

Mortalities of S. glomerata reported in Moreton Bay after floods in the mid 1890’s were attributed to

“mudworm disease” (Brisbane Courier 1898), however today, while heavy mudworm infections reduce the

growth rate of cupped oysters, they are seldom fatal (Read 2010, B.K. Diggles, personal observations),

although hyperinfections of P. hoplura and B. knoxi in confined rearing systems have caused mortalities of

up to 50% in cultured abalone (Lleonart et al. 2003a, 2003b). Indeed, the correlation with flooding suggests

it is possible that the mortalities in Moreton Bay in the mid and late 1890’s (Brisbane Courier 1898,

Lergessner 2006) were actually the first epizootics due to QX disease. Oyster farmers and scientists in the

late 1800’s had a rudimentary ability to diagnose oyster pathogens, and they would have been far more likely

to blame the very visible mudworm for any oyster deaths, rather than an invisible protozoan (Read 2010).

However, Whitelegge (1890) described mudworm hyperinfections in subtidal oysters resulting in mortality,

which is similar to that described by Lleonart et al. (2003a, 2003b), hence it remains plausible that the

mudworm epizootics of the late 19th and early 20th centuries were indicators of structural changes to the biota

of the affected estuaries due to increasing sediment loads bought down after floods. Given the existence of

many species of endemic spionid polychaetes (Walker 2011), increased sedimentation and organic

enrichment in estuaries resulting in proliferation of endemic polychaetes (in the form of both mudworms and

the polychaete intermediate hosts of M. sydneyi) appears a more parsimonious explanation for the emergence

of both mudworm and QX disease, with QX becoming more prominent since the 1970s due to further

declines in water quality causing more frequent immunosuppression of the oysters (Peters and Raftos 2003).


Several species of spionid mudworms appear to be widespread throughout Australia, and indeed many

species of concern to mollusc farmers have cosmopolitan distributions (Walker 2009, 2011), although some

endemic species may have restricted distributions, such as B. knoxi, which may prefer cooler waters as it has

been recorded in Australia only from south Western Australia and Tasmania (Walker 2011). These worms

require specialist knowledge to accurately identify them, and thus the exact distribution of the various species of

_________________________________________________________________________________________________________________________________________________________________________________


215

www.digsfish.com

spionid mudworms in Australia remains to be determined using both morphological and molecular techniques

(Walker 2011).

Spionid mudworms can infect virtually any molluscs that occur near muddy substrates, which means they

can infect several species of molluscs used as bait in Australia, particularly abalone and rock oysters, but not

species that are generally found completely buried in the sediments (e.g. venerid clams (or cockles, Anadara

spp., Katelysia spp.) and pipis (Plebidonax spp.)). Molluscs are widely used as bait throughout Australia,

with “other shellfish” (not including abalone) being used by an average of 33-38% of recreational anglers in

all states except the NT (Kewagama Research 2002, 2007), with highest use being in SA (84-88% of anglers)

and Victoria (55-65% of anglers) (Kewagama Research 2002, 2007). The majority of the molluscs

commercially available are likely to be bivalves such as live or frozen pipis (Plebidonax spp.) and venerid

clams (Katelysia spp., Anadara spp.), but these are unlikely to be infected with mudworms because they live

under sediments. The main route of translocation would appear to be movements of live oysters or mussels

collected by recreational fishers at or near the fishing site. Large quantities of oysters are widely distributed

as seafood and are sold live, chilled or frozen. However, these are unlikely to have heavy mudworm

infections as affected oysters are not marketable and are usually removed from sale (Nell 2001). Hence only

lightly infected oysters would be translocated via the seafood route, and some of these could also be diverted

to use as bait or berley around the country.

Larval mudworms do not survive drying in air for as little as 2 to 4 hours (Lleonart 2003a), while adult

mudworms also cannot survive drying for more than 4 or 5 days out of the water (Nell 2001). Hence

removal of the oysters from the water during processing, grading and delivery to market greatly reduces the

viability of any mudworms that are present. Taking into account the relatively small quantities of susceptible

molluscs used as bait or berley, as well as the fact that spionid mudworm infections are highly prevalent in

wild populations of molluscs in several areas of the country, the likelihood estimations for the occurrence of

spionid mudworms in these commodities are listed below.

Release assessment for infection of molluscs with spionid mudworms

Commodity type Live

molluscs

Whole fresh

dead molluscs

Frozen whole

molluscs

Mollusc shells

Likelihood of

release

Moderate Low Negligible Moderate


Molluscs in all parts of Australia are already at risk of exposure to spionid mudworms, however,

translocation of infected molluscs from areas where different species of mudworms are enzootic via their use

as bait or berley could transport these agents to new regions. Infection and establishment of spionid

mudworms in new hosts would occur only if viable mudworms in shells discarded into the water at the

_________________________________________________________________________________________________________________________________________________________________________________


216

www.digsfish.com

fishing site survived and reproduced via sexual or asexual reproduction. Asexual reproduction occurs via

either architomy (regeneration of fragments of the body into new individuals) or paratomy (division of the

parent body into two halves with reconstitution of the missing halves by regeneration) (Walker 2011).

Sexual reproduction in spionids does not appear to involve copulation even though populations have male

and female individuals. Sperm is released from the male within spermatophores that float freely from the

tube and these are picked up by the palps of the female, which store the sperm in seminal receptacles to be

used as required for egg fertilization (Walker 2011). Fertilized eggs are deposited in capsules within the tube

of the female, which is able to reproduce at an age of 3 months, producing up to 4 broods of eggs a year with

up to 2200 eggs per brood. The eggs hatch into planktonic larvae and larval settlement is stimulated by high

counts of some types of microbial flora associated with sediments (Sebesvari et al. 2006). Settlement of

larvae is also highly seasonal (Handley 2000, Lleonart et al. 2003a, 2003b)

Under natural circumstances, susceptible molluscs do not need to be in close proximity to mudworm infected

molluscs for horizontal transmission to occur, as under ideal environmental conditions favourable to the

mudworm, a single larvae is sufficient to initiate colonization of the host mollusc (Whitelegge 1890). As

these worms are ectocommensals, the immune system of the host is likely to play little or no role in

modulation of the infection process, hence the main factor controlling infection intensity (and therefore

whether disease occurs at all), will be environmental conditions. Proliferation of spionid mudworms will be

encouraged in Australian estuaries wherever increased sedimentation and organic enrichment occurs. Taking

these various factors into consideration, the risk of exposure and establishment of spionid mudworms via use

of bait and berley remains non-negligible, and the likelihood of exposure and establishment of spionid

mudworms in new mollusc populations via translocation is considered to be moderate.


Spionid mudworms are already present in most or all regions of Australia, however the distribution of some

species may be restricted to certain areas at this time. There is evidence that spionid mudworms have been

associated with disease outbreaks in cultured abalone reared at high densities in enclosed systems, however

they are not considered primary pathogens in wild and cultured oysters, and rather are opportunistic

commensals of oysters reared under suboptimal conditions. Spionid mudworms are not listed by the OIE or

NACA as a reportable disease, but some species of spionids remain listed as a reportable disease in SA

(Table 3b). However, control of these agents in cultured molluscs is relatively straightforward, and they do

not cause disease or mortality unless their numbers increase due to environmental conditions of organic

enrichment and sedimentation that favour the polychaete. Their presence does, however, cause marketability

issues for cultured molluscs, and the potential for economic loss due to this must be considered. Considering

all of these factors, establishment of spionid mudworms in new areas would potentially have mild biological

consequences and could cause minor and short term economic problems for mollusc aquaculture industries,

together with largely insignificant environmental effects for ecosystems and wild mollusc fisheries. It is

therefore estimated that the consequences of introduction of spionid mudworms into different parts of the

Australian environment via use of infected bait would likely be Very low.

_________________________________________________________________________________________________________________________________________________________________________________


217

www.digsfish.com


The unrestricted risk associated with spionid mudworm infections is determined by combining the likelihood


risk estimate for spionid mudworm infections does not exceed the ALOP for any of the commodity types,

suggesting that additional risk management for these agents is not required at this time.

Risk estimate for infection of molluscs with spionid mudworms

Commodity type Live molluscs Whole fresh dead

molluscs

Frozen whole

molluscs

Mollusc shells

Combined likelihood

of release and

exposure

Low Low Negligible Low

Consequences of

establishment

Very low Very low Very low Very Low

Risk estimation Negligible risk Negligible risk Negligible risk Negligible risk

_________________________________________________________________________________________________________________________________________________________________________________


218

www.digsfish.com

6.0 Risk Mitigation

6.1. Risk Evaluation

In Section 5, the 44 diseases of concern were placed into 35 different categories and a detailed risk

assessment was undertaken on each one. The outcomes of the risk assessment indicated 21 diseases for

which the unmitigated risk exceeded the ALOP (Table 11). Two diseases were classified as high risk of

spread via translocation of bait and berley, namely EHNV of finfish and AVG of abalone. Three diseases

were classified as moderate risk, including EUS of finfish, and infection of molluscs with Bonamia and

Perkinsus. Sixteen diseases were classified as low risk, including VER of finfish, goldfish ulcer disease,

microsporidian infections of finfish and crustaceans, infections of live finfish, molluscs and crustaceans with

introduced species of digeneans, nematodes and cestodes, infections of live finfish with introduced copepods

and Caligus epidemicus, infection of finfish and annelids with myxosporeans, viral infections of freshwater

crayfish, GAV, SMV and WTD of prawns, infections of crustaceans with Hematodinium spp. and Sacculina

spp., infections of molluscs with Haplosporidians, and infections of molluscs and anneids with Marteilia

sydneyi (Tables 11, 12)

Live finfish, crustaceans, molluscs and annelids were by far the commodities with the highest risk of

introduction and establishment of disease agents via translocation. Fresh dead (chilled) commodities were

also generally high risk commodities, however, the risks posed by EUS and many protozoan and metazoan

disease agents was significantly lowered or made negligible by the process of freezing or freeze drying

(Table 12). Freezing did not reduce the risk of establishment of some viruses however, particularly for

EHNV and AVG, and Perkinsus olseni is highly resistant to freezing and this parasite was found to represent

a particularly high risk of establishment in all types of translocated mollusc commodities (excluding mollusc

shells). Some options for reducing these risks to within the ALOP are listed in section 6.2.

Of course, knowledge regarding new and emerging diseases always evolves rapidly. It is also important to

realise that the status of some of the existing disease agents with respect to the ALOP may change at any

time in the future, especially if industries based on movement of live finfish, crustaceans, molluscs or

amphibians for bait become established, as they have in some other parts of the world (e.g. in the USA, see

Goodwin et al. 2004, Pernet et al. 2008, Picco and Collins 2008, Miller 2009). Because of this, the hazard

list, and this RA, will require regular updating to consider new information on diseases of bait and berley

commodities as it becomes available, as well as whenever there are significant changes to either Australias

aquatic animal disease status or how bait is used in Australia. There remain several large data gaps in

relation to disease agents that infect aquatic animals in Australia, particularly for species such as pipis,

cockles, bait crabs, callianassids, cephalopod molluscs, annelids, echinoderms, and ascideans, all of which

are commonly used as bait or berley (Appendix 1). Surveillance should be undertaken to begin to fill these

data gaps. Importantly, it must be realised that there remains a significant risk of transfer of as yet unknown

disease agents, even in the absence of their identification (Gaughan 2002), and that active surveillance may

be the only way to minimise the risk of transfer of these unknown disease agents.

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

21

9

w

ww

.dig

sfis

h.co

m

Tab

le 1

1. T

he d

isea

se a

gen

ts id

entif

ied

as r

equi

ring add

itio

nal r

isk

man

agem

ent,

incl

udin

g es

timat

ed c

onse

quenc

es o

f est

ablis

hmen

t and

the

hig

hest

unm

itiga

ted

risk

estim

atio

n in

var

ious

co

mm

oditi

es.

Add

itio

nal r

isk

man

agem

ent i

s re

quire

d fo

r th

ose

dise

ases

hig

hlig

hted

in

red

.

Dis

ease

C

om

bine

d lik

lihoo

d of

rel

ease

+ e

xpos

ure

Con

sequ

ence

s o

f es

tabl

ishm

ent

Hig

hest

unm

itiga

ted

risk

estim

atio

n R

isk

miti

gatio

n re

quire

d ?

Ris

ky c

om

mo

ditie

s

Infe

ctio

n of

fin

fish

and

mol

lusc

s w

ith A

qua

tic B

irna

viru

s

Low

- V

ery

Low

Lo

w

Ver

y Lo

w r

isk

No

Non

e

Infe

ctio

n of

fin

fish

with

Ep

izoo

tic H

aem

ato

poie

tic N

ecr

osis

V

irus

(EH

NV

) M

oder

ate

- L

ow

Hig

h H

igh

risk

Yes

A

ll fin

fish

com

mod

ities

Infe

ctio

n of

fin

fish

with

V

iral

Enc

epha

lopa

thy

and

R

etin

opa

thy

(Nod

avi

rus)

M

oder

ate

- L

ow

Low

Lo

w R

isk

Yes

Li

ve f

infis

h w

hole

fre

sh f

infis

h

who

le f

roze

n fin

fish,

fr

ozen

fis

h he

ads

E

mer

genc

e of

a n

ew p

revi

ousl

y u

nkno

wn

viru

s of

fin

fish

Lo

w

Unk

now

n N

ot p

ossi

ble

N

ot p

ossi

ble

N

ot p

ossi

ble

In

fect

ion

of f

infis

h w

ith A

ero

mo

na

s sa

lmo

nic

ida (G

oldf

ish

ulce

r di

sea

se)

Mod

era

te -

Low

Lo

w

Low

Ris

k

Yes

liv

e fin

fish

w

hole

fre

sh f

infis

h In

fect

ion

of s

alm

onid

s w

ith La

cto

cocc

us

ga

rvie

ae

Low

- V

ery

Low

Lo

w

Ver

y Lo

w R

isk

No

Non

e In

fect

ion

of

salm

onid

s w

ith P

isci

rick

etts

ia-lik

e b

act

eria

(P

LBs)

Lo

w -

Ext

rem

ely

Low

Lo

w

Ver

y Lo

w R

isk

No

Non

e

Infe

ctio

n of

fin

fish

with

Yer

sin

ia r

uck

eri (Y

ersi

nosi

s)

Ver

y Lo

w -

Ext

rem

ely

Low

V

ery

Low

N

eglig

ible

Ris

k N

o N

one

Infe

ctio

n of

fin

fish

with

Ap

ha

no

myc

es i

nva

da

ns

(E

piz

ootic

U

lcer

ativ

e S

yndr

ome)

Mod

era

te –

Ext

rem

ely

Low

M

oder

ate

M

oder

ate

Ris

k Y

es

live

finfis

h

Who

le f

resh

fin

fish

Infe

ctio

n of

fin

fish

with

Mic

rosp

orid

ians

Mod

era

te -

Low

Lo

w

Low

Ris

k Y

es

live

finfis

h

Who

le f

resh

fin

fish

Sys

tem

ic a

moe

bic

infe

ctio

ns o

f go

ldfis

h

Neg

ligib

le

Neg

ligib

le

Neg

ligib

le R

isk

No

Non

e In

fect

ion

of f

infis

h w

ith s

cutic

ocili

ate

s (in

clu

ding

U

ron

ema

spp.

) N

eglig

ible

N

eglig

ible

N

eglig

ible

Ris

k N

o N

one

Infe

ctio

n of

fin

fish

with

mon

ogen

eans

Low

- n

eglig

ible

Lo

w

Ver

y Lo

w R

isk

No

Non

e In

fect

ion

of f

infis

h a

nd m

ollu

scs

with

dig

enea

ns Lo

w -

neg

ligib

le

Mod

era

te

Low

Ris

k Y

es

live

finfis

h

Infe

ctio

n of

fin

fish

and

cr

usta

cea

ns

with

ne

ma

tode

s a

nd

cest

odes

Low

- n

eglig

ible

M

oder

ate

Lo

w R

isk

Yes

liv

e fin

fish

Infe

ctio

n of

fin

fish

with

cop

epod

s Lo

w -

neg

ligib

le

Mod

era

te

Low

Ris

k Y

es

live

finfis

h

Infe

ctio

n of

fin

fish

and

ann

elid

s w

ith m

yxos

pore

ans

M

oder

ate

- L

ow

Low

Lo

w R

isk

Yes

liv

e fin

fish

W

hole

fre

sh f

infis

h V

iral i

nfec

tions

of

fres

hwa

ter

cra

yfis

hes

Lo

w -

Ver

y Lo

w

Mod

era

te

Low

ris

k Y

es

live

cra

yfis

h W

hole

fre

sh c

rayf

ish

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

22

0

w

ww

.dig

sfis

h.co

m

Dis

ease

C

om

bine

d lik

lihoo

d of

rel

ease

+ e

xpos

ure

Con

sequ

ence

s o

f es

tabl

ishm

ent

Hig

hest

unm

itiga

ted

risk

estim

atio

n R

isk

miti

gatio

n re

quire

d ?

Ris

ky c

om

mo

ditie

s

Infe

ctio

n of

pra

wns

with

Gill

Ass

ocia

ted

Viru

s (G

AV

)

Mod

era

te –

Ver

y Lo

w

Mod

era

te

Low

ris

k Y

es

Live

pra

wns

W

hole

fre

sh p

raw

ns

Infe

ctio

n of

cru

sta

cea

ns

with

Hep

ato

panc

rea

tic P

arv

oviru

s (H

PV

)

Low

- V

ery

Low

Lo

w

Ver

y Lo

w R

isk

No

Non

e

Infe

ctio

us

Hyp

oder

ma

l a

nd

Ha

ema

top

oiet

ic

Nec

rosi

s of

pr

aw

ns (

IHH

NV

) Lo

w -

Ver

y Lo

w

Low

V

ery

Low

Ris

k N

o N

one

Infe

ctio

n of

pra

wns

with

Mon

odon

Ba

culo

viru

s (M

BV

)

Low

- V

ery

Low

V

ery

Low

N

eglig

ible

Ris

k N

o N

one

Infe

ctio

n of

pra

wns

with

Mou

rily

an

viru

s (M

oV)

Low

- V

ery

Low

V

ery

Low

N

eglig

ible

Ris

k N

o N

one

Infe

ctio

n of

pr

aw

ns

and

cr

ayf

ish

with

S

paw

ner

Isol

ate

d M

orta

lity

Viru

s (S

MV

) Lo

w -

Ver

y Lo

w

Mod

era

te

Low

ris

k Y

es

Live

pra

wns

W

hole

fre

sh p

raw

ns

Whi

te T

ail

Dis

ease

of

fres

hwa

ter

gia

nt p

raw

ns

Low

- V

ery

Low

M

oder

ate

Lo

w r

isk

Yes

Li

ve F

W p

raw

ns

Who

le f

resh

FW

pra

wns

In

fect

ion

of c

rust

ace

ans

with

RLO

s

Low

- V

ery

Low

V

ery

Low

N

eglig

ible

Ris

k N

o N

one

Infe

ctio

n of

cru

sta

cea

ns w

ith

Hem

ato

din

ium

spp.

Lo

w -

Ver

y Lo

w

Mod

era

te

Low

ris

k Y

es

Live

cru

sta

cea

ns

Who

le f

resh

cru

sta

cea

ns

Infe

ctio

n of

cru

sta

cea

ns w

ith M

icro

spor

idia

ns M

oder

ate

- L

ow

Low

Lo

w R

isk

Yes

Li

ve c

rust

ace

ans

W

hole

fre

sh c

rust

ace

ans

In

fect

ion

of

cra

bs

with

Sa

ccu

lina

spp.

a

nd

othe

r rh

izoc

epha

lan

barn

acl

es Lo

w -

Neg

ligib

le

Mod

era

te

Low

ris

k Y

es

Live

cra

bs

Vira

l Ga

nglio

neur

itis

of A

balo

ne (

AV

G)

Mod

era

te-

Low

H

igh

Hig

h ris

k Y

es

All

aba

lone

com

mod

ities

(e

xclu

ding

she

lls)

Infe

ctio

n of

mol

lusc

s w

ith Bo

na

mia

spp.

, a

nd/o

r un

ide

ntifi

ed

mic

roce

lls

Mod

era

te -

Neg

ligib

le

Mod

era

te

Mod

era

te R

isk

Yes

Li

ve o

yste

rs

Who

le f

resh

oys

ters

In

fect

ion

of m

ollu

scs

with

Ha

plos

pori

dia

ns Lo

w -

Neg

ligib

le

Mod

era

te

Low

ris

k Y

es

Live

oys

ters

Infe

ctio

n of

oys

ters

and

ann

elid

s w

ith

Ma

rtei

lia s

ydn

eyi (Q

X

dise

ase

) Lo

w -

Ver

y Lo

w

Mod

era

te

Low

Ris

k

Yes

Li

ve o

yste

rs a

nd a

nnel

ids

Who

le f

resh

oys

ters

and

a

nnel

ids

Infe

ctio

n of

mol

lusc

s w

ith Per

kin

sus

ols

eni

Hig

h -

Mod

era

te

Mod

era

te

Mod

era

te R

isk

Yes

A

ll m

ollu

sc c

omm

oditi

es

(exc

ludi

ng s

hells

) In

fect

ion

of m

ollu

scs

with

spi

onid

mu

dwor

ms

Low

- N

eglig

ible

V

ery

Low

N

eglig

ible

Ris

k N

o N

one

_________________________________________________________________________________________________________________________________________________________________________________


221

www.digsfish.com

Table 12. Commodities potentially harbouring disease agents that require additional risk

management. �� = high risk, �� = moderate risk, ��= low risk, X = within ALOP.


FINFISH EHNV EUS VER GUD Microsporidians Myxosporeans Digeneans cestodes

nematodes copepods+

Live finfish ��

Whole fresh dead finfish

�� X

Frozen whole finfish

�� X �� X X X X

Frozen fish fillets �� X X X X X X

Frozen fish heads �� X �� X X X X

Frozen fish guts/offal

�� X X X X X X

CRUSTACEANS Viruses of FW

crayfish

GAV SMV WTD Microsporidians Hematodinium spp.

Sacculina spp.

Live prawns X �� * �� X X

Live crayfish/ lobsters

�� X �� X �� ? X

Live crabs X X X X ��

Whole fresh dead prawns

X �� * �� X X

Whole fresh dead crayfish / lobsters

�� X X X �� X X

Whole fresh dead crabs

X X X X �� X

Frozen whole prawns

X X X X X X X

Frozen whole crayfish / lobsters

X X X X X X X

Frozen whole crabs

X X X X X X X

Frozen prawn tails X X X X X X X

Frozen crayfish / lobster tails

X X X X X X X

Frozen prawn heads

X X X X X X X

Frozen crayfish / lobster heads

X X X X X X X

_________________________________________________________________________________________________________________________________________________________________________________


222

www.digsfish.com


MOLLUSCS AVG Perkinsus olseni

Bonamia spp.

Marteilia sydneyi

Haplosporidians Digeneans

Live molluscs X �� X X ��

Live oysters X ��

Live abalone �� X X ��

Whole fresh dead molluscs

X �� X X X X

Whole fresh dead oysters

X �� X X

Whole fresh dead abalone

�� X X X X

Frozen whole molluscs

X �� X X X X

Frozen whole oysters

X �� X X X X

Frozen whole abalone

�� X X X X

Frozen mollusc meat

X �� X X X X

Frozen abalone meat and viscera

�� X X X X

ANNELIDS Marteilia sydneyi

Myxo-sporeans

Live annelids ��

Fresh dead annelids

��

Frozen annelids X X

Freeze dried annelids

X X

+ = Introduced species, as well as Caligus epidemicus, * = freshwater prawns (Macrobrachium spp.) only,

? = unknown, as marine crayfish/lobsters in Australia have not been actively surveyed for Hematodinium

spp. at this time.

_________________________________________________________________________________________________________________________________________________________________________________


223

www.digsfish.com

6.2 Options for Risk Mitigation

There are many options available for mitigating the risk of disease translocation posed by those commodities

that exceed the ALOP. Some of these include:

1. Controls on use of particular high risk commodities as bait (e.g. extend nationally and strengthen

existing bans on use of abalone products as bait to reduce the risk of spread of AVG).

2. Controls on use of aquacultured commodities as bait, and/or compulsory disease surveillance/ testing

of aquacultured products used as bait (such as live polychaetes).

3. Controls on translocation of live bait (which would reduce the risk of spread of all disease agents of

concern for finfish, crustaceans, molluscs and annelids)

4. Compulsory freezing/freeze drying of all bait commodities, which would reduce the risk of spread of

all diseases to within the ALOP, except for EHNV and VER in finfish, and AVG and Perkinsus

olseni in molluscs (Table 12). Methods of inactivation of these other pathogens could also be

investigated and implemented, if necessary and practical.

5. Increased active surveillance of bait commodities, particularly those where data gaps were identified

(e.g. pipis, cockles, bait crabs, callianassids, cephalopod molluscs, annelids, echinoderms, and

ascideans). Disease surveillance should also be undertaken in the early stages of development of

fisheries for new species likely to be used as bait, and/or whenever significant quantities of bait are

being translocated to a new geographical region.

6. Educating fishers of the risks involved with transfer of disease via translocation of bait. Examples of

education and extension could include, posters (Appendix 2) erected in tackle shops, pet shops

fishing co-ops, or boat ramps, educational pamphlets distributed with fishing or boating licenses or

at tackle shows, and even signage at retail points of sale for seafood, encouraging consumers not to

use at risk types of seafood as bait or berley.

These and other options for risk mitigation should be examined in more detail and prioritised, preferably by a

national working group including representatives from all states and territories (with stakeholder

involvement wherever necessary), in order to develop the most appropriate and effective options for risk

mitigation during the risk management and risk communication phases of this risk analysis process.

_________________________________________________________________________________________________________________________________________________________________________________


224

www.digsfish.com

7.0 References

Abollo E, Ramilo A, Casas SM, Comesaña P, Cao A, Carballal MJ, Villalba A (2008). First detection of the

protozoan parasite Bonamia exitiosa (Haplosporidia) infecting flat oyster Ostrea edulis grown in European

waters. Aquaculture 274: 201–207.

Adlard RD, Ernst I (1995). Extended range of the oyster pathogen Marteilia sydneyi. Bulletin of the

European Association of Fish Pathologists 15: 119-121.

Adlard RD, Lester RJG (1995). Development of a diagnostic test for Mikrocytos roughleyi, the aetiological

agent of Australian winter mortality of the commercial rock oyster, Saccostrea commercialis (Iredale and

Roughley). Journal of Fish Diseases 18: 609-614.

Adlard RD, Nolan MJ (2008). Aquatic animal health subprogram: Identification of host interactions in the

life cycle of QX disease. Report prepared by QLD Museum for FRDC project 2006/062. 60 pgs.

AFFA (2002). Import Risk Analysis (IRA) of non-viable bivalve molluscs. Technical Issues Paper.

September 2002. Biosecurity Australia – Agriculture, Fisheries and Forestry Australia. 67 pgs.

Alderman DJ (1996) Geographical spread of bacterial and fungal diseases of crustaceans. Revue Scientifique

et Technique Office International des Epizooties 15(2): 603-632.

Alvarez-Pellitero P, Sitja-Bobadilla A (1993). Pathology of Myxosporea in marine fish culture. Diseases of

Aquatic Organisms 17: 229-238.

Alvarez-Pellitero P, Palenzuela O, Padros F, Sitjà-Bobadilla A, Riaza A, Silva R, Arán J (2004).

Histophagous scuticociliates (Cilophora) parasitizing turbot Scophthalmus maximus: morphology, in vitro

culture and virulence. Folia Parasitolology 51: 177–187.

Amigo JM, Gracia MP, Rius M, Salvado H, Maillo PA, Vivares CP (1996). Longevity and effects of

temperature on the viability and polar-tube extrusion of spores of Glugea stephani, a microsporidian parasite

of commercial flatfish. Parasitology Research 82: 211-214.

Anderson C, Knowles G, de Lisle G (1994). A survey for Yersinia ruckeri and Aeromonas salmonicida in

farmed and wild fish. Surveillance 21(3): 39-40.

Anderson IG, Prior HC (1992). Baculovirus infections in the mudcrab, Scylla serrata, and a freshwater

crayfish, Cherax quadricarinatus, from Australia. Journal of Invertebrate Pathology 60: 265-273.

Anderson IG, Oakey HJ (2008). The production of nodavirus-free fish fry and the nodaviruses natural

distribution : FRDC final report FRDC project ; no. 2002/043. 63 pgs.

_________________________________________________________________________________________________________________________________________________________________________________


225

www.digsfish.com

Anderson IG, Shariff M, Nash G, Nash M (1987). Mortalities of juvenile shrimp, Penaeus monodon,

associated with Penaeus monodon baculovirus, cytoplasmic reo-like virus, and rickettsial and bacterial

infections, from Malaysian brackishwater ponds. Asian Fisheries Science 1: 47-64.

Anderson IG, Shariff M, Nash G (1989). A hepatopancreatic microsporidian in pond-reared tiger shrimp,

Penaeus monodon, from Malaysia. Journal of Invertebrate Pathology 53: 278-280.

Anderson IG, Law AT, Shariff M, Nash F (1990). A parvo-like virus in the giant freshwater prawn,

Macrobrachium rosenbergii. Journal of Invertebrate Pathology 55: 447-449.

Anderson TJ, Hine PM, Lester RJG (1995) A Steinhausia-like infection in the ovocytes of Sydney rock

oysters Saccostrea commercialis. Diseases of Aquatic Organisma 22: 143-146.

Anderson SA, Hulston DA, McVeagh M, Webb VL, Smith PJ (2009). In vitro culture and cryopreservation

of Uronema marinum isolated from farmed New Zealand groper (Polyprion oxygeneios). Journal of

Microbiological Methods 79: 62–66.

Andrews JD (1968). Oyster mortality studies in Virginica. VII Review of epizootiology and origin of

Minchinia nelsoni. Proceedings of the National Shellfisheries Association 58: 23-36.

Andrews JD (1996). History of Perkinsus marinus , a pathogen of oysters in Chesapeake Bay 1950-1984.

Journal of Shellfish Research 15: 13-16.

Aohagi Y, Shibahara T, Machida N, Yamaga Y, Kagota K (1992). Clinostomum complanatum (Trematoda:

Clinostomatidae) in five new fish hosts in Japan. Journal of Wildlife Diseases 28: 467-469.

AQIS (1999a). Import risk analysis on live ornamental finfish. Australian Quarantine and Inspection

Service July 1999. 172 pgs.

AQIS (1999b). Import risk analysis on non-viable salmonids and non-salmonid marine finfish. Australian

Quarantine and Inspection Service July 1999. 409 pgs.

Archer DL (2004). Freezing: an underutilized food safety technology? International Journal of Food

Microbiology 90: 127-138.

Arcier JM, Herman F, Lightner DV, Redman RM, Mari J, Bonami JR (1999). A viral disease associated

with mortalities in hatchery reared post larvae of the giant freshwater prawn Macrobrachium rosenbergii.

Diseases of Aquatic Organisms 38: 177-181.

Ariel E, Owens L (1997). Epizootic mortalities in tilapia Oreochromis mossambicus. Diseases of Aquatic

Organisms 29: 1-6.

_________________________________________________________________________________________________________________________________________________________________________________


226

www.digsfish.com

Arsan EL, Bartholomew JL (2008). Potential for dissemination of the nonnative salmonid parasite

Myxobolus cerebralis in Alaska. Journal of Aquatic Animal Health 20: 136-149.

Arthington AH, Bluhdorn DR (2005). Improved management of exotic aquatic fauna: R&D for Australian

rivers. The Land and Water Resources Research and Development Corporation Occasional Paper 04/95.

Arthur JR, Subasinghe RP (2002). Potential adverse socio-economic and biological impacts of aquatic

animal pathogens due to hatchery-based enhancement of inland open-water systems, and possibilities for

their minimisation. pgs. 113-126. In: J.R. Arthur, M.J. Phillips, R.P. Subasinghe, M.B. Reantaso and I.H.

MacRae. (eds.) Primary Aquatic Animal Health Care in Rural, Small-scale, Aquaculture Development. FAO

Fish. Tech. Pap. No. 406.

Arzul I, Renault T, Lipart C (2001). Experimental herpes-like viral infections in marine bivalves:

demonstration of interspecies transmission. Diseases of Aquatic Organisms 46: 1-6.

Ashburner LD (1976). Australian fish face threat from foreign diseases. Australian Fisheries 36: 18-21.

Ashburner LD (1978). Management and diseases of hatchery fish. Proceedings No.36 of Course for

Veterinarians, University of Sydney, pgs. 387-449.

Audemard C, Barnaud A, Collins CM, Le Roux F, Sauriau PG, Coustau C, Blachier P, Berthe FCJ (2001).

Claire ponds as an experimental model for Marteilia refringens life-cycle studies: new perspectives. Journal

of Experimental Marine Biology and Ecology 257: 87-108.

Audemard C, Le Roux F, Barnaud A, Collins C, Sautour B, Sauriau PG, De Montaudouin X, Coustau C,

Combes C, Berthe FCJ (2002). Needle in a haystack: involvement of the copepod Paracartia grani in the life

cycle of the oyster pathogen Marteilia refringens. Parasitology 124: 315–323.

Audemard C, Sajus M, Barnaud A, Sautour B, Sauriau P, Berthe F (2004). Infection dynamics of Marteilia

refringens in flat oyster Ostrea edulis and copepod Paracartia grani in a claire pond of Marennes-Oleron

Bay. Diseases of Aquatic Organisms 61: 103–111.

Austin B, Austin DA (2007). Bacterial fish pathogens: disease in farmed and wild fish. 4th edition.

Springer. 552 pgs.

Bain MB, Cornwell ER, Hope KM, Eckerlin GE, Casey RN, Groocock GH, Getchell RG, Bowser PR,

Winton JR, Batts WN, Cangelosi A, Casey JW (2010). Distribution of an invasive aquatic pathogen (Viral

Hemorrhagic Septicemia Virus) in the Great Lakes and its relationship to shipping. PLoS ONE 5(4): e10156.

doi:10.1371/journal.pone.0010156

Bakke TA, Harris PD, Jansen PA, Hansen LP (1992). Host specificity and dispersal strategy in gyrodactylid

monogeneans, with particular reference to Gyrodactylus salaris (Platyhelminthes: Monogenea). Diseases of


_________________________________________________________________________________________________________________________________________________________________________________


227

www.digsfish.com

Bangyeekhun E, Ryynänen HJ, Henttonen P, Huner JV, Cerenius L, Söderhäll K (2001). Sequence analysis

of the ribosomal internal transcribed spacer DNA of the crayfish parasite Psorospermium haeckeli. Diseases

of Aquatic Organisms 46: 217-222.

Barber RD, Ford SE (1992). Occurrence and significance of ingested haplosporidian spores in the eastern

oyster, Crassostrea virginica (Gmelin, 1791). Journal of Shellfish Research 11: 371-375.

Barlow CG, Hogan AE, Rodgers LJ (1987). Implication of translocated fishes in the apparent extinction in

the wild of the Lake Eacham rainbowfish, Melanotaenia eachamensis. Australian Journal of Marine and

Freshwater Research 38: 897-902.

Barta JR, Desser SS (1984). Blood parasites of amphibians from Algonquin Park, Ontario. Journal of

Wildlife Diseases 20: 180-189.

Bartholomew JL, Reno PW (2002). The history and dissemination of Whirling Disease. American Fisheries

Society Symposium 29: 3-24.

Barton DP (1994). A checklist of helminth parasites of Australian amphibia. Records of the South

Australian Museum 27: 13-30.

Bauer ON (1991). Spread of parasites and diseases of aquatic organisms by acclimatization: a short review.

Journal of Fish Biology 39: 679-686.

Bearham D, Spiers Z, Raidal S, Jones JB, Nicholls PK (2007). Molecular characterisation of a

haplosporidian parasite infecting rock oysters Saccostrea cuccullata in northern Western Australia. Journal

of Invertebrate Pathology 95:33–40.

Bearham D, Spiers Z, Raidal S, Jones JB, Nicholls PK (2008a). Detection of Minchinia sp. in rock oysters

Saccostrea cuccullata (Born 1778) using DNA probes. Journal of Invertebrate Pathology 97: 50-60

Bearham D, Spiers Z, Raidal S, Jones JB, Burreson EM, Nicholls PK (2008b). Spore ornamentation of

Haplosporidium hinei n. sp. (Haplosporidia) in pearl oysters Pinctada maxima (Jameson, 1901).

Parasitology 135:521–527.

Bearham D, Spiers Z, Raidal S, Jones JB, Nicholls PK (2008c). Spore ornamentation of Minchinia occulta n.

sp. (Haplosporidia) in rock oysters Saccostrea cuccullata (Born, 1778). Parasitology 135:1271–1280.

Bearham D, Spiers Z, Raidal S, Jones JB, Nicholls PK (2009). Detection of Minchinia occulta in samples of

pearl oysters Pinctada maxima infected by Haplosporidium hinei. Australian Veterinary Journal 87: 430-

437.

Bell TA, Lightner DV (1984). IHHN virus: infectivity and pathogenicity studies in Penaeus stylirostris and

Penaeus vannamei. Aquaculture 38: 185–194.

_________________________________________________________________________________________________________________________________________________________________________________


228

www.digsfish.com

Bennett I (1987). Australian Seashores. Angus and Robertson. 411 pgs.

Bergmann SM, Sadowski J, Kielpinski M, M Bartlomiejczyk, Fichtner D, Riebe R, Lenk M, Kempter J

(2010). Susceptibility of koi x crucian carp and koi x goldfish hybrids to koi herpesvirus (KHV) and the

development of KHV disease (KHVD). Journal of Fish Diseases 33: 267-272.

Berthe FCJ, Hine PM (2004). Corrigendum: Bonamia exitiosa Hine et al. 2001 is proposed as the valid

name of Bonamia sp. infecting flat oysters, Ostrea chilensis, in New Zealand. Diseases of Aquatic

Organisms 57: 181.

Berthe FCJ, Le Roux F, Adlard RD, Figueras A (2004). Marteiliosis in molluscs: A review. Aquatic Living

Resources 17: 433-448.

Beumer JP, Ashburner LD, Burbury ME, Jette E, Latham DJ (1982). A checklist of the parasites of fishes

from Australia and its adjacent Antarctic Territories. Technical Communication 48 of the Commonwealth

Institute of Parasitology.

Bigarr L, Cabon J, Baud M, Heimann M, Body A, Lieffrig F, Castric J (2009). Outbreak of betanodavirus

infection in tilapia, Oreochromis niloticus (L.), in fresh water. Journal of Fish Diseases 32: 667-673.

Biosecurity Australia (2006). Revised Draft Generic Import Risk Analysis Report for Prawns and Prawn

Products, Part B. Biosecurity Australia, Canberra, Australia. 284 pgs.

Biosecurity Australia (2008). Biosecurity Australia advice 2008/30. Importation of prawns and prawn

products – amended interim quarantine measures. 12 September 2008. .

http://www.daff.gov.au/__data/assets/word_doc/0006/810825/2008-30.doc

Biosecurity Australia (2009) Generic Import Risk Analysis Report for Prawns and Prawn Products. Final

Report. Biosecurity Australia, Canberra, Australia. 7 October 2009, 292 pgs. .

http://www.daff.gov.au/__data/assets/pdf_file/0012/1302411/Final_prawn_RA_report_7_Oct_

09.pdf

Birkbeck TH, Rennie S, Hunter D, Laidler LA, Wadsworth S (2004). Infectivity of a Scottish isolate of

Piscirickettsia salmonis for Atlantic salmon Salmo salar and immune response of salmon to this agent.


Bishop RK, Cannon LRG (1979). Morbid behaviour of the commercial sand crab, Portunus pelagicus (L.),

parasitized by Sacculina granifera Boschma, 1973 (Cirripedia: Rhizocephala). Journal of Fish Diseases 2:

131–144.

Bishop ML, Carnegie RB, Stokes NA, Peterson CH, Burreson EM (2006) Complications of a non-native

oyster introduction: facilitation of a local parasite. Marine Ecology Progress Series 325:145–152

http://www.daff.gov.au/__data/assets/word_doc/0006/810825/2008-30.doc

http://www.daff.gov.au/__data/assets/pdf_file/0012/1302411/Final_prawn_IRA_report_7_Oct_09.pdf

_________________________________________________________________________________________________________________________________________________________________________________


229

www.digsfish.com

Bisset KA (1948). Seasonal changes in the normal bacterial flora of freshwater fish. Journal of Hygiene 46:

94-97.

Blakemore RJ (1999). Diversity of exotic earthworms in Australia – a status report. In: The Other 99%. The

Conservation and Biodiversity of Invertebrates ed. Ponder W, Lunney D (Eds). Transactions of the Royal

Zoological Society of New South Wales. Pgs 182-187

Blazek RD, Bagge A, Valtone ET (2008). Monogenean assemblages and the apparent transmission

capability of monogeneans between related fish species: an experimental study. Parasitology Research 102:

1359–1366.

Blazer VS, Lilley J, Schill WB, Kiryu Y, Densmore CL, Panyawachira V, Chinabut S (2002). Aphanomyces

invadans in Atlantic menhaden along the east coast of the United States. Journal of Aquatic Animal Health

14: 1–10.

Bonami JR, Trumper B, Mari J, Brehelin M, Lightner DV (1990). Purification and characterization of the

infectious hypodermal and haematopoietic necrosis virus of penaeid shrimps. Journal of General Virology

71: 2657-2664.

Bonami JR, Mari J, Poulos BT, Lightner DV (1995). Characterization of hepatopancreatic parvo-like virus, a

second unusual parvovirus pathogenic for penaeid shrimp. Journal of General Virology 76: 813–817.

Bonami JR, Widada FS (2003). Characterisation of 2 virues (MrNV and XSV), pathogenic agents of the

white tail disease (WTD) of the giant freshwater prawn Macrobrachium rosenbergii. In Freshwater Prawns

2003, International Symposium, Cochin , 20-23 August 2003.

Bonami JR, Shi Z, Qian D, Sri Widada J (2005). White tail disease of the giant freshwater prawn,

Macrobrachium rosenbergii: separation of the associated virions and characterization of MrNV as a new

type of nodavirus. Journal of Fish Diseases 28: 23-31.

Bond NR (2004). Observations on the effects of the introduced parasite Lernaea cypriniacea on a lowland

population of a small native Australian fish, mountain galaxias Galaxias olidus. Victorian Naturalist 121:

194-198.

Boschma H (1955). The described species of the family Sacculinidae. Zoologische Verhandelingen 27: 1-76.

Bott N, Healey JM, Cribb TH (2005). Patterns of digenean parasitism of bivalves from the Great Barrier

Reef and associated waters. Marine and Freshwater Research 56: 387-394.

Bovo G, Hill B, Husby A, Hastein T, Michel C, Olesen NJ, Storset A and Midtlyng P (2005). Work package

3 report: Pathogen survival outside the host, and susceptibility to disinfection. VESO, P.O. Box 8109 Dep.,

N-0032 Oslo, Norway. Available at: www.crl-fish.eu/upload/sites/eurl-fish/links/fisheggtrade%20wp_3.pdf

http://www.crl-fish.eu/upload/sites/eurl-fish/links/fisheggtrade wp_3.pdf

_________________________________________________________________________________________________________________________________________________________________________________


230

www.digsfish.com

Bowater RO, Wingfield M, Fisk A, Condon K, Reid A, Prior H, Kulpa E (2002). A parvo-like virus in

cultured redclaw crayfish Cherax quadricarinatus from Queensland, Australia. Diseases of Aquatic

Organisms 50: 79-86.

Bower SM, McGladdery SE, Price IM (1994). Synopsis of infectious diseases and parasites of commercially

exploited shellfish. Annual Review of Fish Diseases 4: 1-199.

Bower SM, Meyer GR, Boutillier JA (1996). Stained prawn disease (SPD) of Pandalus platyceros in British

Columbia, Canada, caused by a rickettsial infection. Diseases of Aquatic Organisms 24: 41-54.

Bower SM, Hervio D, Meyer GR (1997). Infectivity of Mikrocytos mackini, the causative agent of Denman

Island disease in Pacific oysters Crassostrea gigas, to various species of oysters. Diseases of Aquatic


Bray RA (1990). Hemiuridae (Digenea) from marine fishes of the southern Indian Ocean: Dinurinae,

Elytrophallinae, Glomericirrinae and Plerurinae. Systematic Parasitology 17: 183-217.

Bray RA, Cribb TH (1998). Lepocreadiidae (Digenea) of Australian coastal fishes: new species of Opechona

Looss, 1907, Lepotrema Ozaki, 1932 and Bianium Stunkard, 1930 and comments on other species reported

for the first time or poorly known in Australian waters. Systematic Parasitology 41: 123-148.

Breed GM, Olson RE (1977). Biology of the microsporidian parasite Pleistophora crangoni n. sp. in three

species of crangonid sand shrimps. Journal of Invertebrate Pathology 30: 387-405.

Bridle AR, Crosbie PB, Cadoret K, Nowak BF (2010). Rapid detection and quantification of Neoparamoeba

perurans in the marine environment. Aquaculture 309: 56–61.

Brisbane Courier (1898). The Oyster Industry. Ravages of the Worm Disease. Certain areas seriously

affected. Brisbane Courier, Tuesday 1 February 1898, page 5. .

http://trove.nla.gov.au/ndp/del/article/3665726

Broadhurst MK, Millar RB, Young DJ, Wooden ME, Rowland S (2004). Atypical size selection of captive

school prawns, Metapenaeus macleayi, by three recreational fishing gears in south-eastern Australia. New

Zealand Journal of Marine and Freshwater Research 38: 755-766.

Brock JA, Lightner DV (1990). Diseases of crustacea. Diseases caused by microorganisms. In: Kinne O (ed)

Diseases of Marine Animals, Vol. III. Biologische Anstalt Helgoland, Hamburg, Germany, pgs. 245-349.

Brock JA, Nakagawa LK, Hayashi T, Teruya S, Van Campen H (1986). Hepatopancreatic rickettsial

infection of the penaeid shrimp, Penaeus marginatus (Randall), from Hawaii. Journal of Fish Diseases 9:

73-77.

http://trove.nla.gov.au/ndp/del/article/3665726

_________________________________________________________________________________________________________________________________________________________________________________


231

www.digsfish.com

Browdy CL, Holloway JD Jr, King CO, Stokes AD, Hopkins JS, Sandifer PA (1993). IHHN virus and

intensive culture of Penaeus vannamei: effects of stocking density and water exchange rates. Journal of

Crustacean Biology 13: 87-94.

Bruce NL (1987). Australian species of Nerocila Leach, 1818, and Creniola n. gen. (Isopoda: Cymothoidae),

crustacean parasites of marine fishes. Records of the Australian Museum 39: 355-412.

Bruce NL (1990). Hysterothylacium Ward and Magath, 1917, and Ichthyascaris Wu, 1949, Ascaridoid

nematodes from Australian demersal fishes. Memoirs of the Queensland Museum 28: 389-426.

Budy P, Thiede GP, de la Hoz EA, Vatland S (2003). Logan River Whirling Disease Study: Factors

Affecting Trout Population Dynamics, Abundance, and Distribution in the Logan River, Utah. 2004 Annual

Report to Utah Division of Wildlife Resources No. F-47–R. Salt Lake City, UT: Utah Division of Wildlife

Resources, 69 pp.

Burreson EM, Stokes NA, Friedman CS (2000). Increased virulence in an introduced pathogen:

Haplosporidian nelsoni (MSX) in the Eastern oyster Crassostrea virginica. Journal of Aquatic Animal

Health 12: 1-8.

Burreson EM, Stokes NA, Carnegie RB (2004). Bonamia sp. (Haplosporidia) found in nonnative oysters

Crassostrea ariakensis in Bogue Sound, North Carolina. Journal of Aquatic Animal Health 16: 1-9.

Burrows DW (2004). Translocated Fishes in Streams of the Wet Tropics Region, North Queensland:

Distribution and Potential Impact. Cooperative Research Centre for Tropical Rainforest Ecology and

Management, Cairns. 83 pgs. http://www.wettropics.gov.au/res/downloads/translocatedFishesInStreams.pdf

Bushek D, Ford SE, Chintala MM (2002). Comparison of in vitro cultured and wild-type Perkinsus marinus.

III. Fecal elimination and its role in transmission. Diseases of Aquatic Organisms 51: 217-225.

Butler MJ, Behringer DC, Shields JD (2008). Transmission of Panulirus argus virus 1 (PaV1) and its effect

on the survival of juvenile Caribbean spiny lobster. Diseases of Aquatic Organisms 79: 173-182.

Butt D, Raftos DA (2007). Immunosuppression in Sydney rock oysters (Saccostrea glomerata) and QX

disease in the Hawkesbury River, Sydney. Marine and Freshwater Research 58: 213–221.

Butt D, Shaddick K, Raftos D (2006). The effect of low salinity on phenoloxidase activity in the Sydney

rock oyster, Saccostrea glomerata. Aquaculture 251: 159-166.

Byrnes T (1986). Five species of monogenea from Australian bream, Acanthopagrus spp. Australian

Journal of Zoology 34: 65-86.

Byrnes T (1987). Caligids (Copepoda: Caligidae) found on the bream (Acanthopagrus spp.) of Australia.

Journal of Natural History 21: 363- 404.

http://www.wettropics.gov.au/res/downloads/translocatedFishesInStreams.pdf

_________________________________________________________________________________________________________________________________________________________________________________


232

www.digsfish.com

Callinan RB, Paclibare JO, Bondad-Reantaso MG, Chin JC, Gogolewski RP (1995). Aphanomyces species

associated with epizootic ulcerative syndrome (EUS) in the Philippines and red spot disease (RSD) in

Australia: preliminary comparative studies. Diseases of Aquatic Organisms 21: 233-238.

Canning EU, Lom J (1986). The Microsporidia of Vertebrates. Toronto, Academic Press, 1986.

Campbell RE, Lilley JH, Taukhid, Panyawachira V, Kanchanakhan S (2001) In vitro screening of novel

treatments for Aphanomyces invadans. Aquaculture Research 32: 223–233.

Canadian Ministry of Natural Resources (CMNR) (2009) Bait for personal use. Available at:

www.mnr.gov.on.ca/en/Business/LetsFish/2ColumnSubPage/STEL02_165361.html

Caputi N, Stevens R, Christianopoulos D, McCoy D (2001). Off season trial of artificial rock lobster bait.

Fisheries Research and Development Corporation Report FRDC Project No. 99/372. ISBN 0-7309-8435-4

Carnegie RB, Burreson EM, Hine PM, Stokes NA, Audemard C, Bishop MJ, Peterson CH (2006). Bonamia

perspora n. sp. (Haplosporidia), a parasite of the oyster Ostreola equestris, is the first Bonamia species

known to produce spores. Journal of Eukaryotic Microbiology 53: 232-245.

Carson J, Handlinger J (1988). Virulence of the aetiological agent of goldfish ulcer disease in Atlantic

salmon, Salmo salar L. Journal of Fish Diseases 11: 471-479.

Carson J, Gudkovs N (2001). Aeromonas salmonicida: a sleeping giant. Microbiology Australia July 2001:

28-30.

Carson J, Wilson T (2009). Yersinosis in fish. Australian and New Zealand Standard Diagnostic Procedure.

http://www.scahls.org.au/__data/assets/pdf_file/0010/1516519/Yersiniosis.pdf

Carson J, Gudkovs N, Austin B (1993). Characteristics of an Enterococcus-like bacterium from Australia

and South Africa, pathogenic for rainbow trout (Oncorhynchus mykiss Walbaum). Journal of Fish Diseases

16: 381-388.

Castric J, Thiery R, Jeffroy J, deKinkelin P, Raymond JC (2001). Sea bream Sparus aurata, an

asymptomatic contagious fish host for nodavirus. Diseases of Aquatic Organisms 47: 33-38.

Catalano SR (2010a). Parasites of Australian herring/Tommy Ruff. .

http://www.marineparasites.com/pdf/Australian_herring.pdf

Catalano SR (2010b). Parasites of western Australian salmon (Arripis truttaceus). .

http://www.marineparasites.com/pdf/Australian%20salmon.pdf

http://www.mnr.gov.on.ca/en/Business/LetsFish/2ColumnSubPage/STEL02_165361.html

http://www.scahls.org.au/__data/assets/pdf_file/0010/1516519/Yersiniosis.pdf

http://www.marineparasites.com/pdf/Australian_herring.pdf

http://www.marineparasites.com/pdf/Australian salmon.pdf

_________________________________________________________________________________________________________________________________________________________________________________


233

www.digsfish.com

Catap ES, Lavilla-Pitago CR, Maeno Y, Travina RD (2003). Occurrence, histopathology and experimental

transmission of hepatopancreatic parvovirus infection in Penaeus monodon postlarvae. Diseases of Aquatic


Chang PH, Lin CW, Lee YC (2002). Lactococcus garvieae infection in cultured rainbow trout,

Oncorhynchus mykiss, in Taiwan and associated biophysical characteristics and histopathology. Bulletin of

the European Association of Fish Pathologists 22: 319-326.

Chang PH, Kuo ST, Lai SH, Yang HS, Ting YY, Hsu CL, Chen HC (2005). Herpes-like virus infection

causing mortality of cultured abalone Haliotis diversicolor supertexta in Taiwan. Diseases of Aquatic


Chatton E, Poisson R (1931). Sur l’existence, dans le sang des crabs, de peridiniens parasites:

Hematodinium perezi n.g., n.sp. (Syndinidae). CR Sceances Soc Biol Paris 105: 553–557.

Chapman A, Hobbs RP, Morgan DL, Gill HS (2006). Helminth parasitism of Galaxias maculatus (Jenyns

1842) in southwestern Australia. Ecology of Freshwater Fish 15: 559–564.

Chen SN, Chang PS, Kou GH (1990). Infection route and eradication of Penaeus monodon baculovirus

(MBV) in larval giant tiger prawns, Penaeus monodon. In: Diseases of Cultured Penaeid Shrimp in Asia and

the United States, Fulks W. & Main K.L., eds. Oceanic Institute, Honolulu, Hawaii, USA, pgs. 177-184.

Chen MF, Yun S, Marty GD, McDowell TS, House ML, Appersen JA, Guenther TA, Arkush KD, Hedrick

RP (2000a). A Piscirickettsia salmonis-like bacterium associated with mortality in white seabass

Atractoscion noblis. Diseases of Aquatic Organisms 43: 117-126.

Chen SC, Wang PC, Tung MC, Thompson KD, Adams A (2000b). A Piscirickettsia salmonis-like organism

in grouper, Epinephelus melanostigma, in Taiwan. Journal of Fish Diseases 23: 415-418.

Chen HG, Chen HY, Wang CS, Chen SN, Shih HH (2010). Effects of various treatments on egg hatching of

Dendromonocotyle pipinna (Monogenea: Monocotylidae) infecting the blotched fantail ray, Taeniurops

meyeni, in Taiwan. Veterinary Parasitology 171: 229–237.

Cheslett D, McKiernan F, Hickey C, Collins E (2009). Viral gametocytic hypertrophy of the Pacific oyster

Crassostrea gigas in Ireland. Diseases of Aquatic Organisms 83:181-185.

Choi DL, Lee NS, Choi HJ, Park MA, McGladdery SE, Park MS (2004). Viral gametocytic hypertrophy

caused by a papova-like virus infection in the Pacific oyster Crassostrea gigas in Korea. Diseases of Aquatic


Chong YC and Loh H (1984). Hepatopancreas chlamydial and parvoviral infections of farmed and marine

prawns in Singapore. Singapore Veterinary Journal 9: 51-56.

_________________________________________________________________________________________________________________________________________________________________________________


234

www.digsfish.com

Choudhury A, Charipar E, Nelson P, Hodgson JR, Bonar S, Cole RA (2006). Update on the distribution of

the invasive Asian fish tapeworm, Bothriocephalus acheilognathi, in the U.S. and Canada. Comparative

Parasitology 73: 269-273.

Chu FL, Greene KH (1989). Effect of temperature and salinity on the invitro culture of the oyster pathogen

Perkinsus marinus (Apicomplexa: Perkinsea). Journal of Invertebrate Pathology 53: 260-268.

Cigarria J, Elston R (1997). Independent introduction of Bonamia ostreae, a parasite of Ostrea edulis, to

Spain. Diseases of Aquatic Organisms 29: 157-158.

Cochennec-Laureau N, Reece KS, Berthe FJC, Hine PM (2003). Mikrocytos roughleyi taxonomic affiliation

leads to the genus Bonamia (Haplosporidia). Diseases of Aquatic Organisms 54: 209-217.

Cohen AN, Weinstein A, Emmett MA, Lau W, Carlton JT (2001). Investigations into the introduction of

non-indigenous marine organisms via the cross-continental trade in marine baitworms. Report for the U.S.

Fish and Wildlife Service, Sacramento CA. 29 pgs.

Colorni A, Samocha T and Colorni B (1987). Pathogenic viruses introduced into Israeli mariculture systems

by imported penaeid shrimp. Bamidgeh 39: 21-28.

Colquhoun DJ, Duodu S (2011). Francisella infections in farmed and wild aquatic organisms. Veterinary

Research 42: 47

Comtet T, Garcia C, LeCoguic Y, Joly JP (2004). First record of the microsporidian parasite Steinhausia

mytilovum in Mytilus sp. (Bivalvia: Mytilidae) from France. Diseases of Aquatic Organisms 58: 261-264.

Corbeil S, Crane MSJ (2009). Piscirickettsia salmonis. Australian and New Zealand Standard Diagnostic

Procedure. http://www.scahls.org.au/__data/assets/pdf_file/0005/1516514/Piscirickettsia_salmonis.pdf

Corbeil S, Hyatt AD, Crane MSJ (2005). Characterisation of an emerging rickettsia-like organism in

Tasmanian farmed Atlantic salmon Salmo salar. Diseases of Aquatic Organisms 64: 37-44.

Corbeil S, Arzul I, Robert M, Berthe F, Besnard-Cochennec N, Crane M, (2006). Molecular characterisation

of an Australian isolate of Bonamia exitiosa. Diseases of Aquatic Organisms 71: 81–85.

Corbeil S, Handlinger J, Crane MSJ (2009). Bonamiosis in Ostrea angasi. Australian and New Zealand

Standard Diagnostic Procedure. http://www.scahls.org.au/__data/assets/pdf_file/0003/1516512/Bonamia.pdf

Corbeil S and 16 other co-authors (2010). Development and validation of a TaqMan® PCR assay for the

Australian abalone herpes-like virus. Diseases of Aquatic Organisms 92: 1-10.

Corfield J, Diggles B, Jubb C, McDowall RM, Moore A, Richards A, Rowe DK (2007). Draft final report for

public comment. Review of the impacts of introduced aquarium fish species that have established wild

http://www.scahls.org.au/__data/assets/pdf_file/0005/1516514/Piscirickettsia_salmonis.pdf

http://www.scahls.org.au/__data/assets/pdf_file/0003/1516512/Bonamia.pdf

_________________________________________________________________________________________________________________________________________________________________________________


235

www.digsfish.com

populations in Australia. Prepared for the Australian Government Department of the Environment and Water

Resources. http://www.environment.gov.au/biodiversity/invasive/publications/pubs/wild-aquarium-fish.pdf

Coughran J, McCormack RB, Daly G (2009). Translocation of the Yabby Cherax destructor into eastern

drainages of New South Wales, Australia. Australian Zoologist 35: 100-103.

Coughran J, Leckie SR (2007). Invasion of a New South Wales stream by the Tropical Crayfish, Cherax

quadricarinatus (von Martens). In: Lunney D, Ebs P, Hutchings P, Burgin S (Eds.). Pest or Guest: the

Ecology of Overabundance. Royal Zoological Society of New South Wales. Pgs 40-46.

Cowley JA, Dimmock CM,Wongteerasupaya C, Boonsaeng V, Panyim S, Walker PJ (1999). Yellow head

virus from Thailand and gill-associated virus from Australia are closely related but distinct viruses. Diseases

of Aquatic Organisms 36: 153–175.

Cowley JA, Dimmock CM, Spann KM, Walker PJ (2000). Detection of Australian gill-associated virus

(GAV) and lymphoid organ virus (LOV) of Penaeus monodon by RT-nested PCR. Diseases of Aquatic


Cowley JA, Hall MR, Cadogan LC, Spann KM, Walker PJ (2002). Vertical transmission of gill-associated

virus (GAV) in the black tiger prawn Penaeus monodon. Diseases of Aquatic Organisms 50: 95-104.

Cowley JA, Cadogan LC, Wongteerasupaya C, Hodgson RAJ, Boonsaeng V, Walker PJ (2004). Multiplex

RT-nested PCR differentiation of gill-associated virus (Australia) from yellow head virus (Thailand) of

Penaeus monodon. Journal of Virological Methods 117: 49-59.

Cowley JA, McCulloch RJ, Rajendran KV, Cadogan LC, Spann KM, Walker PJ (2005). RT-nested PCR

detection of Mourilyan virus in Australian Penaeus monodon and its tissue distribution in healthy and

moribund prawns. Diseases of Aquatic Organisms 66: 91-104.

Cowley JA, Coman GJ, Salmon ML, Young ND, Rajendran KV, Wilson KJ, Preston NP (2009). In situ

stress testing to identify Australian black tiger prawns (Penaeus monodon) free of gill-associated virus and

Mourilyan virus. Australian Veterinary Journal 87: 244-248.

Crane MJ and 8 other authors. (2000). First isolation of an aquatic birnavirus from farmed and wild fish

species in Australia. Diseases of Aquatic Organisms 43: 1-14.

Crane MJ, Fegan M, Corbeil S, Warner S (2009). Development of Molecular Diagnostic Procedures for the

Detection and Identification of Herpes-like Virus of Abalone (Haliotis spp.). Final Report to the Fisheries

Research and Development Corporation. Project Number 2007-006.

Cranfield HJ, Dunn A, Doonan IJ, Michael KP (2005). Bonamia exitiosa epizootic in Ostrea chilensis from

Foveaux Strait, southern New Zealand between 1986 and 1992. ICES Journal of Marine Science 62: 3-13.

http://www.environment.gov.au/biodiversity/invasive/publications/pubs/wild-aquarium-fish.pdf

_________________________________________________________________________________________________________________________________________________________________________________


236

www.digsfish.com

Crawford SE (2001). Live Rockweed (Ascophyllum) used as a Shipping Medium for the Live Transport of

Marine Baitworms from Maine. In: Paust BC and Rice AA (eds.) Marketing and shipping live aquatic

products: Proceedings of the Second International Conference and Exhibition, November 1999, Seattle, WA.

University of Alaska Sea Grant, AK-SG-01-03. http://nsgl.gso.uri.edu/aku/akuw99003.pdf

Cribb J (2010). Science brings salvation for oyster farmer. FRDC News FISH 18(1): 4-5.

Crockford M (2007). Partial characterization of pilchard herpesvirus and the associated disease in pilchards.

Thesis presented for the degree of Doctor of Philosophy at Murdoch University. 108 pgs.

Crosbie PB, Munday BL (1999). Environmental factors and chemical agents affecting the growth of the

pathogenic marine ciliate Uronema nigricans. Diseases of Aquatic Organisms 36: 213-219.

Daszak PA, Berger L, Cunningham A, Hyatt AD, Green DE, Speare R (1999). Emerging infectious diseases

and amphibian population declines. Emerging Infectious Diseases 5: 735-748.

Daszak PA, Cunningham A, Hyatt AD (2000). Emerging infectious diseases of wildlife: Threats to

biodiversity and human health. Science 287: 443-449.

Daszak PA, Cunningham A, Hyatt AD (2003). Infectious diseases and amphibian population declines.

Diversity and Distributions 9: 141-150.

Dauer DM, Maybury CA, Ewing RM (1981). Feeding behavior and general ecology of several spionid

polychaetes from the Chesapeake Bay. Journal of Experimental Marine Biology and Ecology 54: 21-38.

Davies RL (1990). O-serotyping of Yersinia ruckeri with special emphasis on European isolates. Veterinary


Davies S, Mair G, Harris J (2008). Baitshop survey report 2008. With a focus on the supply and demand of

baitworms in South Australia. Flinders University. 27 pgs.

Davies S, Mair G, Harris J, Safari A (2010a). Worm aquaculture. Is there a demand in South Australia ?

Austasia Aquaculture 24(1): 56-59.

Davies KR, McColl KA, Wang LF, Yu M, Williams LM, Crane MJ (2010b). Molecular characterisation of

Australasian isolates of aquatic birnaviruses. Diseases of Aquatic Organisms 93: 1-15

DEEDI (2010). Recreational fishing rules and regulations for Queensland. A brief guide. Department of

Employment, Economic Development and Innovation, Queensland Government January 2010.

DeJonckheere JF (1979). Occurrence of Naegleria and Acanthamoeba in aquaria. Applied Environmental


http://nsgl.gso.uri.edu/aku/akuw99003.pdf

_________________________________________________________________________________________________________________________________________________________________________________


237

www.digsfish.com

De La Pena LD, Lavilla-Pitogo CR, Villar CB, Paner MG, Capulos GC (2008). Prevalence of monodon

baculovirus (MBV) in wild shrimp Penaeus monodon in the Philippines. Aquaculture 285: 19-22.

Delvinquier BLJ (1989). Occurrence of the protozoans, Lankesterella hylae, and Haemogregarina sp., in the

blood of the green tree frog, Litoria caerulea. Memoirs of the Queensland Museum 1989: 27: 267-273.

Delvinquier BLJ, Freeland WJ (1988). Protozoan parasites of the cane toad, Bufo marinus, in Australia.

Australian Journal of Zoology 36: 301-316.

Dennis DM, Munday BL (1994). Microsporidosis of palinurid lobsters from Australian waters. Bulletin of

the European Association of Fish Pathologists 14: 16-18.

Desportes I, Perkins FO (1990). Phylum Paramyxea. In: Handbook of Protoctista, L. Margulis, J. O.

Corliss, M. Melkonian and D.J. Chapman, (eds.). Jones and Bartlett Publ., Boston, pgs. 30-35.

Deveney MR, Bayly TJ, Johnston CJ, Nowak BF (2005). A parasite survey of farmed southern bluefin tuna ,

Thunnus maccoyii (Castelnau). Journal of Fish Diseases 28: 279-284.

Diamant A (1997). Fish to fish transmission of a marine myxosporean. Diseases of Aquatic Organisms 30:

99-105.

Diggles BK (2002). Import Risk Assessment: Juvenile yellowtail kingfish (Seriola lalandi) from Spencer

Gulf Aquaculture, South Australia. NIWA Client Report WLG 2002/03. .

http://www.biosecurity.govt.nz/files/regs/imports/risk/yellowtail-kingfish-ra.pdf

Diggles BK (2003). Shellfish Disease Surveillance Programme – Final report June 2003. NIWA Client

Report AUS2002-018, prepared for Primary Industries and Resources, South Australia (PIRSA). 24 pgs.

Diggles BK (2005). Import Risk Analysis: Ornamental fish. Biosecurity NZ, Ministry of Agriculture and

Forestry. IRA prepared by DigsFish Services Pty Ltd for the NZ Government. 264 pgs

Diggles BK (2007). Scientific submission to AQIS supporting an application for an import permit allowing

entry of whole, round menhaden (Brevoortia tyrannus) for use as lobster bait. DigsFish Services Client

Report DF07-01 prepared for the Western Rock Lobster Development Association (WRLDA). 39 pgs.

Diggles BK, Hine PM (2002). Bonamia exitiosus epidemiology in Foveaux Strait oysters. Final research

report, OYS1999/01, Ministry of Fisheries, New Zealand. 51 pgs. (Unpublished report held in NIWA

library, Wellington.)

Diggles BK, Carson J, Hine PM, Tait M (2000). Vibrio species associated with mortalities in hatchery reared

turbot (Colistium nudipinnis) and brill (C. guntheri) in New Zealand. Aquaculture 183: 1-12.

http://www.biosecurity.govt.nz/files/regs/imports/risk/yellowtail-kingfish-ra.pdf

_________________________________________________________________________________________________________________________________________________________________________________


238

www.digsfish.com

Diggles BK, Nichol J, Hine PM, Wakefield S, Cochennec-Laureau N, Roberts RD, Friedman CS (2002).

Pathology of cultured paua (Haliotis iris Martyn, 1784) infected by a novel haplosporidian parasite, with

some observations on the course of disease. Diseases of Aquatic Organisms 50: 219-231.

Diggles BK, Cochennec-Laureau N, Hine M (2003). Comparison of diagnostic techniques for Bonamia

exitiosus from flat oysters Ostrea chilensis in New Zealand. Aquaculture 220: 145-156.

Dinamani P, Hine PM, Jones JB (1987). Occurrence and characteristics of the haemocyte parasite Bonamia

sp. in the New Zealand dredge oyster Tiostrea lutaria. Diseases of Aquatic Organisms 3: 37-44.

Doonan IJ, Cranfield HJ, Michael KP (1994). Catastrophic reduction of the oyster, Tiostrea chilensis (Bivalvia:

Ostreidae), in Foveaux Strait, New Zealand, due to infestation by the protistan Bonamia sp. NZ Journal of

Marine and Freshwater Research 28: 335-344.

Di Stefano RJ, Litvan ME, Horner PT (2009). The bait industry as a potential vector for alien crayfish

introductions: Problem recognition by fisheries agencies and a Missouri evaluation. Fisheries 34(12): 586-

597.

Doubrovsky A, Paynter JL, Sambhi SK, Atherton JG, Lester RJG (1988). Observations on the ultrastructure

of baculovirus in Australian Penaeus monodon and Penaeus merguiensis. Australian Journal of Marine and


Dove ADM (1998). A silent tragedy: parasites and exotic fishes of Australia. Proceedings of the Royal

Society Queensland 107: 109-113.

Dove ADM (2000). Richness patterns in the parasite communities of exotic poeciliid fishes. Parasitology

120: 609-623.

Dove ADM, Ernst I (1998). Concurrent invaders - four exotic species of Monogenea now established on

exotic freshwater fishes in Australia. International Journal of Parasitology 28: 1755-1764.

Dove ADM, Fletcher AS (2000). The distribution of the introduced tapeworm Bothriocephalus

acheilognathi in Australian freshwater fishes. Journal of Helminthology 74: 121-127.

Dove ADM, O’Donoghue PJ (2005). Trichodinids (Ciliophora: Trichodinidae) from native and exotic

Australian freshwater fishes. Acta Protozoologica 44: 51-60.

Dove ADM, Cribb TH, Mockler SP, Lintermans M (1997). The Asian tapeworm, Bothriocephalus

acheilognathi, in Australian freshwater fishes. Marine and Freshwater Research 48: 181-183.

DPIF (2004). Management arrangements for potentially high-risk activities in the context of ecologically

sustainable development (ESD) for approved aquaculture operations, Aquaculture Policy FAMOP001. 53

pgs. http://www2.dpi.qld.gov.au/extra/pdf/fishweb/FAMOP001.pdf

http://www2.dpi.qld.gov.au/extra/pdf/fishweb/FAMOP001.pdf

_________________________________________________________________________________________________________________________________________________________________________________


239

www.digsfish.com

Durand SV, Tang KFJ, Lightner DV (2000). Frozen commodity shrimp: potential avenue for introduction of

white spot syndrome virus and yellowhead virus. Journal of Aquatic Animal Health 12: 128-135.

Dykova I, Lom J, Machackova B, Sawyer TK (1996). Amoebic infections in goldfishes and granulomatous

lesions. Folia Parasitologica 43: 81-90.

Dykova I, Lom J, Machackova B, Peckova H (1998). Vexillifera expectata sp. n. and other non-encysting

amoebae isolated from organs of freshwater fish. Folia Parasitologica 45: 17-26.

Dyková I, Kyselová I, Pecková H, Oborník M, Luke J (2001). Identity of Naegleria strains isolated from

organs of freshwater fishes. Diseases of Aquatic Organisms 46: 115-121.

Dykstra MJ, Levine JF, Noga EJ, hawkins JH, Gerdes P, Hargis WJ Jr., Grier HJ, Te Strake D (1989).

Ulcerative mycosis: a serious menhaden disease of the south eastern coastal fisheries of the United States.

Journal of Fish Diseases 12: 175-178.

Eaton WD, Love DC, Botelho C, Meyers TR, Imamura K, Koeneman T (1991). Preliminary results on the

seasonality and life cycle of the parasitic dinoflagellate causing Bitter Crab Disease in Alaskan Tanner crabs

(Chionoecetes bairdi). Journal of Invertebrate Pathology 57: 426–434.

Eaves LE, Ketterer PJ (1994). Mortalities in red claw crayfish Cherax quadricarinatus associated with

systemic Vibrio mimicus infection. Diseases of Aquatic Organisms 19: 233-237.

Eddy F, Powell A, Gregory S, Nunan LM, Lightner DV, Dyson PJ, Rowley AF, Shields RJ (2007). A novel

bacterial disease of the European shore crab, Carcinus maenas – molecular pathology and epidemiology.


Edgerton BF (1996). Viruses of freshwater crayfish. PhD Thesis, James Cook University of North

Queensland, Townsville, Australia, 152 pp.

Edgerton BF, Owens L (1997). Age at first infection of Cherax quadricarinatus by Cherax quadricarinatus

bacilliform virus and Cherax giardiavirus-like virus, and production of putative virus-free crayfish.

Aquaculture 152: 1– 12.

Edgerton BF, Prior HC (1999). Description of a hepatopancreatic rickettsia-like organism in the redclaw

crayfish Cherax quadricarinatus .Diseases of Aquatic Organisms 36: 77-80.

Edgerton BF, Webb, R, Wingfield M (1997). Systemic parvo-like virus in the freshwater crayfish Cherax

destructor. Diseases of Aquatic Organisms 29: 73-78.

Edgerton BF, Evans LH, Stephens FJ, Overstreet RM (2002). Synopsis of freshwater crayfish diseases and

commensal organisms. Aquaculture 206: 57-135.

_________________________________________________________________________________________________________________________________________________________________________________


240

www.digsfish.com

Eldar A, Ghittino C (1999). Lactococcus garvieae and Streptococcus iniae infections in rainbow trout

Oncorhynchus mykiss : similar but different diseases. Diseases of Aquatic Organisms 36: 227-231.

Eldar A, Ghittino C, Asanta L, Bozzetta E, Goria M, Prearo M, Bercovier H (1996). Enterococcus

seriolicida is a junior synonym of Lactococcus garvieae, a causative agent of septicaemia and

meningoencephalitis in fish. Current Microbiology 32: 85-88.

Eldar A, Goria M, Ghittino C, Zlotkin A, Bercovier H (1999). Biodiversity of Lactococcus garvieae strains

isolated from fish in Europe, Asia and Australia. Applied and Environmental Microbiology 65: 1005-1008.

Elliot J (2001). Tasmania - quarterly report- Nodavirus in striped trumpeter, Rickettsia-like organisms in

salmonids. Animal Health Surveillance 6 (1): 13-14.

El-Matbouli M, Hoffmann RW (1991). Effects of freezing, aging, and passage through the alimentary canal

of predatory animals on the viability of Myxobolus cerebralis spores. Journal of Aquatic Animal Health 3:

260–262.

El-Matbouli M, Soliman H (2011). Transmission of Cyprinid herpesvirus-3 (CyHV-3) from goldfish to

naïve common carp by cohabitation. Research in Veterinary Science doi:10.1016/j.rvsc.2010.07.008

Elsayed E, Faisal M, Thomas M, Whelan G, Batts W and Winton J (2006). Isolation of viral haemorrhagic

septicaemia virus from muskellunge, Esox masquinongy (Mitchill), in Lake St-Clair, Michigan, USA reveals

a new sublineage of the North American genotype. Journal of Fish Diseases 29: 611–619.

Elston R (1980). Ultrastructural aspects of a serious disease of hatchery reared larval oysters, Crassostrea

gigas Thünberg. Journal of Fish Diseases 3: 1-10.

Elston R (1997). Special topic review: bivalve mollusc viruses. World Journal of Microbiology and

Biotechnology 13: 393-403.

Elston RA, Farley CA, Kent ML (1986). Occurrence and significance of bonamiosis in European flat oysters

Ostrea edulis in North America. Diseases of Aquatic Organisms 2: 49-54.

Englund RA, Eldredge LG (2001). Fishes. In: Hawaii's invasive species (Staples, G.W.; Cowie, R.H., eds),

Bishop Museum, Honolulu, pgs. 32-40.

EPA (2007). Recreational fishing in protected areas. Operational Policy, Visitor Management. Queensland

Government, Environmental Protection Agency. http://www.derm.qld.gov.au/register/p01235aa.pdf

Ernst I, Whittington I (2001). Experimental susceptibility of some reef fish species to Benedenia lutjani

(Monogenea: capsalidae), a parasite of Lutjanus carponotatus (Pisces: Lutjanidae). Parasitology Research

87: 345-348.

http://www.derm.qld.gov.au/register/p01235aa.pdf

_________________________________________________________________________________________________________________________________________________________________________________


241

www.digsfish.com

Ernst I, Fletcher A, Hayward C (2000). Gyrodactylus anguillae (Monogenea: Gyrodactylidae) from anguillid

eels (Anguilla australis and Anguilla reinhardtii) in Australia: A Native or an Exotic? Journal of


Ernst I, Whittington ID, Corneillie S, Talbot C (2005). Effects of temperature, salinity, desiccation and

chemical treatments on egg embryonation and hatchng success of Benedenia seriolae (Monogenea:

Capsalidae), a parasite of farmed Seriola spp. Journal of Fish Diseases 28: 157-164.

Evans BB, Lester RJG (2001). Parasites of ornamental fish imported into Australia. Bulletin of the

Euri\opean Association of Fish Pathologists 21: 51-55.

Evans JJ, Klesius PH, Shoemaker CA (2009). First isolation and characterization of Lactococcus garvieae

from Brazilian Nile tilapia, Oreochromis niloticus (L.), and pintado, Pseudoplathystoma corruscans (Spix &

Agassiz). Journal of Fish Diseases 32: 943-951.

Evans L, Edgerton BF, Stephens FJ, Cabalu HJ, Overstreet RM (1998). Scientific Review of Freshwater

Crayfish Diseases and Commensal Organisms. Report Prepared for AQIS .

http://www.daff.gov.au/__data/assets/word_doc/0018/11583/fw_crayfish_dis_pro_rev.doc

Ewing GP, Lyle JM (2009). Reproductive dynamics of redbait, Emmelichthys nitidus (Emmelichthyidae),

from south-eastern Australia. Fisheries Research 97: 206-215.

Faisal M, Schulz CA (2009). Detection of Viral Haemorrhagic Septicaemia virus (VHSV) from the leech

Myzobdella lugubris Leidy, 1851. Parasites and Vectors 2: 45 .

http://www.parasitesandvectors.com/content/2/1/45

Farley CA, Wolf PH, Elston RA (1988). Long term study of microcell disease in oysters with a description

of a new genus, Mikrocytos (g.n.) and two new species, Mikrocytos mackini and Mikrocytos roughleyi.

Fisheries Bulletin 86: 581-593.

Fauquet CM, Mayo MA, Maniloff J, Desselberger U, Ball LA (2005). Virus taxonomy: classification and

nomenclature of viruses: eighth report of the International Committee on the Taxonomy of Viruses Elsevier

Academic Press. San Diego, California.

Fegan DF, Flegel TW, Sriurairatana S, Waiyakruttha M (1991). The occurrence, development and

histopathology of monodon baculovirus in Southern Thailand. Aquaculture 96: 205-217.

Ferguson G, Mayfield S (2006). The South Ausralian Goolwa Cockle (Donax deltoides) Fishery. SARDI

Aquatic Sciences Publication No. RD06/0005-1, SARDI Research Report Series No. 150. 30 pgs.

Field RH, Chapman CJ, Taylor AC, Neil DM, Vickerman K (1992). Infection of the Norway lobster

Nephrops norvegicus by a Hematodinium-like species of dinoflagellate on the west coast of Scotland.

Diseaes of Aquatic Organisms 13: 1–15.

http://www.daff.gov.au/__data/assets/word_doc/0018/11583/fw_crayfish_dis_pro_rev.doc

http://www.parasitesandvectors.com/content/2/1/45

_________________________________________________________________________________________________________________________________________________________________________________


242

www.digsfish.com

Fisher MC, Garner TWJ (2007). The relationship between the emergence of Batrachochytrium

dendrobatidis, the international trade in amphibians and introduced amphibian species. Fungal Biology

Reviews 21: 2-9.

Flegel TW (1997). Special topic review: Major viral diseases of the black tiger prawn (Penaeus monodon)

in Thailand. World J. Microbiol. Biotechnol. 13: 433-442.

Flegel TW (2006). Detection of major penaeid shrimp viruses in Asia, a historical perspective with emphasis

on Thailand. Aquaculture 258: 1-33.

Flegel TW (2009). Review of disease transmission risks from prawn products exported for human

consumption. Aquaculture 290: 179-326.

Flegel TW, Boonyaratpalin S, Fegan DF, Guerin M, Sriurairatana S (1992). High mortality of black tiger

prawns from cotton shrimp disease in Thailand. In: Diseases in Asian Aquaculture I: 181-197.

Flegel TW, Sriurairatana S, Wongteerasupaya C, Boonsaeng V, Panyim S, Withyachumnarnkul B (1995).

Progress in characterization and control of yellow-head virus of Penaeus monodon. In: Browdy C, Hopkins S

(eds) Proc Special Session on Shrimp Farming, Aquaculture ’95, San Diago, February 1995. World

Aquaculture Society, Baton Rouge, FL, pgs. 76–83.

Flegel TW, Sriurairatana S, Wongteerasupaya C, Boonsaeng V, Panyim S, Withyachumnarnkul B (1997).

Progress in characterization and control of yellow-head virus of Penaeus monodon. In: Flegel TW,

Menasveta P, Paisarnrat S. Shrimp biotechnology in Thailand. National Center for Genetic Engineering and

Biotechnology, Bangkok, pgs. 71–78.

Flegel TW, Thamavit V, Parawipas T, Alday-Sanz V (1999). Statistical correlation between severity of

hepatopancreatic parvovirus infection and stunting of farmed black tiger shrimp (Penaeus monodon).

Aquaculture 174: 197–206.

Flegel TW, Nielsen L, Thamavit V, Kongtim S, Pasharawipas T (2004). Presence of multiple viruses in non-

diseased, cultivated shrimp at harvest. Aquaculture 240: 55-68.

Fletcher AS and Whittington ID (1998). A parasite-host checklist for Monogenea from freshwater fishes in

Australia, with comments on biodiversity. Systematic Parasitology 41: 159-168

Fletcher WJ, Jones JB, Pearce AF, Hosja W (1997). Environmental and biological aspects of the mass

mortality of pilchards (Autumn 1995) in Western Australia. Fisheries Department of WA Fisheries Research

Report No. 106. 112 pgs.

Font WF (1998). Parasites in paradise: patterns of helminth distribution in Hawaiian stream fishes. Journal

of Helminthology 72: 307-311.

_________________________________________________________________________________________________________________________________________________________________________________


243

www.digsfish.com

Font WF (2003). The global spread of parasites: What do Hawaiian streams tell us ? BioScience 53: 1061-

1067.

Font WF, Tate DC (1994). Helminth parasites of native Hawaiian freshwater fishes: an example of extreme

ecological isolation. Journal of Parasitology 80: 682-688.

Ford SE, Xu Z, Debrosse G (2001). Use of particle filtration and UV irradiation to prevent infection by

Haplosporidium nelsoni (MSX) and Perkinsus marinus (Dermo) in hatchery reared larval and juvenile

oysters. Aquaculture 194: 37-49.

Ford SE, Allam B, Xu Z (2009). Using bivalves as particle collectors with PCR detection to investigate the

environmental distribution of Haplosporidium nelsoni. Diseases of Aquatic Organisms 83: 159-168.

Foschino R, Nucera D, Volponi G, Picozzi C, Ortoffi M, Bottero MT (2008). Comparison of Lactococcus

garvieae strains isolated in northern Italy from dairy products and fishes through molecular typing. Journal

of Applied Microbiology 105: 652-662.

Fraser CA, Owens L (1996). Spawner-isolated mortality virus from Australian Penaeus monodon. Diseases


Fraser GC, Callinan RB, Calder LM (1992) Aphanomyces species associated with red spot disease: an

ulcerative disease of estuarine fish from eastern Australia. Journal of Fish Diseases 15: 173–181.

Freeman MA, Turnbull JF, Yeomans WE, Bean CW (2009). Prospects for management strategies of invasive

crayfish populations with an emphasis on biological control. Aquatic Conservation 20: 211-223.

Frerichs GN, Tweedie A, Starkey WG and Richards RH (2000). Temperature, pH and electrolyte sensitivity

and heat, UV and disinfectant inactivation of sea bass (Dicentrarchus labrax) neuropathy nodavirus.

Aquaculture 185: 13-24.

Friedman CS (1996). Haplosporidian infections of the pacific oyster, Crassostrea gigas (Thunberg), in

California and Japan. Journal of Shellfish Research 15: 597-600.

Friedman CS, Andree KB, Beauchamp KA, Moore JD, Robbins TT, Shields JD (2000). Candidatus

Xenohaliotis californiensis is a newly described pathogen of abalone, Haliotis spp., along the west coast of

North America. International Journal of Systematic Bacteriology 50: 847–855.

Friedman CS, Estes RM, Stokes NA, Burge CA, Hargrove JS, Barber BJ, Elston RA, Burreson EM, Reece

KS (2005). Herpes virus in juvenile Pacific oysters Crassostrea gigas from Tomales Bay, California,

coincides with summer mortality episodes. Diseases of Aquatic Organisms 63: 33-41.

Frischer ME, Lee RF, Sheppard MA, Mauer A and others (2006). Evidence for a free-living life stage of the

blue crab parasitic dinoflagellate, Hematodinium sp. Harmful Algae 5: 548–557.

_________________________________________________________________________________________________________________________________________________________________________________


244

www.digsfish.com

Gagne´ N, Mackinnon AM, Boston L, Souter B, Cook-Versloot M, Griffiths S and Olivier G (2007).

Isolation of viral haemorrhagic septicaemia virus from mummichog, stickleback, striped bass and brown

trout in eastern Canada. Journal of Fish Diseases 30:213–223.

Gangnonngiw W, Kiatpathomchai W, Sriurairatana S, Laisutisan K, Chuchird N, Limsuwan C, Flegel TW

(2009). Parvo-like virus in the hepatopancreas of freshwater prawns Macrobrachium rosenbergii cultivated

in Thailand. Diseases of Aquatic Organisms 85: 167-173.

Gangnonngiw W, Laisutisan K, Sriurairatana S, Senapin S, Chuchird N, Limsuwan C, Chaivisuthangkura P,

Flegel TW (2010). Monodon baculovirus (MBV) infects the freshwater prawn Macrobrachium rosenbergii

cultivated in Thailand. Virus Research 148: 24-30.

Gao F, Fan X, Yi Z, Strüder-Kypke M, Song W (2010). Phylogenetic consideration of two scuticociliate

genera, Philasterides and Boveria (Protozoa, Ciliophora) based on 18 S rRNA gene sequences. Parasitology

International 59: 549–555.

Garcia C, Robert M, Arzul I, Chollet B, Joly JP, Miossec L, Comtet T, Berthe F (2006). Viral gametocytic

hypertrophy of Crassostrea gigas in France: from occasional records to disease emergence? Diseases of


Gardener GR, Harshbarger JC, Lake JL, Sawyer TK, Price KL, Stephenson MD, Haaker PL, Togstad HA

(1995). Association of prokaryotes with symptomatic appearance of withering syndrome in black abalone

Haliotis cracherodii. Journal of Invertebrate Pathology 66: 111-120.

Gates KK, Guy CS, Zale AV, Horton TB (2008). Adherence of Myxobolus cerebralis myxospores to

waders: implications for disease dissemination. North American Journal of Fisheries Management 28: 1453–

1458.

Gates KK, Guy CS, Zale AV, Horton TB (2009). Angler awareness of aquatic nuisance species and

potential transport mechanisms. Fisheries Management and Ecology 16: 448-456.

Gaughan DJ, Mitchell RW, Blight SJ (2000). Impact of mortality, possibly due to herpesvirus, on pilchard

Sardinops sagax stocks along the south coast of Western Australia in 1998–99. Marine and Freshwater

Research 51: 601–612.

Gaughan DJ (2002). Disease-translocation across geographic boundaries must be recognized as a risk even

in the absence of disease identification : the case with Australian Sardinops. Reviews in Fish Biology and

Fisheries 11: 113-123.

Ghisalberti EL, Chubb C, Park JO, Glendenning L (2004). Chemoattraction and the development of an

artificial bait for the western rock lobster (Panulirus cygnus). Fisheries Research and Development

Corporation Report FRDC Project No. 2000/255. ISBN 0-646-43190-0

_________________________________________________________________________________________________________________________________________________________________________________


245

www.digsfish.com

Gill PA, Callinan RB (1997). Ulcerative dermatitis associated with Uronema sp infection of farmed sand

whiting Sillago ciliata. Australian Veterinary Journal 75: 357.

Gleeson DJ, McCallum HI, Owens PF (2000). Differences in initial and acquired resistance to

Ichthyophthirius multifiliis between populations of rainbowfish. Journal of Fish Biology 57: 466-475.

Glenner H, Thomsen PF, Rybakov AV, Galil BS, Hoeg JT (2008). The Phylogeny of Rhizocephalan

Parasites of the Genus Heterosaccus using Molecular and Larval Data (Cirripedia: Rhizocephala;

Sacculinidae). Israel Journal of Ecology and Evolution 54: 223-238.

Goggin CL, Lester RJG (1987). Occurrence of Perkinsus species (Protozoa, Apicomplexa) in bivalves from

the Great Barrier Reef. Diseases of Aquatic Organisms 3: 113-117.

Goggin CL, Lester RJG (1995). Perkinsus, a protistan parasite of abalone in Australia: a review. Marine

and Freshwater Research 46: 639-646.

Goggin CL, Sewell KB, Lester RJG (1989). Cross-infection experiments with Australian Perkinsus species.

Diseases of Aquatic Organisms 7: 55-59

Goggin CL, Sewell KB, Lester RJG (1990). Tolerances of Perkinsus spp. (Protozoa, Apicomplexa) to

temperature, chlorine and salinity. Journal of Shellfish Research 9: 145–148.

Goodwin AE (2002). First report of spring viremia of carp virus (SVCV) in North America. Journal of

Aquatic Animal Health 14:161-164.

Goodwin AE, Peterson JE, Meyers TR, Money DJ (2004). Transmission of exotic fish viruses: the relative

risks of wild and cultured bait. Fisheries: Reviews in Fish Health, American Fisheries Society Fish Health

Section, 19-23.

Goodwin AE, Sadler J, Merry GE, Marecaux EN (2009). Herpesviral haematopoietic necrosis virus (CyHV-

2) infection: case studies from commercial goldfish farms. Journal of Fish Diseases 32: 271-278.

Gray MJ, Miller DL, Hoverman JT (2009). Ecology and pathology of amphibian ranaviruses. Diseases of


Green TJ, Barnes AC (2010). Reduced salinity, but not estuarine acidification, is a cause of immune-

suppression in the Sydney rock oyster Saccostrea glomerata. Marine Ecology Progress Series 402: 161-

170.

Green TJ, Raftos D, O'Connor W, Adlard RD, Barnes AC (2011). Disease Prevention Strategies for QX

Disease (Marteilia sydneyi) of Sydney Rock Oysters (Saccostrea glomerata). Journal of Shellfish Research

30: 47-53.

_________________________________________________________________________________________________________________________________________________________________________________


246

www.digsfish.com

Grindley RM, Keogh JA, Friedman CS (1998). Shell lesions in New Zealand Haliotis sp. (Mollusca,

Gastropoda). Journal of Shellfish Research 17: 805-811.

Grixti D, Conron SD Jones PL (2007). The effect of hook/bait size and angling technique on the hooking

location and the catch of recreationally caught black bream Acanthopagrus butcheri. Fisheries Research 84:

338-344.

Grizel H, Comps M, Bonami JR, Cousserans F, Duthoit JL, LePennec MA (1974). Research on the agent of

digestive gland disease of Ostrea edulis Linné. Science et Pêche Bulletin d'Information et de Documentation

de l'Institut Scientifique et Technique des Pêches Maritimes 240: 7-30. (In French).

Grizel H (1985). Etude des recentes epizooties de l'huitre plate Ostrea edulis Linne et de leur impact sur

l'osteiculture Bretonne. These, Academie de Montpellier, Universite des Sciences et Techniques du

Languedoc.

Grizzle JM, Altinok I, Fraser WA, Francis-Floyd R (2002). First isolation of largemouth bass virus. Diseases


Grizzle JM, Brunner CJ (2003). Review of Largemouth Bass Virus. Fisheries 28(11): 10-14.

Groocock GH, Getchell RG, Wooster GA, Britt KL, Batts WN, Winton JR, Casey RN, Casey JW, Bowser

PR (2007). Detection of viral hemorrhagic septicemia in round gobies in New York State (USA) waters of

Lake Ontario and the St. Lawrence River. Diseases of Aquatic Organisms 76: 87–192.

Groocock GH, Grimmett SG, Getchell RG, Wooster GA, Bowser PR (2008). A survey to determine the

presence and distribution of largemouth bass virus in wild freshwater bass in New York State. Journal of

Aquatic Animal Health 20: 158-164.

Grossel GW, Dykova I, Handlinger J, Munday BL (2003). Pentacapsula neurophila sp. N. (Multivalvulida)

from the central nervous system of striped trumpeter Latris lineata (Forster). Journal of Fish Diseases 26:

315-320.

Grutter AS (1994). Spatial and temporal variations of the ectoparasites of seven reef fish species from

Lizard Island and Heron Island, Australia. Marine Ecology Progress Series 115: 21-30.

Gunter NL, Burger MAA, Adlard RD (2010). Morphometric and molecular characterisation of four new

Ceratomyxa species (Myxosporea: Bivalvulida: Ceratomyxidae) from fishes off Lizard Island, Australia.

Folia Parasitologica 57: 1-10.

Gurney RH, Nowak BF, DykovaI, Kuris AM (2004). Histopathological effects of trypanorhynch

metacestodes in the digestive gland of a novel host, Carcinus maenas (Decapoda). Diseases of Aquatic

Organisms 58: 63-69

_________________________________________________________________________________________________________________________________________________________________________________


247

www.digsfish.com

Gustafson L (2009). Summary of VHS surveillance data analysis. NAHSS Outlook, Dec 2009.

http://www.aphis.usda.gov/vs/ceah/ncahs/nsu/outlook/issue23_dec09/outlook_dec09_vhs_surveillance.pdf

Halder M, Ahne W (1988). Freshwater crayfish Astacus astacus – a vector for infectious pancreatic necrosis

virus (IPNV). Diseases of Aquatic Organisms 4: 205-209.

Hall DN (1988). Occurrence of copepod parasite Lernea cyprinacea L. on the Australian grayling

Prototroctes maraena Gunther. Proceedings of the Royal Society of Victoria. 95: 273-274.

Hallett SL, Roubal FR (1995). Experiments on the infection dynamics of Caligus epidemicus (Copepoda:

Caligidae) on the small marine fish, Ambassis marianus (Günther). Journal of Fish Diseases 18: 59-66.

Hallett SL, Atkinson SD, Erseus C, El-Matbouli M (2006). Myxozoan parasites disseminated via

oligochaete worms as live food for aquarium fishes: descriptions of aurantiactinomyxon and raabeia

actinospore types. Diseases of Aquatic Organisms 69: 213-225.

Hamasaki K, Hatai K (1993). Prevention of fungal infection in the eggs and larvae of the swimming crab

Portunus trituberculatus and the mud crab Scylla serrata by bath treatment with formalin. Nippon Suisan

Gakkaishi 59: 1067-1072.

Handley SJ (2000). Larval development of Boccardia knoxi, a shell infesting spionid polychaete. New

Zealand Journal of Marine and Freshwater Research 34: 681-686.

Handlinger, J and 16 other co-authors (2006). Abalone Aquaculture Subprogram: A National Survey of

Diseases of Abalone Species to Support Trade and Translocation Issues and the Development of Health

Surveillance programs. FRDC Project No. 2002/201.

Hannon B (2008). An overview of the impacts of translocated native fish species in Australia. Sinclair

Knight Mertz report for the Australian Government Department of the Environment, Water, Heritage and

the Arts. August 2008. 66 pgs.

Haskin HH, Ford SE (1982). Haplosporidium nelsoni (MSX) on Delaware Bay seed oyster beds: A host-

parasite relationship along a salinity gradient. Journal of Invertebrate Pathology 40: 388-405.

Haskin HH, Andrews JD (1988). Uncertainties and speculations about the life cycle of the eastern oyster

pathogen Haplosporidium nelsoni (MSX). In: Fisher WS (ed) Disease processes in marine bivalve molluscs.

American Fisheries Society Special Publication 18: 5-22.

Hassan M. (2008). Parasites of native and exotic freshwater fishes in the south-west of Western Australia.

PhD Thesis, Murdoch University, Perth, Western Australia.

http://www.aphis.usda.gov/vs/ceah/ncahs/nsu/outlook/issue23_dec09/outlook_dec09_vhs_surveillance.pdf

_________________________________________________________________________________________________________________________________________________________________________________


248

www.digsfish.com

Hassan M, Beatty SJ, Morgan DL, Doupe RG, Lymbery AJ (2008). An introduced parasite, Lernaea

cyprinacea L. , found on native freshwater fishes in the south west of Western Australia. Journal of the

Royal Society of Western Australia 91: 149-153.

Hasson KW, Fan Y, Reisinger T, Venuti J, Varner PW (2006). White spot syndrome virus (WSSV)

introduction into the Gulf of Mexico and Texas freshwater systems through imported frozen bait shrimp.


Haswell WA (1885). On a destructive parasite of the rock oyster. Proceedings of the Linnean Society of New

South Wales /10: 272-275.

Hatai K (1982). On the fungus Haliphthoros milfordensis isolated from temporarily held abalone (Haliotis

sieboldii). Fish Pathology 17: 199-204.

Hayward CJ (1997). Distribution of external parasites indicates boundaries to dispersal of sillaginid fishes in

the Indo-West Pacific. Marine and Freshwater Research 48: 391-400.

Heasman M, Diggles BK, Hurwood D, Mather P, Pirozzi I, Dworjanyn S. (2004). Paving the way for

continued rapid development of the flat oyster (Ostrea angasi) farming industry in New South Wales. Final

Report to the Department of Transport & 590 Regional Services, Project No. NT002/0195. NSW Fisheries

Final Report No. 66. 40pgs. .

http://www.dpi.nsw.gov.au/__data/assets/pdf_file/0003/134481/Output-448.pdf

Heckmann RA, Greger PD, Furtek RC (1993). The Asian fish tapeworm, Bothriocephalus acheilognathi, in

fishes from Nevada. Journal of the Helminthological Society of Washington 60: 127-128.

Hedrick RP, McDowell TS (1995). Properties of iridoviruses from ornamental fish. Veterinary Research 26:

423–427.

Heiniger H, Gunter NL, Adlard RD (2011). Re-establishment of the family Coccomyxidae and description

of five novel species of Auerbachia and Coccomyxa (Myxospora: Bivalvulida) parasites of Australian fishes.


Hemaprasanth, RS, Raghavendra A, Sridhar N, Raghunath MR, Eknath AE (2011). Comparative

susceptibility of carp fingerlings to Lernaea cyprinacea infection. Veterinary Parasitolology

doi:10.1016/j.vetpar.2010.12.029

Hendrix PF, Callaham MA, Drake JM, Huang CY, James SW, Snyder BA, Zhang W (2008). Pandora’s box

contained bait: the global problem of introduced earthworms. Annual Review of Ecology, Evolution and

Systematics 39: 593–613.

Henry GW, Lyle JM (2003). The National Recreational and Indigenous Fishing Survey. FRDC Project No.

99/158. 188 pgs.

http://www.dpi.nsw.gov.au/__data/assets/pdf_file/0003/134481/Output-448.pdf

_________________________________________________________________________________________________________________________________________________________________________________


249

www.digsfish.com

Herbert BW (1987). Notes on the diseases and epibionts of Cherax quadricarinatus and C. tenuimanus

(Decapoda: Parastacidae). Aquaculture 64: 165-173.

Herbert BW (1988). Infection of Cherax quadricarinatus (Decapoda: Parastacidae) by the microsporidian

Thelohania sp. (Microspora: Nosematidae). Journal of Fish Diseases 11: 301-308.

Herfort A (2004). Aquatic animal diseases significant to Australia: identification field guide, Australian

Government Department of Agriculture, Fisheries and Forestry, Canberra. 91 pgs.

Hervio D, Bachere E, Boulo V, Cochennec N, Vuillemin V, LeCoguic Y, Cailletaux G, Mazurie J, Mialhe E.

(1995). Establishment of an experimental infection protocol for the flat oyster, Ostrea edulis, with the

intrahaemocytic protozoan parasite, Bonamia ostreae: application in the selection of parasite-resistant

oysters. Aquaculture 132: 183-194.

Hewitt GC (1971). Two species of Caligus (Copepoda, Caligidae) from Australian waters, with a

description of some developmental stages. Pacific Science 25: 145-164.

Hill KM, Carnegie RB, Aloui-Bejaoui N, Gharsalli RE, White DM, Stokes NA, Burreson EM (2010).

Observation of a Bonamia sp. infecting the oyster Ostrea stentina in Tunisia, and a consideration of its

phylogenetic affinities. Journal of Invertebrate Pathology 103: 179-185.

Hills J (2007). A review of the abalone virus ganglioneuritis (AVG). NZ Ministry of Fisheries .

http://www.biosecurity.govt.nz/files/pests/paua-avg-virus/avg-virus-review-julie-hills-nov07.pdfn

Hine PM (1995). Herpesviruses associated with mortalities among pilchards (Sardinops neopilchardus)

around New Zealand. Surveillance 22(3): 45-48.

Hine PM (1996a). The ecology of Bonamia and decline of bivalve molluscs. New Zealand Journal of

Ecology 20: 109–116.

Hine PM (1996b). Southern hemisphere mollusc diseases and an overview of associated risk assessment

problems. Rev. Sci. Tech. Off. Int. Epiz. 15: 563-577.

Hine PM (2002). Severe apicomplexan infection in the oyster Ostrea chilensis: a predisposing factor in

bonamiosis. Diseases of Aquatic Organisms 51: 49–60.

Hine PM, Wesney B (1997). Virus-like particles associated with cytopathology in the digestive gland

epithelium of scallops Pecten novaezelandiae and toheroa Paphies ventricosum. Diseases of Aquatic


Hine PM, MacDiarmid SC (1997). Contamination of fish products: risks and prevention. Rev. Sci. Tech.

Off. Int. Epiz. 16: 135-145.

http://www.biosecurity.govt.nz/files/pests/paua-avg-virus/avg-virus-review-julie-hills-nov07.pdfn

_________________________________________________________________________________________________________________________________________________________________________________


250

www.digsfish.com

Hine PM, Thorne T (1998). Haplosporidium sp. (Haplosporidia) in hatchery-reared pearl oysters, Pinctada

maxima (Jameson, 1901), in north Western Australia. Journal of Invertebrate Pathology 71: 48–52.

Hine PM, Thorne T (2000). A survey of some parasites and diseases of several species of bivalve mollusc in

northern Western Australia. Diseases of Aquatic Organisms 40: 67-78.

Hine PM, Thorne T (2002). Haplosporidium sp. (Alveolata: Haplosporidia) associated with mortalities

among rock oysters Saccostrea cuccullata in north Western Australia. Diseases of Aquatic Organisms 51:

123–133.

Hine PM, Diggles BK (2002a). Prokaryote infections in the New Zealand scallops Pecten novaezelandiae

and Chlamys delicatula. Diseases of Aquatic Organisms 50: 137-144.

Hine PM, Diggles BK (2002b). The distribution of Perkinsus olseni in New Zealand bivalve molluscs.

Surveillance 29: 8-11.

Hine PM, Wesney B, Besant P (1998). Replication of a herpes-like virus in larvae of the flat oyster Tiostrea

chilensis at ambient temperatures. Diseases of Aquatic Organisms 32: 161-171.

Hine PM, Cochennec-Laureau N, Berthe FCJ (2001). Bonamia exitiosus n. sp. (Haplosporidia) infecting flat

oysters Ostrea chilensis (Philippi) in New Zealand. Diseases of Aquatic Organisms 47: 63-72.

Hine PM, Wakefield S, Diggles BK, Webb VL, Maas EW (2002a). The ultrastructure of a haplosporidian

containing Rickettsiae, associated with mortalities among cultured paua Haliotis iris. Diseases of Aquatic


Hine PM, Diggles BK, Parsons M, Pringle A, Bull B (2002b). The effects of stressors on the dynamics of

Bonamia exitiosus Hine, Cochennec-Laureau & Berthe, infections in flat oysters Ostrea chilensis (Philippi).


Hine PM, Carnegie RB, Burreson EM, Engelsma MY (2009). Inter-relationships of haplosporidians deduced

from ultrastructural studies. Diseases of Aquatic Organisms 83: 247-256.

Hsieh, CY, Chuang PC, Chen LC, Tu C, Chien MS, Huang KC, Kao HF, Tung MC, Tsai SS (2006).

Infectious hypodermal and haematopoietic necrosis virus (IHHNV) infections in giant freshwater prawn

Macrbrachium rosenbergii. Aquaculture 258: 73-79.

Ho JS, Kim IH, Cruz-Lacierda ER, Nagasawa K (2004). Sea lice (Copepoda: Caligidae) parasitic on marine

cultured and wild fishes of the Philippines. Journal of the Fisheries Society of Taiwan 31: 235-249.

Hochberg FG (1983). The parasites of cephalopods: a review. Memoirs of the National Museum Victoria

No. 44: 109-145.

_________________________________________________________________________________________________________________________________________________________________________________


251

www.digsfish.com

Hoeg JT (1990). Akentrogonid host invasion and an entirely new type of life cycle in the rhizocephalan

parasite Clistosaccus paguri (Thecostraca: Cirripedia). Journal of Crustacean Biology 10: 37-52.

Hoese HD (1964). Studies on oyster scavengers and their relation to the fungus Dermocystidium marinum.

Proceedings of the National Shellfisheries Association 53: 161-174.

Holland JW, Okamura B, Hartikainen H, Secombes CJ (2010). A novel minicollagen gene links cnidarians

and myxozoans. Proceedings of the Royal Society, Biological Sciences B 278: 546-553.

Hooper C, Hardy-Smith P, Handlinger J (2007). Ganglioneuritis causing high mortalities in farmed

Australian abalone (Haliotis laevigata and Haliotis rubra). Australian Veterinary Journal 85: 188-193.

Horvat P (2010). A king of an idea turns waste into treasure. FRDC FISH 18(1): 26.

Horwitz P (1990). The translocation of freshwater crayfish in Australia: Potential impact, the need for

control and global relevance. Biological Conservation 54: 291-305.

House ML, Bartholomew JL, Winton JR, Fryer JL (1999). Relative virulence of three isolates of

Piscirickettsia salmonis for coho salmon Oncorhynchus kisutch. Diseases of Aquatic Organisms 35: 107-

113.

Howard AE (1994). The possibility of long distance transmission of Bonamia by fouling on boat hulls.

Bulletin of the European Association of Fish Pathologists 14: 211-212.

Hudson DA, Lester RJG (1994). Parasites and symbionts of wild mud crabs Scylla serrata (Forskal) of

potential significance in aquaculture. Aquaculture 120: 183–199.

Hudson DA, Shields JD (1994). Hematodinium australis n. sp., a parasitic dinoflagellate of the sand crab

Portunus pelagicus from Moreton Bay, Australia. Diseases of Aquatic Organisms 19: 109–119.

Hudson DA, Adlard RD (1994). PCR-techniques applied to Hematodinium spp and Hematodinium-like

dinoflagellates in decapod crustaceans. Diseases of Aquatic Organisms 20: 203–206.

Hudson DA, Adlard RD (1996). Nucleotide sequence determination of the partial SSU rDNA gene and ITS1

region of Hematodinium cf. perezi and Hematodinium-like dinoflagellates. Diseases of Aquatic Organisms

24: 55–60.

Hudson DA, Hudson NB, Pyecroft SB (2001). Mortalities of Penaeus japonicus prawns associated with

microsporidian infection. Australian Veterinary Journal 79: 504-505.

Humphrey JD (1995a). Australian quarantine policies and practices for aquatic animals and their products: A

review for the Scientific Working Party on Aquatic Animal Quarantine, Part 1: Review and Annexes, Bureau

of Resource Sciences, Canberra.

http://www.ingentaconnect.com/content/bpl/avj

_________________________________________________________________________________________________________________________________________________________________________________


252

www.digsfish.com

Humphrey JD (1995b). Australian quarantine policies and practices for aquatic animals and their products: A

review for the Scientific Working Party on Aquatic Animal Quarantine, Part 2: Appendices. Bureau of

Resource Sciences, Canberra.

Humphery JD, Ashburner LD (1993). Spread of the bacterial fish pathogen Aeromonas salmonicida after

importation of infected goldfish, Carassius auratus, into Australia. Australian Veterinary Journal 70: 453–

454.

Humphrey JD, Pearce M (2004). Fishnote; Epizootic Ulcerative Syndrome (Red-spot Disease). NT

Fisheries Fishnote No. 1. Oct 2004. 5 pgs

Humphrey JD, Lancaster CE, Gudkovs N, Copland JW (1987). The disease status of Australian salmonids:

bacteria and bacterial diseases. Journal of Fish Diseases 10: 403-410.

Hung LV, Tuan LA (2009). Lobster seacage culture in Vietnam. In: Williams K.C. (ed.) 2009. Spiny

lobster aquaculture in the Asia–Pacific region. Proceedings of an international symposium held at Nha

Trang, Vietnam, 9–10 December 2008. ACIAR Proceedings No. 132. Australian Centre for International

Agricultural Research: Canberra. 162 pgs. .

http://www.aciar.gov.au/system/files/node/11536/ACIAR_PR132(online).pdf

Hunter VA, Knittel MD, Fryer JL (1980). Stress induced transmission of Yersinia ruckeri infection from

carriers to recipient steelhead trout Salmo gairdneri Richardson. Journal of Fish Diseases 3: 467-472.

Hutson KS, Ernst I, Whittington ID (2007). Risk assessment for metazoan parasites of yellowtail kingfish

Seriola lalandi (Perciformes: Carangidae) in South Australian sea-cage aquaculture. Aquaculture 271: 85-

99.

Hyatt AD, Hine PM, Jones JB, Whittington RJ, Kearns C, Wise TG, Crane MS, Williams LM (1997).

Epizootic mortality in the pilchard Sardinops sagax neopilchardus in Australia and New Zealand in 1995. 2.

Identification of a herpesvirus within the gill epithelium. Diseases of Aquatic Organisms 28: 17–29.

ICUN (1987). Position Statement on the Translocation of Living Organisms.

http://intranet.iucn.org/webfiles/doc/SSC/SSCwebsite/Policy_statements/IUCN_Position_Statement_on_Tra

nslocation_of_Living_Organisms.pdf

Idowu AB, Edema MO, Adeyi AO (2006). Distribution of bacteria and fungi in the earthworm Libyodrillus

violaceous (Annelida: Oligochaeta), a native earthworm from Nigeria. Rev. Biol. Trop 54: 49-58.

http://www.scielo.sa.cr/pdf/rbt/v54n1/3355.pdf

Inouye K, Ikeya F, Yamazaki T, Hara T (1990). Studies on the methods for evaluating the virucidal activities

of germicides against IPNV and IHNV. Fish Pathology 25: 69–79.

http://www.aciar.gov.au/system/files/node/11536/ACIAR_PR132(online).pdf

http://intranet.iucn.org/webfiles/doc/SSC/SSCwebsite/Policy_statements/IUCN_Position_Statement_on_Translocation_of_Living_Organisms.pdf

http://www.scielo.sa.cr/pdf/rbt/v54n1/3355.pdf

_________________________________________________________________________________________________________________________________________________________________________________


253

www.digsfish.com

Isshiki T, Kusuda R (1987). Susceptibility of different marine fish fingerlings to yellowtail ascites virus

(YAV). Fish Pathology 22: 191-194.

Isshiki T, Nagano T, Suzuki S (2001). Infectivity of aquabirnavirus strains to various marine fish species.

Diseases of Aquatic Organisms 46: 109-114

Itoh N, Oda T, Ogawa K, Wakabayashi H (2002). Identification and development of a paramyxean ovarian

parasite in the Pacific oyster Crassostrea gigas. Fish Pathology 37: 23-28.

Jancovich JK, Davidson EW, Parameswaran N, Mao J, Chinchars VG, Collins JP, Jacobs BL, Storfer A

(2005). Evidence for emergence of an amphibian iridoviral disease because of human-enhanced spread.

Molecular Ecology 16: 213-224

Johnson PT, Lightner DV (1988). Rod-shaped nuclear viruses of crustaceans: gut infecting species.


Johnston T, Hitchcock L (1923). A bacterial disease destructive to fish in Queensland rivers. Transactions

of the Royal Society of South Australia 48: 157-161.

Johnston T, Deland EW (1929). Australian acanthocephala, No. 1. Census of recorded hosts and parasites.

Transactions of the Royal Society of South Australia 53: 146-154.

Johnston T, Simpson ER (1942). Some nematodes from Australian frogs. Transactions of the Royal Society

of South Australia 66: 172-179.

Jones JB, Gibson AP (1997). Risk analysis for the practice of importing frozen fish as bait. An assessment

by the Western Australian Fishing Industry Council of the diseae risks associated with imports into Australia

of wild caught frozen fish for bait usage by the rock lobster industry. October 1997. WAFIC. 188 pgs.

Jones JB, Lawrence CS (2001) Diseases of yabbies (Cherax albidus) in Western Australia. Aquaculture 194:

221-232.

Jones JB, Creeper J (2006). Diseases of pearl oysters and other molluscs: a Western Australian perspective.


Jones JB, Stephens F (2006). Aquatic Animal Health Subprogram: development of a national translocation

policy using abalone and prawns as templates for other aquatic species. FRDC Project 2004/080. 86 pgs.

Jones JB, Scotti PD, Dearing SC, Wesney B (1996). Virus-like particles associated with marine mussel

mortalities in New Zealand. Diseases of Aquatic Organisms 25: 143-149.

_________________________________________________________________________________________________________________________________________________________________________________


254

www.digsfish.com

Jones TC, Lester RJG (1993). Aspects of the biology and pathogenicity of Diceratocephala boschmai

(Platyhelminthes; Temnocephala), an ectosymbiont on the redclaw crayfish Cherax quadricarinatus.

Australian Journal of Marine and Freswater Research 44: 927-933.

Jung SJ, Miyazaki T (1995). Herpesviral haematopoietic necrosis of goldfish (Carassius auratus). Journal of

Fish Diseases 18: 211-220.

Jung SJ, Kitamura SI, Song JY, Oh MJ (2007). Miamiensis avidus (Ciliophora: Scuticociliatida) causes

systematic infection of olive flounder Paralichthys olivaceus and is a senior synonym of Philasterides

dicentrarchi. Diseases of Aquatic Organisms 73: 227–234.

Kabata Z (1984). Diseases caused by metazoans: Crustaceans. In: Kinne O (ed). Diseases of Marine

Animals, Vol. IV, Part 1, Introduction, Pisces. Biologische Anstalt Helgoland, Hamburg. Pgs. 321- 399.

Kailola PJ, Williams MJ, Stewart PC, Reichelt RE, McNee A, Grieve C (1993). Australian Fisheries

Resources. Bureau of Resource Sciences and the Fisheries Research and Development Corporation. 422 pgs.

Kamaishi T, Yoshinaga T (2002). Detection of Haplosporidium nelsoni in Pacific oyster Crassostrea gigas

in Japan. Fish Pathology 37: 193-195.

Kearn GC (1986). The eggs of monogeneans. Advances in Parasitology 25: 175-273.

Kennedy CR (1993). Introductions, spread and colonization of new localities by fish, helminth and

crustacean parasites in the British Isles: a perspective and appraisal. Journal of Fish Biology 43: 287-301.

Kent ML, Speare DJ (2005). Review of the sequential development of Loma salmonae (Microsporidia)

based on experimental infections of rainbow trout (Oncorhynchus mykiss) and Chinook salmon (O.

tshawytscha). Folia Parasitologica 52: 63-68.

Kent ML, Moser M, Fournei JW (1989). Coccidian parasites (Apicomplexa: Eucoccidorida) in hardy head

fish, Atherinomorus capricornensis (Woodland). Journal of Fish Diseases 12: 179-183.

Kent ML, Margolis L, Corliss JO (1994). The demise of a class of protests: taxonomic and nomenclatural

revisions proposed for the protist phylum Myxozoa Grasse, 1970. Canadian Journal of Zoology 72: 932-

937.

Kent ML, Dawe SC, Speare DJ (1995). Transmission of Loma salmonae (Microsporea) to chinook salmon in

sea water. Canadian Veterinary Journal 36: 98-101.

Kent ML, and 15 other co-authors (2001). Recent advances in our knowledge of the myxozoa. Journal of

Eukaryotic Microbiology 48: 395-413.

_________________________________________________________________________________________________________________________________________________________________________________


255

www.digsfish.com

Kewagama Research (2002). National survey of bait and berley use by recreational fishers. Report to

Biosecurity Australia, AFFA. December 2002. Kewagama Holdings, Pty. Ltd., Noosaville, Queensland,

Australia. 137 pgs.

Kewagama Research (2007). National survey of bait and berley use by recreational fishers: a follow-up

survey focusing on prawns/shrimp. Report to Biosecurity Australia. January 2007. Kewagama Holdings

Pty. Ltd., Noosaville, Queensland, Australia. 194 pgs.

Kilpatrick AM, Briggs CJ, Daszak P (2010). The ecology and impact of chytridiomycosis: an emerging

disease of amphibians. Trends in Ecology and Evolution 25: 109-118.

Kim JH, Hayward CJ, Joh SJ, Heo GJ (2002a). Parasitic infections in live freshwater tropical fishes imported

to Korea. Diseases of Aquatic Organisms 52: 169-173.

Kim JH, Hayward CJ, Heo GJ (2002b). Nematode worm infections (Camallanus cotti, Camallanidae) in

guppies (Poecilia reticulata) imported to Korea. Aquaculture 205: 231-235.

Kim R, Faisal M (2010). Experimental studies confirm the wide host range of the Great Lakes viral

haemorrhagic septicaemia virus genotype IVb. Journal of Fish Biology 33: 83-88.

Kiryu Y, Shields JD, Vogelbein WK, Kator H, Blazer VS (2003). Infectivity and pathogenicity of the

oomycete Aphanomyces invadans in Atlantic menhaden Brevoortia tyrannus. Diseases of Aquatic


Kiryu Y, Blazer VS, Vogelbein WK, Kator H, Shields JD (2005). Factors influencing the sporulation and

cyst formation of Aphanomyces invadans, etiological agent of ulcerative mycosis in Atlantic menhaden

Brevoortia tyrannus. Mycologica 97: 569-575.

Kitamura SI, Ko JY, Lee WL, Kim SR, Song JY, Kim DK, Jung SJ, Oh MJ (2007). Seasonal prevalence of

lymphocystis disease virus and aquabirnavirus in Japanese flounder, Paralichthys olivaceus and blue mussel,

Mytilus galloprovincialis. Aquaculture 266: 26-31.

Kitancharoen N, Nakamura K, Wada S, Hatai K (1994). Atkinsiella awabi sp. nov. isolated from stocked

abalone, Haliotis sieboldii. Mycoscience 35: 265-270.

Kleeman SN, Adlard RD, Lester RJG (2002a). Detection of the initial infective stages of the protozoan

parasite Marteilia sydneyi in Saccostrea glomerata and their development through to sporogenesis.

International Journal for Parasitology 32: 767-784.

Kleeman SN, Le Roux F, Berthe F, Adlard RD (2002b). Specificity of PCR and in situ hybridization assays

designed for detection of Marteilia sydneyi and M. refringens. Parasitology 125: 131-141.

_________________________________________________________________________________________________________________________________________________________________________________


256

www.digsfish.com

Korotaeva VD (1975). (Contribution to the fauna of helminths and parasitic crustaceans of marine

commercial fishes of the Australian and New Zealand waters). Trudy Academii Nauk SSR (Novaja Serija)

26: 46-60 (in Russian).

Korsnes K, Karlsbakk E, Devold M, Nerland AH, Nylund A (2009). Tissue tropism of nervous necrosis

virus (NNV) in Atlantic cod, Gadus morhua L., after intraperitoneal challenge with a virus isolate from

diseased Atlantic halibut, Hippoglossus hippoglossus (L.). Journal of Fish Diseases 32: 655-665.

Koudela B, Kucerova S, Hudcovic T (1999). Effect of low and high temperatures on infectivity of

Encephalitozoon cuniculi spores suspended in water. Folia Parasitologica 46: 171-174.

Kovacs-Gayer E, Csaba G, Ratz F, Bekesi L, Szakolczai J (1986). Granulomatosis of common carp

(Cyprinus carpio L.). Bulletin of the European Association of Fish Pathologists 6: 72-75.

Krabsetsve K, Cullen BR, Owens L (2004). Rediscovery of the Australian strain of infectious hypodermal

and haematopoietic necrosis virus. Diseases of Aquatic Organisms 61: 153-158.

Krol RM, Hawkins WE, Overstreet RM (1991). Rickettsial and mollicute infections in hepatopancreatic

cells of cultured pacific white shrimp (Penaeus vannamei). Journal of Invertebrate Pathology 57: 362-370.

Kusuda R, Salati F (1993). Major bacterial diseases affecting mariculture in Japan. Annual Review of Fish

Diseases 1993: 69-85.

Kusuda R, Salati F (1999). Enterococcus seriolicida and Streptococcus iniae. In: Woo PTK, Bruno DW

(eds.) Fish Diseases and Disorders, Volume 3: Viral, Bacterial and Fungal Infections, pgs. 303-317.

Kvingedal R, Owens L, Jerry DR (2006). A new parasite that infects eggs of the mud crab Scylla serrata, in

Australia. Journal of Invertebrate Pathology 93: 54-59.

LaFauce KA, Owens L (2007). Investigation of the pathogenicity of Penaeus merguiensis densovirus

(PmergDNV) to juvenile Cherax quadricarinatus. Aquaculture 271: 31-38.

LaFauce KA, Owens L (2008). The use of insects as a bioassay for Penaeus merguiensis densovirus

(PmergDNV). Journal of Invertebrate Pathology 98: 1-6.

LaFauce KA, Elliman J, Owens L (2007a). Molecular characterisation of hepatopancreatic parvovirus

(PmergDNV) from Australian Penaeus merguiensis. Virology 362: 397-403.

La Fauce KA, Layton R, Owens L (2007b). TaqMan real-time PCR for detection of hepatopancreatic

parvovirus from Australia. Journal of Virological Methods 140: 10–16.

Landsberg JH, Paperna I (1992). Systemic granuloma in goldfish caused by a Dermocystidium-like

aetiological agent. Diseases of Aquatic Organisms 13: 75-78.

_________________________________________________________________________________________________________________________________________________________________________________


257

www.digsfish.com

Langdon JS (1988). Diseases of introduced Australian fish. In: Fish Diseases, Post- Graduate Committee in

Veterinary Science, Sydney, pp. 225–276.

Langdon JS (1989a). Disease risks of fish introductions and translocations. Workshop on Introduced and

Translocated Fishes and their Ecological Effects Australia Society for Fish Biology, Townsville, August

1989.

Langdon JS (1989b). Experimental transmission and pathogenicity of epizootic haematopoietic necrosis

virus (EHNV) in redfin perch, Perca fluviatilis L., and 11 other teleost species. Journal of Fish Diseases 12:

295-310.

Langdon JS (1990). Disease risks of fish introductions and translocations. Bureau of Rural Resources

Proceedings 8: 98–107.

Langdon JS (1991a). Myoliquefaction post-mortem ('milky flesh') due to Kudoa thyrsites (Gilchrist)

(Myxosporea: Multivalvulida) in mahi mahi, Coryphaena hippurus L. Journal of Fish Diseases 14: 45-54.

Langdon JS (1991b). Description of Vavraia parastacida sp. nov. (Microspora: Pleistophoridae) from

marron, Cherax tenuimanus Smith (Decapoda: Parastacidae). Journal of Fish Diseases 14: 619–629.

Langdon JS, Humphrey JD (1987). Epizootic haematopoietic necrosis, a new viral disease in redfin perch,

Perca fluviatilis L. Journal of Fish Diseases 10: 289-297.

Langdon JS, Thorne T (1992). Experimental transmission per-os of microsporidosis due to Vavraia

parastacida in the marron, Cherax tenuimanus (Smith) and yabby Cherax albidus Clark. Journal of Fish

Diseases 15: 315-322.

Langdon JS, Humphrey JD, Williams LM (1986). First virus isolation from Australian fish: an iridovirus-

like pathogen from redfin perch, Perca fluviatilis L. Journal of Fish Diseases 9: 263-268.

Langdon JS, Humphrey JD, Williams LM (1988). Outbreaks of an EHNV-like iridovirus in cultured rainbow

trout, Salmo gairdneri Richardson, in Australia. Journal of Fish Diseases 11: 93–96.

Langdon JS, Thorne T, Fletcher WJ (1992). Reservoir hosts and new clupeoid host records for the

myoliquefactive myxosporean parasite Kudoa thyrsites (Gilchrist). Journal of Fish Diseases 15: 459-471.

Lannan CN, Fryer JL (1994). Extracellular survival of Piscirickettsia salmonis. Journal of Fish Diseases 17:

545–548.

Laoaroon S, Boonnat A, Poltana P, Kanchanaphum P, Gangnonngiw W, Nash G, Withayachumnankul B

(2005) Infectivity of white spot syndrome virus (WSSV) to the polychaete Perenereis nuntia and a

possibility of WSSV transmission from the polychaete to the black tiger shrimp Penaeus monodon. In:

_________________________________________________________________________________________________________________________________________________________________________________


258

www.digsfish.com

Walker PJ, Lester R, Bondad-Reantaso MB (eds) Diseases in Asian Aquaculture V. Fish Health Section,

Asian Fisheries Soc., Manila, p 353-361

Larsson JIR, Koie M (2005). Ultrastructual study and description of Paramyxoides nephtys gen. n., sp. n. a

parasite of Nephtys caeca (Fabricius, 1780) (Polychaeta: Nephtyidae). Acta Protozoology 44: 175- 187.

Leano EM (2002). Haliphthoros spp. from spawned eggs of captive mud crab, Scylla serrata, broodstocks.

Fungal Diversity 9: 93-103.

Lergessner JG (2006). Oysterers of Moreton Bay. Schuurs Publications. 235 pgs.

Lesel R, Lesel M, Gavini F, Vulllaume A (1983). Outbreak of enteric redmouth disease in rainbow trout,

Salmo gairdneri Richardson, in France. Journal of Fish Diseases 6: 385-387.

Lester RJG (1982). Unicapsula seriolae n. sp. (Myxosporea, Multivalvulida) from Australian yellowtail

kingfish Seriola lalandi. Journal of Protozoology 29: 584-587.

Lester RJG (1986). Abalone dieback caused by protozoan infection? Australian Fisheries 45: 2627.

Lester RJG (1989). Diseases of cultured molluscs in Australia. In: Advances in Tropical Aquaculture ,

TahitiFeb 20 – Mar 4 1989. AQUACOP IFREMER Actes de Colloque 9: 207-216.

Lester RJG, Davis GHG (1981). A new Perkinsus species (Apicomplexa, Perkinsea) from the abalone

Haliotis ruber. Journal of Invertebrate Pathology 37: 181187.

Lester RJG, Sewell KB (1989). Checklist of Parasites From Heron Island, Great Barrier Reef. Australian

Jounal of Zoology 37: 101-128.

Lester RJG, Hayward CJ (2005). Control of Perkinsus disease in abalone. Final Report for FRDC Project

no. 2000/151. 41 pgs.

Lester RJG, Doubrovsky A, Paynter JL, Sambhi SK, Atherton JG (1987). Light and electron microscope

evidence of baculovirus infection in the prawn Penaeus plebejus. Diseases of Aquatic Organisms 3: 217-

219.

Levine ND, Nye RR (1977). A survey of blood and other tissue parasites of leopard frogs Rana pipiens in

the United States. Journal of Wildlife Diseases. 13: 17-23.

Levsen A, Berland B (2002). Post-embryonic development of Camallanus cotti (Nematoda:Camallanidae),

with emphasis on growth of some taxonomically important somatic characters. Folia Parasitology 49: 231-

238.

_________________________________________________________________________________________________________________________________________________________________________________


259

www.digsfish.com

Levsen A, Jakobsen PJ (2002). Selection pressure towards monoxeny in Camallanus cotti (Nematoda:

Camallanidae) facing an intermediate host bottleneck situation. Parasitology 124: 625-629.

Li X, Fayer R (2006). Infectivity of microsporidian spores exposed to temperature extremes and chemical

disinfectants. Journal of Eukaryotic Microbiology 53: S77-S79.

Li YY, Xia XA, Wu QY, Liu WH, Lin YS (2008). Infection with Hematodinium sp. in mud crabs Scylla

serrata cultured in low salinity water in southern China. Diseases of Aquatic Organisms 82: 145-150.

Lightner DV (1996). Epizootiology, distribution and the impact on international trade of two penaeid shrimp

viruses in the Americas. Revue Scientifique et Technique Office International des Epizooties 15: 579-601.

Lightner DV (1999). The penaeid shrimp viruses TSV, IHHNV, WSSV, and YHV: current status in the

Americas, available diagnostic methods, and management strategies. Journal of Applied Aquaculture 9:27–

52.

Lightner DV, Redman RM (1981). A baculovirus-caused disease of the penaeid shrimp, Penaeus monodon.

Journal of Invertebrate Pathology 38: 299-302.

Lightner DV, Redman RM (1985). A parvo-like virus disease of penaeid shrimp. Journal of Invertebrate

Pathology 45: 47-53.

Lightner DV, Redman RM, Bell TA, Brock JA (1983a). Detection of IHHN virus in Penaeus stylirostris and

Penaeus vannamei imported into Hawaii. Journal of the World Mariculture Society 14: 212-225.

Lightner DV, Redman RM, Bell TA (1983b). Infectious hypodermal and hematopoietic necrosis, a newly

recognized virus disease of penaeid shrimp. Journal of Invertebate. Pathology 42: 62–70.

Lightner DV, Redman RM, Bell TA (1983c). Observations on the geographic distribution, pathogenesis and

morphology of the baculovirus from Penaeus monodon Fabricius. Aquaculture 32: 209-233.

Lightner DV, Redman RM, Williams RR, Mohney LL, Clerx JPM, Bell TA, Brock JA (1985). Recent

advances in penaeid virus disease investigations. Journal of the World Mariculture Society 16: 267-274.

Lightner DV, Hedrick RP, Fryer JL, Chen SN, Liao IC, Kou GH (1987). A survey of cultured penaeid

shrimp in Taiwan for viral and other important diseases. Fish Pathology 22: 127-140.

Lightner DV, Redman RM, Poulos BT, Nunan LM, Mari JL, Hasson KW (1997). Risk of spread of penaeid

shrimp viruses in the Americas by the international movement of live and frozen shrimp. Rev. Sci. Tech. -

Off. Int. Epizoot. 16: 146–160.

Lightner DV, Pantoja CP, Redman RM, Poulos BT, Nguyen HD, Do TH, Nguyen TC (2008). Collaboration

on milky disease of net-pen-reared spiny lobsters in Vietnam. OIE Bulletin 2008 (2): 46-47.

_________________________________________________________________________________________________________________________________________________________________________________


260

www.digsfish.com

Lightner DV, Redman RM, Pantoja CR, Navarro SA, Tang-Nelson KFJ, Noble BL, Nunan LM (2009).

Emerging non-viral infectious and noninfectious diseases of farmed penaeid shrimp and other decapods. Pgs

46-52. In: Browdy, CL, Jory DE, editors. The Rising Tide, Proceedings of the Special Session on

Sustainable Shrimp Farming, Aquaculture 2009. The World Aquaculture Society, Baton Rouge Louisiana,

USA.

Lilley JH, Roberts RJ (1997). Pathogenicity and culture studies comparing the Aphanomyces involved with

epizootic ulcerative syndrome (EUS) with other similar fungi. Journal of Fish Diseases 20: 135-144.

Lilley JH, Thompson KD, Adams A (1997). Characterization of Aphanomyces invadans by electrophoretic

and western blot analysis. Diseases of Aquatic Organisms 30: 187-197.

Limsuwan C (1991). Handbook for cultivation of black tiger prawns. Tansetakit Co. Ltd., 222 Tansetakit

Building, Wipawadee-Rangsit Road, Ladyaw, Chatujak, Bangkok, 202 pgs.

Lin CK (1989). Prawn culture in Taiwan, What went wrong? World Aquaculture 20: 19–20.

Lin CL, Ho JS, Chen SN (1996). Developmental stages of Caligus epidemicus Hewitt, a copepod parasite of

tilapia cultured in brackish water. Journal of Natural History 30: 661-684.

Lindgren CJ (2006). Angler awareness of aquatic invasive species in Manitoba. Journal of Aquatic Plant

Management 44: 103-108. http://www.apms.org/japm/vol44/v44p103.pdf

Lintermans M, Rutzou M, Kukolic K (1990). Introduced fish of the Canberra Region - recent range

expansions. Bureau of Rural Resources Proceedings 8: 50- 60.

Lintermans M (2004). Human-assisted dispersal of alien freshwater fish in Australia. New Zealand Journal

of Marine and Freshwater Research 38: 481-501.

Lintermans M (2009). Fishes of the Murray Darling Basin: An introductory guide. Murray Darling Basin

Authority Publication No. 10/07. 157 pgs. http://www.mdba.gov.au/files/publications/MDBA-Fish-species-

book.pdf

Lio-Po GD, Sanvictores ME (1986). Tolerance of Penaeus monodon eggs and larvae to fungicides against

Lagenidium sp. & Haliphthoros philippinensis. Aquaculture 51: 161- 168

Litvak MK, Mandrak NE (1993). Ecology of freshwater baitfish use in Canada and the United States.

Fisheries 18: 6-13.

Lleonart M, Handlinger J, Powell M (2003a). Treatment of spionid mud worm (Boccardia knoxi Rainer)

infestation of cultured abalone. Aquaculture 217: 1-10.

http://www.apms.org/japm/vol44/v44p103.pdf

http://www.mdba.gov.au/files/publications/MDBA-Fish-species-book.pdf

_________________________________________________________________________________________________________________________________________________________________________________


261

www.digsfish.com

Lleonart M, Handlinger J, Powell M (2003b). Spionid mudworm infestation of farmed abalone (Haliotis

spp.). Aquaculture 221: 85-96.

Lohrmann KB, Hine PM, Campalans M (2009). Ultrastructure of Bonamia sp. in Ostrea chilensis in Chile.


Lom J, Dykova I (1992). Protozoan Parasites of Fishes. Developments in Aquaculture and Fisheries

Science 26. Elsevier .315 pgs.

Lom J, Dykova I (1995). Studies on protozoan parasites of Australian fishes. Notes on coccidian parasites

with description of three new species. Systematic Parasitology 31: 147-156.

Lom J, Rohde K, Dykova I (1992). Studies of protozoan parasites of Australian fishes. I. New species of the

genera Coccomyxa Leger et Hesse, 1907, Ortholinea Shulman, 1962, and Kudoa Meglitsch, 1947 (Myxozoa,

Myxosporea). Folia Parasitologica 39: 289-306.

Lom J, Dykova I, Wang CH, Lo CF, Kou GH (2000). Ultrastructural justification for the transfer of

Pleistophora anguillarum Hoshina, 1959, to the genus Heterosporis Shubert 1969. Diseases of Aquatic


Longshaw M, Feist SW, Matthews RA, Figueras A (2001). Ultrastructural characterisation of Marteilia

species (Paramyxea) from Ostrea edulis, Mytilus edulis and Mytilus galloprovincialis in Europe. Diseases of


Lopez-Flores I, Garrido-Ramos MA, de la Herran R, Ruiz-Rejon C, Ruiz-Rejo´M, Navas JI (2008).

Identification of Marteilia refringens infecting the razor clam Solen marginatus by PCR and insitu

hybridization. Molecular and Cellular Probes 22: 151–155.

Lotz JM (1997). Special topic review: viruses, biosecurity and specific pathogen-free stocks in shrimp

aquaculture. World Journal for Microbiology and Biotechnology 13: 405–413.

Lowers JM, Bartholomew JL (2003). Detection of myxozoan parasites in oligochaetes imported as food for

ornamental fish. The Journal of Parasitology 89: 84-91.

Lu Y, Tapay LM, Loh PC, Brock JA, Gose RB (1995). Distribution of yellow-head virus in selected tissues

and organs of penaeid shrimp Penaeus vannamei. Diseases of Aquatic Organisms 23: 67–70.

Ludwig HR, Leitch JA (1996). Interbasin transfer of aquatic biota via anglers bait buckets. Fisheries 21:

14-18.

Lumsden JS, Morrison B, Yason C, Russell S, Young K, Yazdanpanah A, Huber P, Al-Hussinee L, Stone D

and Way K (2007). Mortality event in freshwater drum (Aplodinotus grunniens) from Lake Ontario, Canada

associated with viral hemorrhagic septicemia virus, type IV. Diseases of Aquatic Organisms 76: 99–111.

_________________________________________________________________________________________________________________________________________________________________________________


262

www.digsfish.com

Lymbery AJ, Hassan M, Morgan DL, Beatty SJ, Doupe RG (2010). Parasites of native and exotic freshwater

fishes in south western Australia. Journal of Fish Biology 76: 1770-1785.

Ma H, Overstreet RM, Jovonovich JA (2008). Stable yellowhead virus (YHV) RNA detection by qRT-PCR

during 6 day storage. Aquaculture 278: 10-13.

Ma H, Overstreet RM, Jovonovich JA (2009). Daggerblade grass shrimp (Palaemonetes pugio): a reservoir

host for yellow-head virus (YHV). Journal of Invertebrate Pathology. 101: 112-118.

Manisseri MK, Spann KM, Lester RJG (1999). Iodine as a disinfectant against Monodon Baculovirus

(MBV). Asian Fisheries Science 12: 105-111.

Manivannan S, Kennedy B, Karunasagar I, Karunasagar I (2004). Prevalence of monodon baculovirus in

wild Metapenaeus species along the southwest coast of India. Aquaculture 232: 63-67.

Mayfield S, McGarvey R, Gorfine HK, Peeters H, Burch P, Sharma S (2011). Survey estimates of fishable

biomass following a mass mortality in an Australian molluscan fishery. Journal of Fish Diseases 34: 287-

302.

McAllister PE (1993). Salmonidfish viruses. In: Stoskopf MK (ed). Fish Medicine. W B Saunders Co.,

Philadelphia. pgs. 380-408.

McAllister PE, Owens WJ (1995). Assessment of the virulence of fish and molluscan isolates of infectious

pancreatic necrosis virus for salmonid fish by challenge of brook trout, Salvelinus fontinalis (Mitchill).


McAllister PE, Bebak J, Wagner BA (2000). Susceptibility of arctic char to experimental challenge with

infectious hematopoietic necrosis virus (IHNV) and infectious pancreatic necrosis virus (IPNV). Journal of


McColl KA, Slater J, Jeyasekaran G, Hyatt AD, Crane MSJ (2004). Detection of White Spot Syndrome virus

and Yellowhead virus in prawns imported into Australia. Australian Veterinary Journal 82: 69-74.

McColl K, Corbeil S, Williams L, Lancaster M, Crane M. 2007. Abalone ganglioneuritis – a transmission

trial, and time-course study. Proceedings of the Third National FRDC Aquatic Animal Health Scientific

Conference, Cairns, 25-27 July 2007.

McColl KA, Davies KR, Young JG, Crane MS (2009). Aquatic birnavirus infections of finfish. Australian

and New Zealand Standard Diagnostic Procedure. .

http://www.scahls.org.au/__data/assets/pdf_file/0009/1516491/Aquatic_birnaviruses.pdf

McCulloch M, Fallon S, Wyndham T, Hendy E, Lough J, Barnes D (2003). Coral record of increased

sediment flux to the inner Great Barrier Reef since European settlement. Nature 421: 727–730.

http://www.scahls.org.au/__data/assets/pdf_file/0009/1516491/Aquatic_birnaviruses.pdf

_________________________________________________________________________________________________________________________________________________________________________________


263

www.digsfish.com

McKenzie RA, Hall WTK (1976). Dermal ulceration of mullet (Mugil cephalus). Australian Veterinary

Journal 52: 230-231.

MacKenzie K, Longshaw M, Begg GS, McVicar AH (1998). Sea lice (Copepoda: Caligidae) on wild sea

trout (Salmo trutta) in Scotland. ICES Journal of Marine Science 55: 151-162.

Mao JH, Hedrick RP, Chinchar VG (1997). Molecular characterization, sequence analysis, and taxonomic

position of newly isolated fish iridoviruses. Virology 229: 212–220.

Mao J, Green DE, Fellers G, Chinchar VG (1999a). Molecular characterization of iridoviruses isolated from

sympatric amphibians and fish. Virus Research 63: 45–52.

Mao J, Wang J, Chinchar GD, Chinchar VG (1999b). Molecular characterization of a ranavirus isolated

from largemouth bass Micropterus salmoides. Diseases of Aquatic Organisms 37: 107–114.

Manjanaik B, Umesha KR, Karunasagar I, Karunasagar I (2005). Detection of hepatopancreatic parvovirus

(HPV) in wild shrimp from India by nested polymerase chain reaction (PCR). Diseases of Aquatic


Mann KH, Tyler MJ (1963). Leeches as endoparasites of frogs. Nature 197: 1224-1225.

Marcquenski S, Dixon P, Le Deuff RM, Sheppard A, Lasee B, Marcino J (2003). Spring viremia of carp

virus in the upper Midwest. Proceedings of the 28th Annual Eastern Fish Health Workshop. National Fish

Health Research Laboratory, Kearneysville, West Virginia.

Mari J, Bonami JR (1988). PC 84, a parvo-like virus from the crab Carcinus mediterraneus: pathological

aspects, ultrastructure of the agent, and first biochemical characterization. Journal of Invertebrate Pathology

51: 145-156.

Marina H, Beatty SJ, Morgan DL, Doupe RG, Lymbery AJ (2008). An introduced parasite, Lernaea

cyprinacea L., found on native freshwater fishes in the south west of Western Australia. Journal of the

Royal Society of Western Australia 91: 149-153.

Markiw ME, Wolf K (1983). Myxosoma cerebralis (myxozoa: Myxosporea) etiologic agent of salmonids

whirling disease requires a tubificid worm (Annelida: Oligochaeta) in its lifecycle. Journal of Protozoology

30: 561-564.

Matthews D, Cribb TH (1998). Digenetic trematodes of the genus Clinostomum Leidy, 1856 (Digenea:

Clinostomidae) from birds of Queensland, Australia, including C. wilsoni n. sp. from Egretta intermedia.

Systematic Parasitology 39: 199-208.

Matthews RA (2005). Ichthyophthirius multifiliis Fouquet and Ichthyophthiriosis in freshwater teleosts.

Advances in Parasitology 59: 159-241.

_________________________________________________________________________________________________________________________________________________________________________________


264

www.digsfish.com

Mauel MJ, Miller DL (2002). Piscirickettsiosis and piskirickettsiosis-like infections in fish: a review.

Veterinary Microbiology 87: 279-289.

Messick GA (1994). Hematodinium perezi infections in adult and juvenile blue crabs Callinectes sapidus

from coastal bays of Maryland and Virginia, USA. Diseases of Aquatic Organisms 19: 77–82.

Meyers TR, Koeneman TM, Bothelho C, Short S (1987). Bitter Crab Disease: a fatal dinoflagellate infection

and marketing problem for Alaskan Tanner crabs Chionoecetes bairdii. Diseases of Aquatic Organisms 3:

195–216.

Meyers TR, Botelho C, Koeneman TM, Short S, Imamura K (1990). Distribution of bitter crab

dinoflagellate syndrome in southeast Alaskan tanner crabs, Chionoecetes bairdi. Diseases of Aquatic

Organisms 9: 37–43.

Meyers TR, Morado JF, Sparks AK, Bishop GH, Pearson T, Urban D, Jackson D (1996). Distribution of

bitter crab syndrome in tanner crabs (Chionoecetes bairdi, C. opilio) from the Gulf of Alaska and the Bering

Sea. Diseases of Aquatic Organisms 26: 221–227.

Michel C, Faivre B, de Kinkelin P (1986). A clinical case of enteric redmouth in minnows (Pimephales

promelas) imported in Europe as baitfish. Bulletin of the European Association of Fish Pathologists 6: 97-

99.

Michigan Department of Natural Resources (2009). Fish Disease Control Order FO245.09.

http://www.michigan.gov/documents/dnr/FO-425_199353_7.pdf

Miller PE (2009). Diagnosis, prevalence and prevention of the spread of the parasite Heterosporis sp.

(Microsporidia: Pleistophoridae) in yellow perch (Perca flavescens) and other freshwater fish in northern

Minnesota, Wisconsin and in Lake Ontario. Thesis Submitted in Partial Fulfillment of the Requirements for

the Degree of Master of Science in Biology. University of Wisconsin. August, 2009, 57 pgs.

Mitchell AJ, Overstreet RM, Goodwin AE, Brandt TM (2005). Spread of an exotic fish-gill trematode: A

far reacing and complex problem. Fisheries 30: 11-16.

Molnar K, Rohde K (1988a). Seven new coccidian species from marine fishes in Australia. Systematic


Molnar K, Rohde K (1988b). New coccidians from freshwater fishes of Australia. Journal of Fish Diseases

11: 161-169.

Montgomery-Brock D, Tacon AG, Poulos B, Lightner DV (2007). Reduced replication of infectious

hypodermal and hematopoietic necrosis virus (IHHNV) in Litopenaeus vannamei held in warm water.


http://www.michigan.gov/documents/dnr/FO-425_199353_7.pdf

_________________________________________________________________________________________________________________________________________________________________________________


265

www.digsfish.com

Moodie EG, Le Jambre LF, Katz ME (2003a). Thelohania montirivulorum sp. nov. (Microspora:

Thelohaniidae), a parasite of the Australian freshwater crayfish, Cherax destructor (Decapoda: Parastacidae):

fine ultrastructure, molecular characteristics and phylogenetic relationships. Parasitology Research 91: 215

-228.

Moodie EG, Le Jambre LF, Katz ME (2003b). Ultrastructural characteristics and small subunit ribosomal

DNA sequence of Vairimorpha cheracis sp. nov., (Microspora: Burenellidae), a parasite of the Australian

yabby, Cherax destructor (Decapoda: Parastacidae). Journal of Invertebrate Pathology 84: 198-213.

Moodie EG, Le Jambre LF, Katz ME (2003c). Thelohania parastaci sp. nov. (Microspora: Thelohaniidae), a

parasite of the Australian freshwater crayfish, Cherax destructor (Decapoda: Parastacidae). Parasitology

Research 91: 151-165.

Moodie NJ, Horwood PF (2008). Betanodavirus infections of finfish. Australian and New Zealand Standard

Diagnostic Procedure. .

http://www.scahls.org.au/__data/assets/pdf_file/0010/1516492/Betanodavirus_Infections_of_Finfish.pdf

Moore JD, Robbins TT, Friedman CS (2000). Withering syndrome in farmed red abalone Haliotis

rufescens: thermal induction and association with a gastrointestinal rickettsiales-like prokaryote. Journal of


Morales-Covarrubias MS, Nunan LM, Lightner DV, Mota-Urbina JC, Garza-Aguirre MC, Chavez-Sanchez

MC (1999). Prevalence of infectious hypodermal and hematopoietic necrosis virus (IHHNV) in wild adult

blue shrimp Penaeus stylirostris from the Northern Gulf of California, Mexico. Journal of Aquatic Animal

Health 11: 296-301.

Moran JDW, Kent ML (1999). Kudoa thyrsites (Myxozoa: Myxosporea) infections in pen reared Atlantic

salmon in the northeast Pacific Ocean with a survey of potential non- salmonid reservoir hosts. Journal of


Moran JDW, Whitaker DJ, Kent ML (1999a). A review of the myxosporean genus Kudoa Meglitsch, 1947,

and its impact on the international aquaculture industry and commercial fisheries. Aquaculture 172: 163-

196.

Moran JDW, Whitaker DJ, Kent ML (1999b). Natural and laboratory transmission of the marine myxozoan

parasite Kudoa thyrsites to Atlantic salmon. Journal of Aquatic Animal Health 11: 110-115.

Mortensen SH, Bachere E, Gall G, Mialhe E (1992). Persistence of infectious pancreatic necrosis virus

(IPNV) in scallops Pecten maximus. Diseases of Aquatic Organisms 12: 221-227.

Mortensen SH (1993). Passage of infectious pancreatic necrosis virus (IPNV) through invertebrates in an

aquatic food chain. Diseases of Aquatic Organisms 16: 41-45.

http://www.scahls.org.au/__data/assets/pdf_file/0010/1516492/Betanodavirus_Infections_of_Finfish.pdf

_________________________________________________________________________________________________________________________________________________________________________________


266

www.digsfish.com

Mortensen SH, Nilsen RK, Hjeltnes B (1998). Stability of an infectious pancreatic necrosis virus (IPNV)

isolate stored under different laboratory conditions. Diseases of Aquatic Organisms 33: 67-71.

Motte E, Yugcha E, Luzardo J, Castro F, Leclercq G, Rodríguez J, Miranda P, Borja O, Serrano J, Terreron

M, Montalvo K, Narváez A, Tenorio N, Cedeño V, Mialhe E, Boulo V (2003). Prevention of IHHNV

vertical transmission in the white shrimp Litopenaeus vannamei. Aquaculture 219: 57–70.

Munday BL (1985). "Ulcer disease" in cod (Pseudophycis bachus) from the Tamar river. Tasmanian

Fisheries Research 27: 15-18.

Munday BL, Nakai T (1997). Nodaviruses as pathogens in larval and juvenile marine finfish. World

Journal of Microbiology and Biotechnology 13: 375-381.

Munday BL, Langdon JS, Hyatt A, Humphrey JD (1992). Mass mortality associated with a viral-induced

vacuolating encephalopathy and retinopathy of larval and juvenile barramundi, Lates calcarifer Bloch.


Munday BL,O'Donoghue PJ, Watts M, Rough KM, Hawkesford TM (1997). Fatal encephalitis due to the

scuticociliate Uronema nigricans in sea-caged, southern bluefin tuna Thunnus maccoyii. Diseases of Aquatic


Munday BL, Su XQ, Harshbarger JC (1998). A survey of product defects in Tasmanian Atlantic salmon

(Salmo salar). Aquaculture 169: 297-302.

Munday BL, Kwang J, Moody N (2002). Betanodavirus infections of teleost fish: a review. Journal of Fish

Diseases 25: 127-142.

Munro J, Owens L (2007). Yellow head-like viruses affecting the penaeid aquaculture industry: a review.

Aquaculture Research 38: 893-908.

Muraosa, Y, Morimoto K, Sano A, Nishimura K, Hatai K (2009). A new peronosporomycete, Halioticida

noduliformans gen. et sp. nov., isolated from white nodules in the abalone Haliotis spp. from Japan.

Mycoscience 50: 106-115.

Muroga K (2001). Viral and bacterial diseases of marine fish and shellfish in Japanese hatcheries.


Murphy NE, Goggin CL (2000). Genetic Discrimination of Sacculinid Parasites (Cirripedia, Rhizocephala):

Implication for Control of Introduced Green Crabs (Carcinus maenas). Journal of Crustacean Biology 20:

153-157.

Murray AG, O’Callaghan M, Jones B (2003). A model of spatially evolving herpesvirus epidemics causing

mass mortality in Australian pilchard Sardinops sagax. Diseases of Aquatic Organisms 54: 1-14.

_________________________________________________________________________________________________________________________________________________________________________________


267

www.digsfish.com

Murrell A., Kleeman SN, Barker SC, Lester RJG (2002). Synonymy of Perkinsus olseni Lester & Davis,

1981 and Perkinsus atlanticus Azevedo, 1989 and an update on the phylogenetic position of the genus

Perkinsus. Bulletin of the European Association of Fish Pathologists 22: 258–265.

NACA (2010). Quarterly Aquatic Animal Disease Report (Asia and Pacific region). January-March 2010.

FAO/Network of Aquaculture Centres in Asia-Pacific. July 2010.

Nagasawa K, Inoue A, Myat S, Umino T (2007). New host records for Lernaea cyprinacea (Copepoda), a

parasite of freshwater fishes, with a checklist of the lernaeidae in Japan (1915-2007). J. Grad. School Biosp.

Sci. Hiroshima University 46: 21-33.

Najiah M, Nadirah M, Sakri I, Shaharom-Harrison F (2010). Bacteria associated with wild mud crab (Scylla

serrata) from Setiu wetland, Malaysia, with emphasis on antibiotic resistances. Pakistan Journal of

Biological Sciences 13: 293-297.

Nakajima K, Maeno Y (1998). Pathogenicity of Red Sea Bream Iridovirus and other fish Iridoviruses to Red

Sea Bream. Fish Pathology 33: 143-144.

Nakajima K, Inouye K, Sorimachi M (1998). Viral diseases in cultured marine fish in Japan. Fish


Nakamura K, Hatai K (1995). Atkinsiella dubia and its related species. Mycoscience 36: 431-438.

Nakamura K, Nakamura Miho, Hatai K, Zafran (1995). Lagenidium infection in eggs and larvae of

mangrove crab (Scylla serrata) produced in Indonesia. Mycoscience 36: 399-444.

Nash G, Poernomo A, Nash MB (1988). Baculovirus infection in brackishwater pond cultured Penaeus

monodon Fabricius in Indonesia. Aquaculture 73: 1-6.

Natividad JM, Lightner DV (1992). Prevalence and geographic distribution of MBV and other diseases in

cultured giant tiger prawns (Penaeus monodon) in the Philippines. In: Diseases of Cultured Penaeid Shrimp

in Asia and the United States, Fulks W. & Main K., eds. The Oceanic Institute, Makapuu Point, Honolulu,

Hawaii, USA, pgs. 139-160.

Neira FJ, Lyle JM, Keane JP (2009). Shelf spawning habitat of Emmelichthys nitidus in south-eastern

Australia – Implications and suitability for egg-based biomass estimation. Estuarine Coastal and Shelf

Science 81: 521–532.

Nell JA (2001). The history of oyster farming in New South Wales. Marine Fisheries Review 63: 14-25.

Nerland AH, Skaar C, Eriksen TB, Bleie H (2007). Detection of nodavirus in seawater from rearing

facilities for Atlantic halibut Hippoglossus hippoglossus larvae. Diseases of Aquatic Organisms 73: 201-

205.

_________________________________________________________________________________________________________________________________________________________________________________


268

www.digsfish.com

Newman MW, Johnson CA (1975). A disease of blue crabs (Callinectes sapidus) caused by a parasitic

dinoflagellate, Hematodinium sp. Journal of Parasitology 63: 554–557.

New York State Department of Conservation (2009). Baitfish of New York State. How to identify them and

legally use them in our freshwaters and the Hudson River. 30 pgs.

http://www.dec.ny.gov/docs/fish_marine_pdf/baitfishofny.pdf

Nickum D (1999). Whirling disease in the United States: a summary of progress and research and

management, Trout Unlimited. 35 pgs. http://whirlingdisease.montana.edu/pdfs/TU_Report_99.pdf

Nishizawa T, Furuhashi M, Nagai T, Nakai T, Muroga K (1997). Genomic classification of fish nodaviruses

by molecular phylogenetic analysis of the coat protein gene. Applied and Environmental Microbiology 63:

1633-1636.

Noga EJ (1996). Fish disease: diagnosis and treatment. 367 pgs.

Nolan MJ, Cribb TH (2004). The lifecycle of Paracardicoloides yamagutii Martin , 1974 (Digenea:

Sanguinicolidae). Folia Parasitologica 51: 320-326.

Nollens HH, Keogh JA, Probert PK (2003). Effects of shell lesions on survival, growth, condition and

reproduction in the New Zealand blackfoot abalone Haliotis iris. Diseases of Aquatic Organisms 57: 127-

133.

Norman M, Reid A (2000). A Guide to Squid, Cuttlefish and Octopuses of Australasia, CSIRO Publishing

96 pgs.

Northern Territory Government (2010). Northern Territory Recreational Fishing Controls. A handy

reference to the regulations for recreational fishing in the Northern Territory. 32 pgs.

Norton JH, Shepherd MA, Perkins FP, Prior HC (1993). Perkinsus-like infection in farmed golden lipped pearl

oyster Pinctada maxima from the Torres Strait, Australia. Journal of Invertebrate Pathology 62: 105-106.

Novoa B, Figueras A, Puentes CF, Ledo A, Toranzo AE (1993). Characterization of a birnavirus isolated

from diseased turbot cultured in Spain. Diseases of Aquatic Organisms 15: 163-169.

NSW DPI (2010). NSW Recreational Freshwater Fishing Guide 2010. Government of New South Wales

Department of Primary Industries. 86 pgs. .

http://www.dpi.nsw.gov.au/__data/assets/pdf_file/0013/202351/freshwater-guide.pdf

Nunan LM, Poulos BT, Lightner DV (2000). Use of polymerase chain reaction for the detection of infectious

hypodermal and hematopoietic necrosis virus in penaeid shrimp. Marine Biotechnology 2: 319-328.

http://www.dec.ny.gov/docs/fish_marine_pdf/baitfishofny.pdf

http://whirlingdisease.montana.edu/pdfs/TU_Report_99.pdf

http://www.dpi.nsw.gov.au/__data/assets/pdf_file/0013/202351/freshwater-guide.pdf

_________________________________________________________________________________________________________________________________________________________________________________


269

www.digsfish.com

Nunan LM, Poulos BT, Redman M, Le Groumellec M, Lightner DV (2003a). Molecular detection methods

developed for a systemic rickettsia-like bacterium (RLB) in Penaeus monodon (Decapoda: Crustacea).


Nunan LM, Noble B, Le Groumellec M, Lightner DV (2003b). Experimental infection of Penaeus vannamei

by a rickettsia-like bacterium (RLB) originating from Penaeus monodon. Diseases of Aquatic Organisms 54:

43-48.

Nunan LM, Poulos BT, Navarro S, Redman RM, Lightner DV (2010) Milky hemolymph syndrome (MHS)

in spiny lobsters, penaeid shrimp and crabs. Diseases of Aquatic Organisms 91: 105-112.

Ogburn DM, White I, McPhee DP (2007). The disappearance of oyster reefs from eastern Australian

estuaries impact of colonial settlement or mudworm invasion? Coastal Management /35: 271-287.

Oanh DTH, van Hulten M, Cowley JA, Walker PJ (2011). Pathogenicity of gill-associated virus and

Mourilyan virus during mixed infections of black tiger shrimp (Penaeus monodon). Journal of General

Virology 92: 893–901.

O’Donoghue PJ, Adlard RD (2000). Catalogue of protozoan parasites recorded in Australia. Memoirs of the

Queensland Museum 45: 1-163.

O'Donoghue PJ, Phillips PH, Shepherd SA (1991). Perkinsus (Protozoa: Apicomplexa) infections in abalone

from South Australian waters. Transactions of the Royal Society of South Australia 115: 77-82.

Oidtmann B, Heitz E, Rogers D, Hoffman RW (2002). Transmission of crayfish plague. Diseases of


OIE (2005). Report of the meeting of the bureau of the OIE Aquatic Animal Health Standards Commission,

Paris, 11–15 October 2004 . Report of Crustacean Team. Pgs. 43-60.

www.oie.int/downld/SG/2005/A_73SG_12_CS4A.PDF

OIE (2007a). Mourilyan Virus. OIE Aquatic Animal Disease Card 2007. .

http://www.oie.int/fileadmin/Home/eng/Internationa_Standard_Setting/docs/pdf/Mourilyan_virus_card_200

7_AN.pdf

OIE (2007b). Milky Haemolymph Disease of spiny lobsters (Panulirus spp.). OIE Aquatic Animal Disease

Card 2007. .

http://www.oie.int/fileadmin/Home/eng/Internationa_Standard_Setting/docs/pdf/Microsoft_Word_-

_Milky_haemolymph_disease_of_lobsters_card__9-04-08_.pdf

OIE (2010a). Aquatic Animal Health Code. OIE Paris, France. .

http://www.oie.int/en/international-standard-setting/aquatic-code/access-online/

http://www.oie.int/downld/SG/2005/A_73SG_12_CS4A.PDF

http://www.oie.int/fileadmin/Home/eng/Internationa_Standard_Setting/docs/pdf/Mourilyan_virus_card_2007_AN.pdf

http://www.oie.int/fileadmin/Home/eng/Internationa_Standard_Setting/docs/pdf/Microsoft_Word_-_Milky_haemolymph_disease_of_lobsters_card__9-04-08_.pdf

http://www.oie.int/en/international-standard-setting/aquatic-code/access-online/

_________________________________________________________________________________________________________________________________________________________________________________


270

www.digsfish.com

OIE (2010b). Manual of Diagnostic Tests for Aquatic Animals. OIE Paris, France.

http://www.oie.int/en/international-standard-setting/aquatic-manual/access-online

OIE (2011). Infection with Bonamia exitiosa, United Kingdom. Immediate notification. .

http://web.oie.int/wahis/public.php?page=single_report&pop=1&reportid=10283

Ostland VE, Hicks BD, Daly JG (1987). Furunculosis in baitfish and its transmission to salmonids. Diseases


Overstreet RM, Whatley EC (1975). Prevention of microsporidiosis in the blue crab with notes on natural

infections. In: Avault, J.W., Miller, R. (Eds.), Proceedings of the Sixth Annual Workshop World Mariculture

Society. Louisiana State University, Louisiana, pgs. 335– 345.

Owens L (1993). Description of the first haemocytic rod-shaped virus from a penaeid prawn. Diseases of


Owens L, Glazebrook JS (1988). Microsporidosis in prawns from northern Australia. Australian Journal of

Marine and Freshwater Research 39: 301-305.

Owens L, McElnea C (2000). Natural infection of the redclaw crayfish Cherax quadricarinatus with

presumptive spawner-isolated mortality virus. Diseases of Aquatic Organisms 40: 219-223.

Owens L, DeBeer S, Smith J (1991). Lymphoidal parvovirus-like particles in Australian penaeid prawns.


Owens L, Anderson IG, Kenway M, Trott L, Benzie JAH (1992). Infectious hypodermal and haematopoietic

necrosis virus (IHHNV) in a hybrid penaeid prawn from tropical Australia. Diseases of Aquatic Organisms

14: 219-228.

Owens L, Haqshenas G, McElnea C, Coelen R (1998). Putative spawner isolated mortality virus associated

with mid-crop mortality syndrome in farmed Penaeus monodon from northern Australia. Diseases of


Owens L, McElnea C, Snape N, Harris L, Smith M (2003). Prevalence and effect of spawner-isolated

mortality virus on the hatchery phases of Penaeus monodon and P. merguiensis in Australia. Diseases of


Owens L, LaFauce K, Juntunen K, Hayakijkosol O, Zeng C (2009). Macrobrachium rosenbergii nodavirus

disease (white tail disease) in Australia. Diseases of Aquatic Organisms 85: 175-180.

Owens L, Liessmann L, LaFauce K, Nyguyen T, Zeng C (2010). Intranuclear bacilliform virus and

hepatopancreatic parvovirus (PmergDNV) in the mud crab Scylla serrata (Forskal) of Australia.


http://www.oie.int/en/international-standard-setting/aquatic-manual/access-online

http://web.oie.int/wahis/public.php?page=single_report&pop=1&reportid=10283

_________________________________________________________________________________________________________________________________________________________________________________


271

www.digsfish.com

Padros F, Palenzuela O, Hispano C, Tosas O, Zarza C, Crespo S, Alvarez-Pellitero P (2001). Myxdium leei

(Myxozoa) infections in aquarium-reared mediterranean fish species. Diseases of Aquatic Organisms 47:

57-62.

Paperna I, Lainson R (1995). Alloglugea bufonis nov. gen., nov.sp. (Microsporea: Glugeidae) a

microsporidian of Bufo marinus tadpoles and metamorphosing toads (Amphibia: Anura) from Amazonian

Brazil. Diseases of Aquatic Organisms 23: 7-16.

Paramá A, Arranz JA, Álvarez MF, Sanmartín ML, Leiro J (2006). Ultrastructure and phylogeny of

Philasterides dicentrarchi (Ciliophora, Scuticociliata) from farmed turbot in NW Spain. Parasitology 132:

555–564.

Parliament of Tasmania (1933). Salmon and Freshwater Fisheries Commission: Report for year ending 30th

June 1933. Government of Tasmania, Hobart.

Parameswaran V, Kumar S, Ahmed VP, Sahul Hameed AS (2008). A fish nodavirus associated with mass

mortality in hatchery-reared Asian Sea bass, Lates calcarifer. Aquaculture 275: 366–369.

Park KI, Park JK, Lee J, Choi KS (2005). Use of molecular markers for species identification of Korean

Perkinsus sp. isolated from Manila clams Ruditapes philippinarum. Diseases of Aquatic Organisms 66: 255-

263.

Parukuhin AM (1985). New trematode species from commercial fish of the Indian Ocean. Nauchnye Doki

Vyss Shkoly Biol Nauki 8 : 29-34.

Paterson KJ, Schreider MJ, Zimmerman KD (2003). Anthropogenic effects on seston quality and quantity

and the growth and survival of Sydney rock oyster (Saccostrea glomerata) in two estuaries in NSW,

Australia. Aquaculture 221: 407–426.

Paxton H (1979). Taxonomy and aspects of the life history of Australian beachworms (Polychaeta:

Onuphidae). Australian Journal of Marine and Freshwater Research 30: 265-294.

Paxton H (1996). Hirsutonuphis (Polychaeta: Onuphidae) from Australia, with a discussion of setal

progression in juveniles. Invertebrate Taxonomy 10: 77-96.

Paynter JL, Lightner DV and Lester RJG (1985). Prawn virus from juvenile Penaeus esculentus. In:

Proceedings of the second Australian national prawn seminar, Cleveland, QLD, Australia, 1985: 61-64.

Paynter JL., Vickers JE, Lester RJG (1992). Experimental transmission of Penaeus monodon-type

baculovirus (MBV). In: Diseases in Asian Aquaculture I, Shariff M., Subasinghe R. & Arthur J.R., eds. Fish

Health Section, Asian Fisheries Society, Manila, Philippines, pgs. 97-110.

_________________________________________________________________________________________________________________________________________________________________________________


272

www.digsfish.com

Pearce M (1990). Epizootic Ulcerative Syndrome Technical Report. Dec 1987- Sept 1989. Northern

Territory Department of Primary Industries and Fisheries Fishery Report No. 22. 82 pgs.

Peeler EJ, Thrush MA (2004). Qualitative analysis of the risk of introducing Gyrodactylus salaris into the

United Kingdom. Diseases of Aquatic Organisms 62: 103-113.

Pennuto C (1989). The bait leech Nephelopsis obscura in North Dakota: an economic assessment. N.D. Fann

Res. 47:21-23. http://faculty.buffalostate.edu/pennutcm/BaitLeech.pdf

Perera KML (1993). No evidence for seasonality in the ectoparasitic fauna of slimy mackerel, Scomber

australasicus. Australian Journal of Marine and Freshwater Research 44: 709 – 719.

Perkins FO, Wolf PH (1976). Fine structure of Marteilia sydneyi sp. n. haplosporidan pathogen of

Australian oysters. The Journal of Parasitology 62: 528538.

Pernet B, Deconinck A, Llaban A, Archie JW (2008). Evaluating risks associated with transport of the ghost

shrimp Neotrypaea californiensis as live bait. Marine Biology 153: 1127-1140.

Pestal GP, Taylor DM, Hoenig JM, Shields JD, Pickavance R (2003). Monitoring the presence of the lethal

parasite Hematodinium sp. in snow crabs from Newfoundland. Diseases of Aquatic Organisms 53: 67–75.

Peters R, Raftos DA (2003). The role of phenoloxidase suppression in QX disease outbreaks among Sydney

rock oysters (Saccostrea glomerata). Aquaculture 223: 29-39.

Phillips WJ, Cannon LRG (1978). Ecological observations on the commercial sand crab, Portunus pelagicus

(L.), and its parasite, Sacculina granifera Boschma, 1973 (Cirripedia: Rhizocephala). Journal of Fish

Diseases 1: 137–149.

Picco AM, Collins JP (2008). Amphibian commerce as a likely source of pathogen pollution. Conservation

Biology 22: 1582–1589.

Pichot Y, Comps M, Tige G, Grizel H, Rabouin MA (1980). Recherches sur Bonamia ostreae gen. n., sp. n.,

parasite nouveau de l'huitre plate Ostrea edulis L. Rev. Trav. Inst. Peches Marit. 43:131-140.

Pierce BE, Phillips PH, Jackson G (1991). Redfin virus (EHN) fish disease confirmed in South Australia.

SAFISH 15(3): 5-6.

Pike AW, Wadsworth SL (1999). Sea lice on salmonids: their biology and control. Advances in


Plumb JA, Grizzle JM, Young HE, Noyes AD, Lamprecht S (1996). An iridovirus isolated from wild

largemouth bass. Journal of Aquatic Animal Health 8: 265–270.

http://faculty.buffalostate.edu/pennutcm/BaitLeech.pdf

_________________________________________________________________________________________________________________________________________________________________________________


273

www.digsfish.com

Powell EN, Klinck JM, Ford SE, Hofmann EE, Jordan SJ (1999). Modeling the MSX parasite in eastern

oyster (Crassostrea virginica) populations. III. Regional application and the problem of transmission.


Pramod Kiran RB, Rajendran KV, Jung SJ, Oh MJ (2002). Experimental susceptibility of different life-

stages of the giant freshwater prawn, Macrobrachium rosenbergii (de Man), to white spot syndrome virus

(WSSV). Journal of Fish Diseases 25: 201-207.

Pregenzer C (1983). Survey of metazoan symbionts of Mytilus edulis (Mollusca: Pelecypoda) in southern

Australia. Australian Journal of Marine and Freshwater Research 34: 387-396.

Primavera JH, Quinitio ET (2000). Runt-deformity syndrome in cultured giant tiger prawn Penaeus

monodon. Journal of Crustacean Biology 20: 796-802.

Pullen RR, Bouska WW, Campbell SW, Paukert CP (2009). Bothriocephalus acheilognathi and other

intestinal helminths of Cyprinella lutrensis in Deep Creek, Kansas. Journal of Parasitology 95: 1224-1226.

Pyecroft S (2008). Phylogenetic, pathological and epidemiological studies of an agent causing systemic

granulomatosis of goldfish (Carassius auratus) (SGAG) in Australia. PhD Thesis, School of Molecular and

Microbial Sciences, The University of Queensland.

Qian D, Shi Z, Zhang S, Cao Z, Liu W, Li L, Xie Y, Cambournac I, Bonami JR (2003). Extra small virus-

like particles (XSV) and nodavirus associated with whitish muscle disease in the giant freshwater prawn,

Macrobrachium rosenbergii. Journal of Fish Diseases 26: 521-527.

Rai P, Pradeep B, Safeena MP, Karunasagar I, Karunasagar I (2009b). Simultaneous presence of infectious

hypodermal and haematopoietic necrosis virus (IHHNV) and Type A virus related sequence in Penaeus

monodon in India. Aquaculture 295: 168-174.

Rainer S, Hutchings P (1977). Nephtyidae (Polychaeta: Errantia) from Australia. Records of the Australian

Museum 31: 307-347.

Rajendran KV, Cowley JA, McCulloch RJ, Walker PJ (2006). A TaqMan real-time RT-PCR for quantifying

Mourilyan virus infection levels in penaeid shrimp tissues. Journal of Virological Methods 137: 265-271.

Ravi M, Nazeer Basha A, Taju G, Ram Kumar R, Sahul Hameed AS (2010). Clearance of Macrobrachium

rosenbergii nodavirus (MrNV) and extra small virus (XSV) and immunological changes in experimentally

injected Macrobrachium rosenbergii. Fish and Shellfish Immunology 28: 428-433.

Ray SM (1966). A review of the culture method for detecting Dermocystidium marinum, with suggested

modifications and precautions. Proceedings of the National Shellfisheries Association 54: 55-69.

_________________________________________________________________________________________________________________________________________________________________________________


274

www.digsfish.com

Ray SM (1996). Historical perspective on Perkinsus marinus disease of oysters in the Gulf of Mexico.


Rayyan A, Chintiroglou CC (2003). Steinhausia mytilovum in cultured mussels Mytilus galloprovincialis in

the Thermaikos Gulf (northern Aegean Sea, Greece). Diseases of Aquatic Organisms 57: 271-273.

Read GB (2010). Comparison and history of Polydora websteri and P. haswelli (Polychaeta: Spionidae) as

mud-blister worms in New Zealand shellfish. New Zealand Journal of Marine and Freshwater Research 44:

83-100.

Reddacliff GL, Quartararo N (1992). Lymphocystis in cultured snapper (Pagrus auratus) and wild kingfish

(Seriola lalandi) in Australia. Australian Veterinary Journal 69(5): 116-117.

Reece KS, Siddall ME, Burreson EM, Graves JE (1997). Phylogenetic analysis of Perkinsus based on actin

gene sequences. The Journal of Parasitology 83: 417-423.

Reece KS, Siddall ME, Stokes NA, Burreson EM (2004). Molecular phylogeny of the haplosporidia based

on two independent gene sequences. Journal of Parasitology 90: 1111-1122.

Renault T, Stokes NA, Chollet B, Cochennec N, Berthe F, Gerad A, Burreson EM (2000). Haplosporidosis

in the Pacific oyster Crassostrea gigas from the French Atlantic coast. Diseases of Aquatic Organisms 42:

207-214.

Reno PW (1999). Infectious pancreatic necrosis and associated aquatic birnaviruses. In: Woo PTK, Bruno

DW (eds.) Fish Diseases and Disorders, Volume 3: Viral, Bacterial and Fungal Infections, pgs 1-55.

Retallick RW, McCallum H, Speare R (2004). Endemic infection of the amphibian chytrid fungus in a frog

community post-decline. PLOS Biology 2: (11): e351. doi:10.1371/journal.pbio.0020351.

Rivas C, Cepeda C, Dopazoa CP, Novoa B, Noya M, Barja JL (1993). Marine environment as reservoir of

birnaviruses from poikilothermic animals. Aquaculture 115: 183-194.

Robeldo JAF, Figueras A (1995). The effects of culture- site, depth, season, and stock source on the

prevalence of Marteilia refringens in cultured mussels (Mytilus galloprovincialis) from Galicia, Spain.

Journal of Parasitology 81: 354-363.

Roberts RJ (2001). Fish Pathology. 3rd edition. W.B. Saunders. 472 pgs.

Roberts SD, Deveney M, Sierp M (2010). Biosecurity and disease status of prawn nurseries in South

Australia. SARDI Publication Number F2010/000593-1. SARDI Research Report Series No. 449 35 pgs.

Robinson M, Hayes M, Allen B, Thorne K, Jenkins E, Stafford P (2005). Monitoring the prevalence of the

parasitic dinoflagellate Haematodinium in Irish brown crab fisheries. National Development Plan

_________________________________________________________________________________________________________________________________________________________________________________


275

www.digsfish.com

Supporting measures for Sea Fisheries Development. 45 pgs .

http://www.vims.edu/research/departments/eaah/programs/crustacean/research/hematodinium/literature/pdf

%20files%202/Robinson%20et%20al.%202005%20REPORT.pdf

Rodriguez-Tovar LE, Speare DJ, Markham RJF (2011). Fish microsporidia: Immune response,

immunomodulation and vaccination. Fish and Shellfish Immunology 30: 999-1006.

Rohde K (1984). Diseases caused by metazoans: Helminths. In: Kinne O (ed). Diseases of Marine

Animals, Vol. IV, Part 1, Introduction, Pisces. Biologische Anstalt Helgoland, Hamburg. Pgs. 193-320.

Romero X, Turnbull JF, Jimenez R (2000). Ultrastructure and cytopathology of a Rickettsia-like Organism

causing systemic infection in the redclaw crayfish, Cherax quadricarinatus (Crustacea: Decapoda), in

Ecuador. Journal of Invertebrate Pathology 76: 95-104.

Ross J (Ed.) (1995). Fish Australia. The essential fishing companion. Penguin Books. 498 pgs.

Rossteuscher S, Wenker C, Jermann T, Wahli T, Oldenberg E, Schmidt-Posthaus H (2008). Severe

scuticociliate (Philasterides dicentrarchi) infection in a population of sea dragons (Phycodurus eques and

Phyllopteryx taeniolatus). Veterinary Pathology 45: 546–550.

Roubal FR (1994). Histopathology caused by Caligus epidemicus Hewitt (Copepoda: Caligidae) on captive

Acanthopagrus australis (Gunther) (Pisces: Sparidae). Journal of Fish Diseases 17: 631-640.

Roubal FR Armitage J, Rohde K (1983). Taxonomy of metazoan ectoparasites of snapper, Chrysophrys

auratus (Family Sparidae), from southern Australia, eastern Australia and New Zealand. Australian Journal

of Zoology Supplementary Series No. 94.

Roubal FR, Paynter JL, Lester RJG (1989a). Electron microscope observation of hepatopancreatic parvo-like

virus (HPV) in the penaeid prawn, Penaeus merguiensis de Man, from Australia. Journal of Fish Diseases

12: 199-201.

Roubal, FR, Masel J, Lester RGJ (1989b). Studies on Marteilia sydneyi, agent of QX disease in the Sydney

rock oyster, Saccostrea commercialis, with implications for its life cycle. Australian Journal of Marine and


Roughley TC (1926). An investigation of the cause of an oyster mortality on the Georges River, NSW,

1924-25. Proceedings of the Linnaean Society of NSW 51: 446-491.

Roughley TC (1939). A review of the scientific investigation of the fisheries of New South Wales

[Presidential address to Linnean Society]. Proceedings of the Linnean Society of New South Wales /64:

vi_xxvii.

http://www.vims.edu/research/departments/eaah/programs/crustacean/research/hematodinium/literature/pdf files 2/Robinson et al. 2005 REPORT.pdf

_________________________________________________________________________________________________________________________________________________________________________________


276

www.digsfish.com

Rowland SJ, Ingram BA (1991). Diseases of Australian indigenous freshwater fishes (with emphasis on the

ectoparasitic and fungal diseases of Murray cod (Maccullochella peeli), golden perch (Macquaria ambigua)

and silver perch (Bidyanus bidyanus)). New South Wales Fisheries Bulletin 4: 33 pgs.

Rucker RR (1966). Redmouth disease of rainbow trout (Salmo gairdneri). Bull. Off. Int. des Epiz. 65: 825-

830.

Sahul-Hameed AS, Yoganandhan K, Widada JS, Bonami JR (2004). Experimental transmission and tissue

tropism of Macrobrachium rosenbergii nodavirus (MrNV) and its associated extra small virus (XSV).


Savin KW, Cocks BG, Wong F, Sawbridge T, Cogan N, Savage D, Warner S (2010). A neurotropic

herpesvirus infecting the gastropod, abalone, shares ancestry with oyster herpesvirus and a herpesvirus

associated with the amphioxus genome. Virology Journal 7: 308.

Saylor RK, Miller DL, Vandersea MW, Bevelhimer MS, Schofield PJ, Bennett WA (2010). Epizootic

ulcerative syndrome caused by Aphanomyces invadans in captive bullseye snakehead Channa marulius

collected from south Florida, USA. Diseases of Aquatic Organisms 88: 169-175.

Scandol J, Rowling K, Graham K (eds) (2008). Status of Fisheries Resources in NSW 2006/07, NSW

Department of Primary Industries, Cronulla, 334 pgs.

Schloegel LM, Picco AM, Kilpatrick AM, Davies AJ, Hyatt AD, Daszak P (2009). Magnitude of the US

trade in amphibians and presence of Batrachochytrium dendrobatidis and ranavirus infection in imported

North American bullfrogs (Rana catesbeiana). Biological Conservation 142: 1420-1428.

Scholz T, Salgado-Maldonado G (2000). The introduction and dispersal of Centrocestus formosanus

(Nishigori, 1924) (Digenea: Heterophyidae) in Mexico: a review. American Midland Naturalist 143: 185-

200.

Sebesvari Z, Esser F, Harder T (2006) . Sediment-associated cues for larval settlement of the infaunal

spionid polychaetes Polydora cornuta and Streblospio benedicti. Journal of Experimental Marine Biology

and Ecology 337: 109-120.

Sellars MJ, Keys SJ, Cowley JA, McCulloch RJ, Preston NP (2006). Association of Mourilyan virus with

mortalities in farm pond reared Penaeus (Marsupenaeus) japonicus transferred to maturation tank systems.


Sewell KB, Cannon LRG (1998). New Temnocephalans from the branchial chamber of Australian Euastacus

and Cherax crayfish hosts. Proceedings of the Linnean Society of New South Wales 119: 21-36.

Sharpe AK, Keough MJ (1998). An investigation of the indirect effects of intertidal shellfish collection.

Journal of Experimental Marine Biology and Ecology 223: 19-36.

_________________________________________________________________________________________________________________________________________________________________________________


277

www.digsfish.com

Shaw RW, Kent ML, Adamson ML (2000). Viability of Loma salmonae (Microsporidia) under laboratory

conditions. Parasitology Research 86: 978–981.

Shaw RW, Kent ML, Adamson ML (2001). Phagocytosis of Loma salmonae (Microsporidia) spores in

Atlantic salmon (Salmo salar), a resistant host, and Chinook salmon (Oncorhynchus tshawytscha), a

susceptible host. Fish and Shellfish Immunology 11: 91-100.

Shields JD (1992). Parasites and symbionts of the crab Portunus pelagicus from Moreton Bay, Eastern

Australia. Journal of Crustacean Biology 12: 94-100.

Shields JD (1994). The parasitic dinoflagellates of marine crustaceans. Annual Review of Fish Diseases 4:

241–271.

Shields JD, Woods FEI (1993). Impact of parasites on the reproduction and fecundity of the blue sand crab

Portunus pelagicus from Moreton Bay, Australia. Marine Ecology Progress Series 92: 159-170.

Shields JD, Behringer DC (2004). A new pathogenic virus in the Caribbean spiny lobster, Panulirus argus,

from Florida. Diseases of Aquatic Organisms 59: 109-118.

Shields JD, Taylor DM, Sutton SG, O’Keefe PO, Collins PW, Ings DW, Pardy AL (2005). Epizootiology of

bitter crab disease (Hematodinium sp.) in snow crabs, Chionoecetes opilio, from Newfoundland, Canada.

Diseases of Aquatic Organisms 64: 253–264.

Shields JD, Stephens FJ, Jones JB (2006). Pathogens, parasites and other symbionts. In: Phillips BF (ed.)

Lobsters: biology, management, aquaculture and fisheries. Blackwell Publishing. Pgs. 146-204.

Shivappa R, Kozlowicz S, Rolland J, Corsin F, Way K, Levine J (2008). Spring viremia of carp in the United

States of America: evaluation of current diagnostics, pp. 143-156. In Bondad-Reantaso, M.G., Mohan, C.V.,

Crumlish, M. and Subasinghe, R.P. (eds.). Diseases in Asian Aquaculture VI. Fish Health Section, Asian

Fisheries Society, Manila, Philippines. 505 pgs.

Skilleter GA, Zharikov Y, Cameron B, McPhee DP (2005). Effects of harvesting callianassid (ghost)

shrimps on subtropical benthic communities. Journal of Experimental Marine Biology and Ecology 320:

133-158.

Smith PA, Contreras JR, Garces LH, Larenas JJ, Oyanedel S, Caswell-Reno S, Fryer JL (1996a).

Experimental challenge of coho salmon and rainbow trout with Piscirickettsia salmonis. Journal of Aquatic

Animal Health 8: 130-134.

Smith PA, Rojas ME, Guajardo A, Contreras JR, Morales MA, Larenas JJ (2004). Experimental infection

of coho salmon Oncorhynchus kisutch by exposure of skin, gills and intestine with Piscirickettsia salmonis.


_________________________________________________________________________________________________________________________________________________________________________________


278

www.digsfish.com

Smith PJ, Holdsworth J, Anderson C, Hine PM, Allen D, Gibbs W, McKenzie L, Taylor PR, Blackwell RH,

Williamson SH (1996b). Pilchard deaths in New Zealand, 1995. New Zealand Fisheries Data Report No.

70. 52 pgs.

Smith PJ, McVeagh SM, Hulston D, Anderson SA, Gublin Y (2009). DNA identification of ciliates

associated with disease outbreaks in a New Zealand marine fish hatchery. Diseases of Aquatic Organisms

86: 163-167.

Smothers JF, von Dohlen CD, Smith LH, Spall RD (1994). Molecular evidence that the myxozoan protists

are metazoans. Science 265: 1719–1721.

Sommer AI, Strand MA, Rasmussen E, Mennen S (2004). Susceptibility of spotted wolffish Anarhichas

minor to experimental infection with nodavirus and infectious pancreatic necrosis virus. Diseases of Aquatic


Song JY, Kitamura SI, Oh MJ, Kang HS, Lee JH, Tanaka SJ, Jung SJ (2009). Pathogenicity of Miamiensis

avidus (syn. Philasterides dicentrarchi), Pseudocohnilembus persalinus, Pseudocohnilembus hargisi and

Uronema marinum (Ciliophora, Scuticociliatida). Diseases of Aquatic Organisms 83: 133–143.

Sosa ER, Landsberg JH, Stephenson CM, Forstchen AB, Vandersea MW, Litaker RW (2007). Aphanomyces

invadans and ulcerative mycosis in estuarine and freshwater fish in Florida. Journal of Aquatic Animal

Health 19: 14-26.

Soto MA, Shervette VR, Lotz JM (2001). Transmission of white spot syndrome virus (WSSV) to

Litopenaeus vannamei from infected cephalothorax, abdomen, or whole shrimp cadaver. Diseases of Aquatic


Southard GM, Fries LT and Terre DR (2009). Largemouth Bass virus in texas: Distribution and

management issues. Journal of Aquatic Animal Health 21: 36-42.

South Australian Museum (2010). Australian helminth collection. South Australian Museum, North

Terrace, Adelaide, South Australia 5000, Australia.

Spann KM, Lester RJG (1996). Baculovirus of Metapenaeus bennettae from the Moreton Bay region of

Australia. Diseases of Aquatic Organisms 27: 53-58.

Spann KM, Lester RJG (1997). Special Topic review: Viral diseases of penaeid shrimp with particular

reference to four viruses recently found in shrimp from Queensland. World ournal of Microbiology and

Biotechnology 13: 419-426.

Spann KM, Lester RJG, Paynter JL (1993). Efficiency of chlorine as a disinfectant against monodon

baculovirus (MBV). Asian Fisheries Science 6: 295-301.

_________________________________________________________________________________________________________________________________________________________________________________


279

www.digsfish.com

Spann KM, Vickers JE, Lester RJG (1995). Lymphoid organ virus of Penaeus monodon from Australia.

Diseases of Aquatic Organisms 23: 127–134.

Spann KM, Cowley JA, Walker PJ, Lester RJG (1997a). A yellow-head-like virus from Penaeus monodon

cultured in Australia. Diseases of Aquatic Organisms 31: 169–179.

Spann KM, Adlard RD, Hudson DA, Pyecroft SW, Jones TC, Voigt MOC (1997b). Hepatopancreatic parvo-

like virus (HPV) of Penaeus japonicus cultured in Australia. Diseases of Aquatic Organisms 31: 239-241.

Spann KM, Donaldson RA, Cowley JA, Walker PJ (2000). Differences in the susceptibility of some penaeid

prawn species to gill associated virus (GAV) infection. Diseases of Aquatic Organisms 42: 221–225.

Sparrow FK (1973). The peculiar marine phycomycete Atkinsiella dubia from crab eggs. Archives Fur

Mikrobiology 93: 137-144.

Sprague V, Couch J (1971). An annotated list of protozoan parasites, hyperparasites and commensals of

decapod crustacea. Journal of Protozoology 18: 526-537.

Sritunyalucksana K, Srisala J, Wangnai W, Flegel TW (2010). Yellow head virus (YHV) transmission risk

from commodity shrimp is reduced to negligible levels by normal processing. Aquaculture 300: 32-36.

Stanley CA (1978). Area of distribution, movements, age composition and mortality rates of the Australian

salmon population in Tasmania, Victoria and New South Wales. Australian Journal of Marine and


Stentiford GD, Shields JD (2005). A review of the parasitic dinoflagellates Hematodinium species and

Hematodinium-like infections in marine crustaceans. Diseases of Aquatic Organisms 66: 47-70.

Stephens FJ, Savage A (2010). Two mortality events in seacaged yellowtail kingfish Seriola lalandi

Valenciennes, 1833 (Nannopercidae) from Western Australia. Australian Veterinary Journal 88: 414-416.

Stephens FJ, Raidal SR, Jones B (2004). Haematopoietic necrosis in a goldfish (Carassius auratus)

associated with an agent morphologically similar to herpesvirus. Australian Veterinary Journal 82: 167-169.

Stewart JE (1991). Introductions as factors in diseases of fish and aquatic invertebrates. Canadian Journal

of Fisheries and Aquatic Sciences 48: 110-117.

St-Hilaire S, Hill M, Kent ML, Whitaker DJ, Ribble C (1997). A comparative study of muscle texture and

intensity of Kudoa thyrsites infection in farm reared Atlantic salmon Salmo salar on the Pacific coast of

Canada. Diseases of Aquatic Organisms 31: 221-225.

Sudhakaran R, Syed Musthaq S, Haribabu P, Mukherjee SC, Gopal C, Sahul Hameed AS (2006).

Experimental transmission of Macrobrachium rosenbergii nodavirus (MrNV) and extra small virus (XSV) in

_________________________________________________________________________________________________________________________________________________________________________________


280

www.digsfish.com

three species of marine shrimp (Penaeus indicus, Penaeus japonicus and Penaeus monodon). Aquaculture

257: 136–141.

Sudhakaran R, Ishaq Ahmed VP, Haribabu P, Mukherjee SC, Sri Widada J, Bonami JR, Sahul Hameed AS

(2007). Experimental vertical transmission of Macrobrachium rosenbergii nodavirus (MrNV) and extra

small virus (XSV) from brooders to progeny in Macrobrachium rosenbergii and Artemia. Journal of Fish

Diseases 30: 27–35.

Sudhakaran R, Haribabu P, Kumar SR, Sarathi M, Ahmed VP, Babu VS, Venkatesan C, Hameed AS (2008).

Natural aquatic insect carriers of Macrobrachium rosenbergii nodavirus (MrNV) and extra small virus

(XSV). Diseases of Aquatic Organisms 79: 141-145.

Sumpton WD, Potter MA, Smith GS (1994). Parasitism of the commercial sand crab Portunus pelagicus by

the rhizocephalan Sacculina granifera Boschma 1973 in Moreton Bay, QLD Australia. Australian Journal

of Marine and Freshwater Research 45: 169-175.

Sunfish (2008). Report by Chairman, Sunfish Queensland. In: QLD Amateur Fishing Clubs Association

Inc. Year Book 2008. pge. 23.

Sunila I, Karolus J, Lang EP, Mroczka ME, Volk J (2000). Transmission of the haplosporidian parasite

Haplosporidium nelsoni to the eastern oyster Crassostrea virginica in an upweller system. Diseases of


Sutherland DR (2002). Heterosporis update, March 12, 2002. .

http://michigan.gov/documents/heterosporis_29381_7.pdf

Swearer SE, Robertson DR (1999). Life history, pathology and description of Kudoa ovivora n. sp.

(Myxozoa, Myxosporea): an ovarian parasite of Carribbean labroid fishes. Journal of Parasitology 85: 337-

353.

Tan CK, Owens L (2000). Infectivity, transmission and 16s rRNA sequencing of a rickettsia, Coxiella

cheraxi sp. nov., from the freshwater crayfish Cherax quadricarinatus. Diseases of Aquatic Orgamisms 41:

115-122.

Tan J, Lancaster M, Hyatt A, van Driel R, Wong F, Warner S (2008). Purification of a herpes-like virus from

abalone (Haliotis spp.) with ganglioneuritis and detection by transmission electron microscopy. Journal of

Virological Methods 149: 338–341.

Tang KF, Poulos BT, Wang J, Redman RM, Shih HH, Lightner DV (2003). Geographic variations among

infectious hypodermal and hematopoietic necrosis virus (IHHNV) isolates and characteristics of their

infection. Diseases of Aquatic Orgamisms 53: 91-99.

http://michigan.gov/documents/heterosporis_29381_7.pdf

_________________________________________________________________________________________________________________________________________________________________________________


281

www.digsfish.com

Tang KF, Lightner DV (2006). Infectious hypodermal and hematopoietic necrosis virus (IHHNV)-related

sequences in the genome of the black tiger prawn Penaeus monodon from Africa and Australia. Virus

Research 118: 185-191.

Tang KF, Navarro SA, Lightner DV (2007). PCR assay for discriminating between infectious hypodermal

and hematopoietic necrosis virus (IHHNV) and virus-related sequences in the genome of Penaeus monodon.

Diseases of Aquatic Orgamisms 74: 165-170.

Tang KF, Pantoja CR, Lightner DV (2008). Nucleotide sequence of a Madagascar hepatopancreatic

parvovirus (HPV) and comparison of genetic variation among geographical isolates. Diseases of Aquatic

Orgamisms 80: 105-112.

Taylor PW (1977). Isolation and experimental infection of free living amebae in freshwater fishes. The

Journal of Parasitology 63: 232-237.

Thoney DA, Hargis WJ (1991). Monogenea (Platyhelminthes) as hazards for fish in confinement. Annual

Review of Fish Diseases 1991: 133-153.

Tidd WM, Shields RJ (1963). Tissue damage inflicted by Lernaea cyprinacea Linnaeus, a copepod parasitic

on tadpoles. The Journal of Parasitology 49: 693-696.

Tisdall DJ, Phipps JC (1987). Isolation and characterisation of a marine birnavirus from returning Quinnat

salmon (Onchorhynchus tshawtscha) in the South Island of New Zealand. New Zealand Veterinary Journal

35: 217-218.

Tobback E, Decostere A, Hermans K, Haesebrouck F, Chiers K (2007). Yersinia ruckeri infections in

salmonid fish. Journal of Fish Diseases 30: 257-268.

Tobback E, Decostere A, Hermans K, Ryckaert J, Duchateau L, Haesebrouck F, Chiers K (2009). Route of

entry and tissue distribution of Yersinia ruckeri in experimentally infected rainbow trout Oncorhynchus

mykiss. Diseases of Aquatic Organisms 84: 219-228.

Toranzo AE, Magarinos B, Romalde JL (2005). A review of the main bacterial fish diseases in mariculture

systems. Aquaculture 246: 37-61.

Tourtip S, Wongtripop S, Stentiford GD, Bateman KS, Sriurairatana S, Chavadej J, Sritunyalucksana K,

Withyachumnarnkul B (2009). Enterocytozoon hepatopenaei sp. nov. (Microsporida: Enterocytozoonidae),

a parasite of the black tiger shrimp Penaeus monodon (Decapoda: Penaeidae): Fine structure and

phylogenetic relationships. Journal of Invertebrate Pathology 102: 21-29.

Trust TJ, Khouri AG, Austen RA, Ashburner LD (1980). First isolation in Australia of atypical Aeromonas

salmonicida. FEMS Microbiology Letters 9(1): 39-42.

_________________________________________________________________________________________________________________________________________________________________________________


282

www.digsfish.com

Tung CW, Wang CS, Chen SN (1999). Histological and electron microscopic study on Macrobrachium

muscle virus (MMV) infection in the giant freshwater prawn, Macrobrachium rosenbergii (De Man),

cultured in Taiwan. Journal of Fish Diseases 22: 319-323.

Turnbull JF, Larkins PE, McPadden C, Matondang R (1994). A histopathological disease survey of cultured

shrimp in North East Sumatra, Indonesia. Journal of Fish Diseases 17: 57-65.

Unestam T (1969) Resistance to the crayfish plague in some American, Japanese and European crayfishes.

Rep. Inst. Freshw. Res. Drottningholm 49:202–209

Unestam T (1972) On the host range and origin of the crayfish plague fungus. Rep. Inst. Freshw. Res.

Drottningholm 52: 192–198

Unestam T (1975) Defence reactions in and susceptibility of Australian and New Guinean freshwater

crayfish to European- crayfish-plague fungus. Aust. J. Exp. Biol. Med. Sci. 53: 349–359

Unzueta-Bustamante ML, Holtschmit KH, Olivas-Valdez JA, Martinez-Cordova LR, Porchas-Cornejo MA,

Lizarraga-Partida ML (1998). Infectious hypodermal and hematopoietic necrosis virus (IHHNV) in wild

parent stocks of blue shrimp, Penaeus stylirostris (Stimpson), in Guaymas bay, Sonora, Mexico. Ciencias

Marinas 24: 491-498

Urquhart K, Murray AG, Gregory A, O’Dea M, Munro LA, Smail DA, Shanks AM, Raynard MS (2008).

Estimation of infectious dose and viral shedding rates for infectious pancreatic necrosis virus in Atlantic

salmon, Salmo salar L., post-smolts. Journal of Fish Diseases 31: 879-887.

USDA (2006). Viral Haemorrhagic septicaemia in the Great Lakes. July 2006. Emerging Disease Notice.

United States Department of Agriculture. .

http://www.aphis.usda.gov/vs/ceah/cei/taf/emergingdiseasenotice_files/vhsgreatlakes.htm

VanPatten KA, Nunan LM, Lightner DV (2004). Seabirds as potential vectors of penaeid shrimp viruses and

the development of a surrogate laboratory model utilizing domestic chickens. Aquaculture 241: 31-46.

Vela A, Vázquez J, Gibello A, Blanco MM, Moreno MA, Liébana P, Albendea C, Alcalá B, Mendez A,

Domínguez L, Fernández-Garayzábal J (2000). Phenotypic and genetic characterization of Lactococcus

garvieae isolated in Spain from lactococcosis outbreaks and comparison with isolates of other countries and

sources. Journal of Clinical Microbiology 38: 3791-3795.

VHSV Expert Panel and Working Group (2010). Viral haemorrhagic septicaemia virus (VHSV-Ib) risk

factors and association measures derived by expert panel. Preventative Veterinary Medicine 94:128-139.

Victoria DPI (2006). Bulletin for Abalone Industry – 9 May 2006, Update on herpes like virus in farmed

abalone.

http://www.aphis.usda.gov/vs/ceah/cei/taf/emergingdiseasenotice_files/vhsgreatlakes.htm

_________________________________________________________________________________________________________________________________________________________________________________


283

www.digsfish.com

Victoria DPI (2010). Recreational fishing guide 2009-2010. State of Victoria, Department of Primary

Industries. 69 pgs. http://www.dpi.vic.gov.au/DPI/nrenfaq.nsf/childdocs/-

B43DB2A1BECEB2A04A256812001DD817-038067B67D8D0260CA2571F6007C592F?open

Vidal-Martinez VM, Jimenez-Cueto AM, Sima-Alvarez R (2002). Parasites and symbionts of native and

cultured shrimps from Yucatan, Mexico. Journal of Aquatic Animal Health 14: 57-64.

Vijayan KK, Raj VS, Alavandi SV, Sekhar VT, Vaseeharan B, Santiago TC (2003). Need of novel

diagnostics in the health management of Macrobrachium rosenbergii, with special reference to an emerging

epizootic in India, the white muscle syndrome (WMS). In Freshwater Prawns 2003, International

Symposium, Cochin , 20-23 August 2003.

Vijayan KK, Stalin Raj V, Balasubramanian CP, Alavandi SV, Thillai Sekhar V, Santiago TC (2005a).

Polychaete worms - A vector for white spot syndrome virus (WSSV). Diseases of Aquatic Organisms 63:

107-11.

Vijayan KK, Raj VS, Alavandi SV, Sekhar VT, Santiago TC (2005b). Incidence of white muscle disease, a

viral like disease associated with mortalities in hatchery reared postlarvae of the giant freshwater prawn

Macrobrachium rosenbergii (deMan) from the south-east coast of India. Aquaculture Research 36: 311-

316.

Villalba A., Reece KS, Ordas MC, Casa SM, Figueras A (2004). Perkinsosis in molluscs: a review. Aquatic

Living Resources 17: 411–432.

Voelker FA, Anver MR, McKee AE, Casey HW, Brenniman GR (1977). Amebiasis in goldfish. Veterinary


Vogelbein WK, Overstreet RM (1988). Life-history and pathology of a heterophyid trematode infecting

Florida-reared ornamental fishes. International Association for Aquatic Animal Medicine Proceedings 19:

138.

Vogelbein WK, Shields JD, Haas LW, Reece KS, Zwerner DE (2001). Skin Ulcers in Estuarine Fishes: A

Comparative Pathological Evaluation of Wild and Laboratory-Exposed Fish. Environmental Health

Perspectives 109: 687-693.

Vossbrinck CR, Andreadis TG, Debrunner-Vossbrinck BA (1998). Verification of intermediate hosts in the

life cycles of microsporidia by small subunit rDNA sequencing. Journal of Eukaryotic Microbiology 45:

290-292

Walker AN, Lee RF, Frischer ME (2009). Transmission of the parasitic dinoflagellates Hematodinium sp.

infection in the blue crabs Callinectes sapidus by cannibalism. Diseases of Aquatic Organisms 85: 193–197.

Walker G (2001). Introduction to the Rhizocephala (Crustacea: Cirripedia). Journal of Morphology 249: 1–8.

http://www.dpi.vic.gov.au/DPI/nrenfaq.nsf/childdocs/-B43DB2A1BECEB2A04A256812001DD817-038067B67D8D0260CA2571F6007C592F?open

_________________________________________________________________________________________________________________________________________________________________________________


284

www.digsfish.com

Walker G, Lester RJG (1998). Effect of salinity on development of larvae of Heterosaccus lunatus

(Cirripedia: Rhizocephala). Journal of Crustacean Biology 18: 650-655.

Walker LM (2009). Polydora and Dipolydora (Polychaeta: Spionidae) of estuaries and bays of subtropical

eastern Australia: A review and morphometric investigation of their taxonomy and distribution. MSc thesis,

Southern Cross University, Lismore, NSW. 180 pgs.

Walker LM (2011). A review of the current status of the Polydora-complex (Polychaeta: Spionidae) in

Australia and a checklist of recorded species. Zootaxa 2751: 40-62.

Walker PJ, Mohan CV (2009). Viral disease emergence in shrimp aquaculture: origins, impact and the

effectiveness of health management strategies. Reviews in Aquaculture 1: 125-154.

Waltzek TB, Kelley GO, Stone DM, Way K, Hanson L, Fukuda H, Hirono I, Aoki T, Davison AJ, Hedrick

RP (2005). Koi herpesvirus represents a third cyprinid herpesvirus (CyHV-3) in the family Herpesviridae.

Journal of General Virology 86: 1659–1667.

Wechsler SJ, Woods LC, Kraeuter JN, Hetrick FM, McAllister PE (1987). Transmission of infectious

pancreatic necrosis virus in striped bass, Morone saxatilis. Journal of Fish Diseases 10: 29-34.

Welsford DC, Lyle JM (2003). Redbait (Emmelichthys nitidus): a synopsis of fishery and biological data.

TAFI Technical Report Series No. 20. 32 pgs.

Weng HT (1987). The parasitic barnacle, Sacculina granifera Boschma, affecting the commercial sand crab,

Portunus pelagicus (L.), in populations from two different environments in Queensland. Journal of Fish

Diseases 10: 221-227.

Wesche SJ (1995). Outbreaks of Marteilia sydneyi in Sydney rock oysters and their relationship with

environmental pH. Bulletin of the European Association of Fish Pathologists 15: 23-27.

Wesche SJ, Adlard RD, Lester RJG (1999). Survival of spores of the oyster pathogen Marteilia sydneyi

(Protozoa, Paramyxea) as assessed using fluorogenic dyes. Diseases of Aquatic Organisms 36: 221-226.

Western Australian Department of Fisheries (2002). The Introduction and Aquaculture of Non-endemic

Species in Western Australia: the ‘Rotund’ Yabby Cherax rotundus and the All-male Hybrid Yabby. A

Discussion Paper. Fisheries Management Paper No. 160. June 2002. 30 pgs.

Westman K (1991). The crayfish fishery in Finland-its past, present and future. Finnish Fisheries Research

12: 187-216.

Wetchateng T, Friedman CS, Wight NA, Lee PY, Teng PH, Sriurairattana S, Wongprasert K,

Withyachumnarnkul B (2010). Withering syndrome in the abalone Haliotis diversicolor supertexta. Diseases


_________________________________________________________________________________________________________________________________________________________________________________


285

www.digsfish.com

Whipps, CM, Adlard, RD, Bryant, MS, Lester, RJG, Findlay, V and Kent, ML (2003) First report of three

Kudoa species from Eastern Australia: Kudoa thyrsites from Mahi mahi (Coryphaena hippurus), Kudoa

amamiensis and Kudoa minithyrsites n. sp. from sweeper (Pempheris ypsilychnus). Journal of Eukaryotic


White ME, Powell EN, Ray SM, Wilson EA (1987). Host to host transmission of Perkinsus marinus in

oyster (Crassostrea virginica) populations by the ectoparasitic snail Boonea impressa (Pyramidellidae).


Whitelegge T (1890). Report on the worm disease affecting the oysters on the coast of New South Wales.

Records of the Australian Museum 1: 1 – 15.

Whittington ID, Horton MA (1996). A revision of Neobenedenia Yamaguti, 1963 (Monogenea: Capsalidae)

including a redescription of N. melleni (McCallum 1927). Journal of Natural History 30: 1113-1156.

Whittington ID, Cribb BW, Hamwood TE, Halliday JA (2000). Host-specificity of monogeneans

(Platyhelminth) parasites: a role for anterior adhesive areas ? International Journal for Parasitology 30:

305-320.

Whittington ID, Deveney MR, Wyborn SJ (2001). A revision of Benedenia Deising, 1858 including a

rediscription of B. sciaenae (van Beneden 1856) Odhner, 1905 and recognition of Menziesia Gibson 1976

(Monogenea: Capsalidae). Journal of Natural History 35: 663-777.

Whittington RJ, Cullis B (1988). The susceptibility of salmonid fish to an atypical strain of Aeromonas

salmonicida that infects goldfish, Carassius auratus (L.) in Australia. Journal of Fish Diseases 11: 461-470.

Whittington RJ, Reddacliff GL (1995). Influence of environmental temperature on experimental infection of

redfin perch (Perca fluviatilis) and rainbow trout (Oncorhynchus mykiss) with epizootic haematopoietic

necrosis virus, an Australian iridovirus. Australian Veterinary Journal 72: 421–424.

Whittington RJ, Chong R (2007). Global trade in ornamental fish from an Australian perspective: The case

for revised import risk analysis and management strategies. Preventive Veterinary Medicine 81 (1-3 Spec.

Iss.): pgs. 92-116.

Whittington RJ, Gudkovs N, Carrigan MJ, Ashburner LD, Thurstan SJ (1987). Clinical, microbiological and

epidemiological findings in recent outbreaks of goldfish ulcer disease due to atypical Aeromonas

salmonicida in south-eastern Australia. Journal of Fish Diseases 10: 353-362.

Whittington RJ, Djordjevic SP, Carson J, Callinan RB (1995). Restriction endonuclease analysis of atypical

Aeromonas salmonicida isolates from goldfish Carassius auratus, silver perch Bidyanus bidyanus, and

greenback flounder Rhombosolea tapirina in Australia. Diseases of Aquatic Organisms 22: 185-191.

_________________________________________________________________________________________________________________________________________________________________________________


286

www.digsfish.com

Whittington RJ, Kearns C, Hyatt AD, Hengstberger S, Rutzou T (1996). Spread of epizootic haematopoietic

necrosis virus (EHNV) in redfin perch (Perca fluviatilis) in southern Australia. Australian Veterinary

Journal 73: 112-114.

Whittington RJ, Jones JB, Hine PM, Hyatt AD (1997). Epizootic mortality in the pilchard Sardinops sagax

neopilchardus in Australia and New Zealand in 1995. 1. Pathology and epizootiology. Diseases of Aquatic


Whittington RJ, Reddacliff LA, Marsh I, Kearns C, Zupanovic Z, Callinan RB (1999). Further observations

on the epidemiology and spread of epizootic haematopoietic necrosis virus (EHNV) in farmed rainbow trout

(Oncorhynchus mykiss) in southeastern Australia and a recommended sampling strategy for surveillance.


Whittington RJ, Crockford M, Jordan D, Jones B (2008). Herpesvirus that caused epizootic mortality in

1995 and 1998 in pilchard, Sardinops sagax neopilchardus (Steindachner), in Australia is now endemic.


Whittington RJ, Becker JA, Dennis MM (2010). Iridovirus infections in finfish – critical review with

emphasis on ranaviruses. Journal of Fish Diseases 33: 95-122.

Wickins SC, Dennis MM, Landos M, Slapeta J, Whittington RJ (2011). Histopathological survey of lesions

and infections affecting sick ornamental fish in pet shops in New�South Wales, Australia. Diseases of


Wicklund T, Dalsgaard I (1998). Occurrence and significance of atypical Aeromonas salmonicida in non-

salmonid and salmonid fish species: a review. Diseases of Aquatic Organisms 32: 49-69

Wijegoonawardane PK, Sittidilokratna N, Petchampai N, Cowley JA, Gudkovs N, Walker PJ (2009).

Holmologous genetic recombination in the yellowhead complex of nidoviruses infecting Penaeus monodon

shrimp. Virology 390: 79-88.

Wilhelm G, Boulo V (1988). Infection de l’etrille Liocarcinus puber (L.) par un dinoflagellate parasite:

Hematodinium sp. Cons. Int. Explor. Mer. Ser. CM K: 32: 1–10.

Wilhelm G, Mialhe E (1996). Dinoflagellate infection associated with the decline of Necora puber crab

populations in France. Diseases of Aquatic Organisms 26: 213–219.

Wilson C, Heckmann R, Goede R, Inchausty V (1995). Intracranial amoebiasis in ornamental goldfish

(Carassius auratus): a case report. Fish Health Section Newsletter, American Fisheries Society 23: 1-3.

Wisconsin Department of Natural Resources (2005). Heterosporis sp. yellow perch parasite. Fact Sheet. 2

pgs. http://dnr.wi.gov/fish/documents/heterosporis_factsheet.pdf

http://dnr.wi.gov/fish/documents/heterosporis_factsheet.pdf

_________________________________________________________________________________________________________________________________________________________________________________


287

www.digsfish.com

Wisconsin Aquaculture Association (2007). VHS found in inland lake in Michigan. The Creel 40(2): 6.

http://www.wisconsinaquaculture.com/Forms/2007_June_Web_Creel.pdf

Withyachumnarnkul B, Chayaburakul K, Lao-Aroon S, Plodpai P, Sritunyalucksana K, Nash G (2006). Low

impact of infectious hypodermal and hematopoietic necrosis virus (IHHNV) on growth and reproductive

performance of Penaeus monodon. Diseases of Aquatic Organisms 69: 129-136.

Wolf PH (1972). Occurrence of a haplosporidan in Sydney rock oysters Crassostrea commercialis from

Moreton Bay, Queensland, Australia. Journal of Invertebrate Pathology 19: 416-417.

Wolf PH (1977). An unidentified protistan parasite in the ova of the blacklipped oyster, Crassostrea

echinata, from northern Australia. Journal of Invertebrate Pathology 29: 244-246.

Wolf K, Markiw ME (1984). Biology contravenes taxonomy in the Myxozoa: new discoveries show

alternation of the invertebrate and vertebrate hosts. Science 225: 1449-1452.

Wolf K, Snieszko SF, Dunbar CE, Pyle E (1960). Virus nature of infectious pancreatic necrosis in trout.

Proceedings of the Society for Experimental Biology and Medicine 104: 105–108.

Woodburn KD, Eldred B, Clark E, Hutton RF, Ingle RM (1957) . The live bait shrimp industry of the west

coast of Florida (Cedar Key to Naples). Florida State Board of Conservation Marine Laboratory Technical

Series 21, 33 pgs.

Woodland JE, Brunner CJ, Noyes AD, Grizzle JM (2002a). Experimental oral transmission of largemouth

bass virus. Journal of Fish Diseases 25: 669-672.

Woodland JE, Noyes AD, Grizzle JM (2002b). A survey to detect largemouth bass virus among fish from

hatcheries in the southeastern USA. Transactions of the American Fisheries Society 131: 308–311.

Xu W, Xie J, Shi H, Li C (2010). Hematodinium infections in cultured ridgetail white prawns, Exopalaemon

carinicauda, in eastern China. Aquaculture 300: 25-31

Yarish C, Whitlatch R, Kraemer G, Lin S (2009). Multi-Component Evaluation to Minimize the Spread of

Aquatic Invasive Seaweeds, Harmful Algal Bloom Microalgae, and Invertebrates via the Live Bait Vector in

Long Island Sound. Dept. Ecol. Evol. Biol. University of Connecticut. 23 pgs. .

http://digitalcommons.uconn.edu/cgi/viewcontent.cgi?article=1001&context=ecostam_pubs

Yasuda H, Ooyama T, Iwata K, Tun T, Yokoyama H, Ogawa K (2002). Fish to fish transmission of

Myxidium spp. (Myxozoa) in cultured tiger puffer suffering from emaciation disease. Fish Pathology 37:

29-33.

Yoganandhan K, Leartvibhas M, Sriwongpuk S, Limsuwan C (2006). White tail disease of the giant

freshwater prawn Macrobrachium rosenbergii in Thailand. Diseases of Aquatic Organisms 69: 255-258.

http://www.wisconsinaquaculture.com/Forms/2007_June_Web_Creel.pdf

http://digitalcommons.uconn.edu/cgi/viewcontent.cgi?article=1001&context=ecostam_pubs

_________________________________________________________________________________________________________________________________________________________________________________


288

www.digsfish.com

Young ND, Crosbie PBB, Adams MB, Nowak BF, Morrison, RN (2007). Neoparamoeba perurans n. sp. an

agent of amoebic gill disease in Atlantic salmon (Salmo salar L.) International Journal for Parasitology 37:

1469-1481.

Young ND, Dykova I, Snekvik K, Nowak BF, Morrison, RN (2008). Neoparamoeba perurans is a

cosmopolitan aetiological agent of amoebic gill disease. Diseases of Aquatic Organisms 78: 217-223.

Zetlmeisel C, Hermann J, Petney T, Glenner H, Griffiths C, Taraschewski H (2011). Parasites of the shore

crab Carcinus maenas (L.): implications for reproductive potential and invasion success. Parasitology 138:

394-401

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

28

9

w

ww

.dig

sfis

h.co

m

App

endi

x 1.

The

dis

ease

age

nts

know

n to

infe

ct a

quat

ic a

nim

als

that

are u

sed

as b

ait a

nd b

erle

y.

Dat

a ga

ps a

re in

dica

ted

where

the

y oc

cur.

Sci

entif

ic n

ame

D

isea

se a

gen

t R

efer

ence

s F

INF

ISH

S

altw

ater

fin

fish

F. A

rrip

ida

e

Cop

epod

a,

Ces

toda

, M

onog

enea

, M

yxoz

oa,

Nem

at

oda

, Tre

ma

toda

O

’Don

oghu

e a

nd

Adl

ard

(2

000)

, C

ata

lano

(2

010a

, 201

0b)

F. A

ther

inid

ae

E

imer

ia s

pp.,

Myx

ozoa

, Tre

ma

toda

K

ent

et a

l. (1

989)

, O

’Don

oghu

e a

nd A

dla

rd

(200

0)

F. C

ara

ngid

ae

V

ER

/Nod

avi

rus,

G

ou

ssia

spp.

, ly

mp

hocy

stis

, Ku

doa

spp

., A

cant

hoce

pha

la,

Cop

epod

a,

Ces

toda

, Is

opod

a,

Mon

ogen

ea,

othe

r M

yxoz

oa,

Nem

ato

da,

Tre

ma

toda

R

edda

cliff

a

nd

Qua

rtar

aro

(1

992)

, P

erer

a

(199

3),

O’D

onog

hue

and

A

dla

rd

(200

0),

Dig

gles

(20

02),

And

erso

n a

nd O

ake

y (2

008)

F

. Clu

peid

ae

P

ilcha

rd h

erp

esvi

rus

(PH

V),

K

ud

oa

spp.

, N

ema

toda

, Tre

ma

toda

, M

icro

spor

ea ,

oth

er M

yxoz

oa

Whi

ttin

gton

et

a

l. 19

97,

O’D

onog

hue

and

A

dla

rd (

2000

), C

rock

ford

(20

07)

F. C

ongr

ida

e

Dat

a ga

p F

. Em

mel

icht

hyid

ae

Mon

ogen

ea,

Tre

ma

toda

, C

esto

da,

Nem

ato

da

P

aruk

uhin

(19

85),

Kor

ota

eva

(19

75)

F

. Eng

raul

ida

e

Ku

do

a sp

p., M

icro

spor

ea,

Nem

ato

da, T

rem

ato

da

O’D

onog

hue

and

Adl

ard

(20

00)

F. G

emp

ylid

ae

Mic

rosp

orea

, Ku

do

a sp

p., C

esto

da

O’D

onog

hue

and

Adl

ard

(20

00)

F. G

errid

ae

E

imer

ia s

pp.,

Myx

ozoa

, Tre

ma

toda

, C

iliop

hora

O

’Don

oghu

e a

nd A

dla

rd (

2000

) F

. Hem

iram

phi

dae

A

cant

hoce

pha

la,

Cop

epod

a, C

esto

da,

Isop

oda

, M

onog

enea

, M

yxoz

oa,

Nem

ato

da,

Tre

ma

toda

Le

ster

a

nd

Sew

ell

(198

9),

Gun

ter

et

al.

(201

0),

Sou

th A

ustr

alia

n M

useu

m (

2010

) F

. La

brid

ae

Mic

rosp

orea

, ot

her

Myx

ozoa

, C

iliop

hora

, Tre

ma

toda

O

’Don

oghu

e a

nd A

dla

rd (

2000

) F

. Lei

ogna

thid

ae

Dat

a ga

p F

. Let

hrin

ida

e

Cili

opho

ra, C

opep

oda

, C

esto

da,

Hiru

dine

a,

Iso

poda

, M

onog

enea

, M

yxoz

oa,

Nem

ato

da,

Tre

ma

toda

Le

ster

a

nd

Sew

ell

(198

9),

Bru

ce

(199

0),

O’D

onog

hue

and

Adl

ard

(20

00)

F. L

utja

nida

e

EU

S/ Ap

ha

no

myc

es in

vad

an

s, B

ened

enia s

pp., G

yro

dac

tylu

s spp

., ot

her

Mon

ogen

ea,

Cili

opho

ra,

Ces

toda

, M

yxoz

oa,

Nem

ato

da

Bru

ce

(199

0),

O’D

onog

hue

and

A

dla

rd

(200

0),

Whi

ttin

gton

et

al.

(200

1)

F. M

onoc

ant

hida

e E

imer

ia s

p, o

ther

coc

cidi

a,

unid

entif

ied

haem

opro

tozo

an,

Mic

rosp

ore

a,

othe

r M

yxoz

oa,

Tre

ma

toda

Iso

pod

a

O’D

onog

hue

and

Adl

ard

(20

00)

F. M

orid

ae

Pse

ud

om

on

as spp

, Vib

rio

spp.

, Nem

ato

da ,

Iso

poda

, Cop

epod

a

Mun

day

(198

5),

Bru

ce (

1990

) F

. Mug

lida

e

EU

S/ A

ph

an

om

yces

inva

da

ns

, E

imer

ia s

p., C

iliop

hora

, Cop

epod

a,

Ces

toda

, Is

opod

a,

Myx

ozoa

, M

onog

enea

, T

rem

ato

da

Dov

e (2

000)

, O

’Don

oghu

e a

nd

Adl

ard

(2

000)

, H

ump

hrey

and

Pea

rce

(200

4)

F. N

emip

teri

dae

C

opep

oda

, M

onog

enea

, N

ema

toda

, Tre

ma

toda

Lest

er a

nd S

ewel

l (19

89),

Gru

tter

(19

94)

F. S

com

brid

ae

C

ard

ico

la s

p. , K

ud

oa

spp.

, Uro

nem

a spp

., C

opep

oda

, Ces

toda

, M

onog

enea

, ot

her

Myx

ozoa

, ot

her

Tre

ma

toda

, N

ema

toda

H

um

phre

y (1

995a

), O

’Don

oghu

e a

nd A

dla

rd

(200

0),

Dev

eney

et

al.

(200

5)

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

29

0

w

ww

.dig

sfis

h.co

m

Sci

entif

ic n

ame

D

isea

se a

gen

t R

efer

ence

s

F. S

illa

gini

dae

E

US

/ Ap

ha

no

myc

es in

vad

an

s, G

yro

da

ctyl

us s

pp., K

ud

oa s

pp., E

imer

ia s

pp.,G

ou

ssia

spp.

, C

opep

oda

, C

esto

da,

Uro

nem

a spp

., ot

her

Cili

opho

ra, H

irudi

nea

, Is

opod

a,

othe

r M

onog

enea

, ot

her

Myx

ozoa

, N

ema

toda

, Cre

pid

oo

din

ium

aus

tra

le

Bru

ce

(199

0),

Gill

a

nd

Ca

llina

n (1

997)

, H

ayw

ard

(1

997)

, O

’Don

oghu

e a

nd

Adl

ard

(2

000)

F

. Spa

rida

e

EU

S/ Ap

ha

no

myc

es in

vad

an

s, B

ened

enia s

pp., C

alig

us

epid

emic

us, G

ou

ssia

spp.

, ly

mp

hocy

stis

, C

iliop

hora

, ot

her

Cop

epod

a,

othe

r M

onog

enea

, M

yxoz

oa, T

rem

ato

da

Rou

bal

et

al.

(198

3),

Red

dacl

iff

and

Q

uart

ara

ro

(199

2),

Ca

llina

n et

a

l. (1

995)

, O

’Don

oghu

e a

nd A

dla

rd (

2000

) T

ailo

r (P

om

ato

mu

s sa

ltatr

ix)

Mon

ogen

ea,

Nem

ato

da,

Aca

ntho

cep

hala

, T

rem

ato

da,

Isop

oda

Jo

hns

ton

and

D

ela

nd

(192

9),

Bru

ce

1987

, B

ray

and

Crib

b (1

998)

, Bra

y (1

990)

Fre

shw

ater

finf

ish

F

. Am

bass

ida

e

EU

S/ Ap

ha

no

myc

es in

vad

an

s, C

lino

sto

mu

m a

ust

ralie

nse

, C

esto

da,

Cili

opho

ra, F

lage

llate

s,

Myx

ozoa

, N

ema

toda

, Tre

ma

toda

D

ove

(200

0),

O’D

onog

hue

and

A

dla

rd

(200

0),

Hum

phr

ey a

nd P

earc

e (2

004)

C

ara

ssiu

s a

ura

tus

Ha

ema

top

oiet

ic n

ecro

sis

herp

esvi

rus

of g

oldf

ish,

Her

pes

-lik

e vi

rus,

Bot

hri

oce

ph

alu

s a

chei

log

na

thi,

EU

S/A

ph

an

om

yces

inva

da

ns,

Aer

om

ona

s sa

lmo

nici

da

(a

typi

cal),

Yer

sin

ia

ruck

eri,

Eim

eria

sp.,

Ho

fere

llus

cara

ssii,

Myx

ob

olu

s spp.

, D

act

ylo

gyr

us

an

cho

ratu

s,

Gyr

od

act

ylu

s ko

ba

yash

ii, L

ern

aea

cyp

rin

ace

a, L

ern

aea

sp

p.,

Mitr

asp

ora

cyp

rin

id, A

rgu

lus

spp.

, C

iliop

hora

, Fla

gella

tes, o

ther

Myx

ozoa

, S

yste

mic

am

oebi

asi

s

Hu

mph

rey

and

Ash

burn

er (

1993

), D

ove

and

F

letc

her

(200

0),

O’D

onog

hue

and

A

dla

rd

(200

0),

Ste

phen

s et

al.

(200

4),

Cor

field

et

al.

(200

7)

F. C

ichl

ida

e

Boh

le ir

idov

irus,

Str

epto

cocc

us s

pp.,

EU

S/ A

ph

an

om

yces

inva

da

ns

, Cen

tro

cest

us s

pp. ,

C

iliop

hora

A

riel

a

nd

Ow

ens

(199

7),

Cor

field

et

a

l. (2

007)

C

ypri

nu

s ca

rpio

Bo

thri

oce

ph

alu

s a

chei

log

na

thi,

Aer

om

on

as

salm

on

icid

a (

aty

pica

l), D

act

ylo

gyr

us

exte

nsu

s,

Ler

na

ea c

ypri

na

cea

/ Ler

na

ea sp

p., S

ph

aer

osp

ora

spp.

, Mitr

asp

ora

spp

., C

iliop

hora

H

um

phre

y (1

995a

), D

ove

et a

l. (1

997)

, D

ove

and

Fle

tche

r (2

000)

, O

’Don

oghu

e a

nd A

dla

rd

(200

0)

Ele

otri

dae

V

ER

/Nod

avi

rus,

EU

S/ Ap

ha

no

myc

es in

vad

ans,

Bot

hrio

cep

ha

lus

ach

eilo

gn

ath

i, C

lino

sto

mu

m

au

stra

lien

se, H

enn

egu

ya sp

p., M

yxo

bo

lus s

pp., P

seu

do

da

ctyl

og

yro

ides s

pp., G

yro

da

ctyl

us s

pp.,

Erg

asi

lus s

pp., Z

sch

okk

ella s

pp.,

Cili

opho

ra, N

ema

toda

, Tre

ma

toda

, C

esto

da,

Mic

rosp

ore

a

Dov

e et

al.

(199

7),

Dov

e (2

000)

, D

ove

and

F

letc

her

(200

0),

O’D

onog

hue

and

A

dla

rd

(200

0),

Hum

phr

ey a

nd P

earc

e (2

004)

F

. Ga

laxi

ida

e L

igu

la in

test

ina

lis, L

ern

aea

cyp

rin

ace

a, Myx

ob

olu

s spp

., C

opep

oda

, C

iliop

hora

O

’Don

oghu

e a

nd A

dla

rd (

2000

), L

ymb

ery

et

al.

(201

0)

F. H

emirh

am

phi

dae

D

ata

gap

F. M

ela

nota

eniid

ae

E

US

/ Ap

ha

no

myc

es in

vad

an

s, C

lino

sto

mu

m a

ust

ralie

nse, C

iliop

hora

, Fla

gella

tes,

Tre

ma

toda

D

ove

(200

0),

O’D

onog

hue

and

A

dla

rd

(200

0),

Hum

phr

ey a

nd P

earc

e (2

004)

M

isg

urn

us

an

gu

illic

au

dat

us

Gyr

od

act

ylu

s m

acr

aca

nth

us

, Cili

opho

ra

Dov

e a

nd E

rnst

(19

98)

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

29

1

w

ww

.dig

sfis

h.co

m

Sci

entif

ic n

ame

D

isea

se a

gen

t R

efer

ence

s F

. Mug

lida

e E

US

/ Ap

ha

no

myc

es in

vad

an

s,

Cili

opho

ra ,

Cop

epod

a,

Myx

ozoa

, Tre

ma

toda

D

ove

(200

0),

O’D

onog

hu

e a

nd

Adl

ard

(2

000)

, H

ump

hrey

and

Pea

rce

(200

4)

Nem

ata

losa

spp.

E

US

/ Ap

ha

no

myc

es in

vad

an

s, H

enn

ygu

ya sp

p., B

act

eria

l dis

ease

, F

unga

l dis

ease

, M

yxoz

oa

John

ston

a

nd H

itchc

ock

(192

3),

O’D

onog

hue

and

A

dla

rd

(200

0),

Hum

phr

ey

and

P

earc

e (2

004)

O

nco

rhyn

chu

s m

ykis

s E

HN

V,

Aqu

atic

Birn

avi

rus,

Yer

sin

ia r

uck

eri, L

act

oco

ccu

s g

arv

iea

e, B

act

eria

l dis

ease

, F

unga

l di

sea

se,

Cili

opho

ra F

lage

llate

s,

Neo

pa

ram

oeb

a sp.

C

arso

n et

a

l. (1

993)

, H

um

phre

y (1

995a

),

Whi

ttin

gton

et

al.

(199

9),

Cra

ne e

t a

l. 20

00,

O’D

onog

hue

and

Adl

ard

(20

00),

P

erca

flu

via

tilis

E

HN

V,

Reo

viru

s, Tri

ang

ula

per

cae,

Cili

opho

ra

Lang

don

et

al.

(198

6),

Lang

don

(198

8,

1989

b,

1990

),

Hum

phre

y (1

995a

),

Whi

ttin

gton

et

al.

(199

9)

F. P

oeci

liida

e

EH

NV

, Bo

thri

oce

ph

alu

s a

chei

log

na

thi

, Gyr

od

act

ylu

s bu

llata

rudi

s, Uro

clei

do

ides

ret

icu

late

s,

Ca

ma

llan

us

cotti,

Cen

tro

cest

us s

p.,

Pro

totr

ansv

erso

trem

a s

teer

i, G

ou

ssia

spp.

, Ler

na

ea

cyp

rin

ace

a, ot

her

Cop

epod

a, C

iliop

hora

, Fla

gella

tes

Lang

don

(198

9a,

1989

b, 1

990)

, D

ove

et a

l. (1

997)

, D

ove

and

Ern

st (

1998

), D

ove

(200

0),

Dov

e a

nd F

letc

her

(200

0),

O’D

onog

hue

and

A

dla

rd (

2000

), M

arin

a e

t a

l. (2

008)

F

. Ret

ropi

nnid

ae

B

oth

rio

cep

ha

lus

ach

eilo

gn

ath

i, G

yro

da

ctyl

us

spp

., Myx

idiu

m s

pp., L

ern

aea

cyp

rin

ace

a /

Ler

na

ea sp

p., C

iliop

hora

, Tre

ma

toda

R

owla

nd a

nd I

ngra

m (

1991

), D

ove

(200

0),

Dov

e a

nd F

letc

her

(200

0),

O’D

onog

hue

and

A

dla

rd (

2000

) F

. Ter

apo

nida

e

VE

R/N

oda

viru

s,

EU

S/

Ap

ha

no

myc

es in

vad

an

s, C

linos

tom

um

spp.

, Eim

eria

spp

., Hen

neg

uya

spp.

, Ler

na

ea c

ypri

na

cea

/ Ler

na

ea sp

p., C

esto

da,

Cop

epod

a,

Cili

opho

ra ,

Fla

gella

tes,

M

onog

enea

, T

rem

ato

da,

oth

er M

yxoz

oa

Row

land

and

Ing

ram

(19

91),

Dov

e (2

000)

, O

’Don

oghu

e a

nd A

dla

rd (

2000

), H

um

phre

y a

nd P

earc

e (2

004)

T

oxo

tes s

p.

EU

S/ A

ph

an

om

yces

inva

da

ns

H

um

phre

y a

nd P

earc

e (2

004)

,

CR

US

TA

CE

AN

S

Pra

wns

and

shr

imp

M

acr

ob

rach

ium

ro

sen

ber

gii

Whi

te T

ail

Dis

ease

/MrN

V,

HP

V,

Cili

opho

ra ,

Mic

rosp

orid

ia

A

nder

son

et

al.

(199

0),

O’D

onog

hue

and

A

dla

rd (

2000

) O

wen

s et

al.

(200

9)

Ma

cro

bra

chiu

m sp

p.

HP

V,

Cili

opho

ra

And

erso

n et

a

l. (1

990)

, O

’Don

oghu

e a

nd

Adl

ard

(20

00)

Met

ap

ena

eop

sis spp

.

Dat

a ga

p M

eta

pen

aeu

s ben

net

tae

Ben

etta

e ba

culo

viru

s/M

BV

S

pann

and

Les

ter

(199

6),

M

eta

pen

aeu

s m

acl

eayi

Cili

opho

ra

O’D

onog

hue

and

Adl

ard

(20

00)

Met

ap

ena

eus s

pp.

Ben

etta

e ba

culo

viru

s /M

BV

, I

HH

NV

, C

iliop

hora

K

rabs

etsv

e et

al.

(200

4)

Pa

laem

on s

pp.

D

ata

gap

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

29

2

w

ww

.dig

sfis

h.co

m

Sci

entif

ic n

ame

D

isea

se a

gen

t R

efer

ence

s P

ara

pen

aeo

psi

s spp

.

Dat

a ga

p P

ena

eus

mo

no

do

n IH

HN

V,

LPV

, H

aem

ocyt

ic r

od s

hap

ed v

irus,

MoV

, M

BV

, G

AV/LO

V,

SM

V,

HP

V,

BM

NV

-like

, M

yco

pla

sma s

pp.,

Aca

ntho

cep

hala

, B

act

eria

l dis

ease

, C

esto

da,

Cili

op

hora

, Fun

gal d

isea

se,

Gre

garin

es,

Nem

ato

da

Ow

ens

et

al.

(199

2,

1998

),

Spa

nn

et

al.

(199

7a),

O

’Don

oghu

e a

nd

Adl

ard

(2

000)

, B

iose

curit

y A

ustr

alia

(20

06, 2

009)

Pen

aeu

s es

cule

ntu

s

IHH

NV

, LP

V,

Ha

emoc

ytic

rod

sha

ped

viru

s, H

PV

, M

BV

, G

AV

/LO

V,

Th

elo

ha

nia

spp.

, Am

eso

n sp

p., A

cant

hoce

pha

la,

Ces

toda

, G

rega

rine

s, N

ema

toda

O

wen

s et

al.

(199

2),

Spa

nn e

t a

l. (1

997b

),

O’D

onog

hue

and

Adl

ard

(20

00),

Bio

secu

rity

Aus

tra

lia (

2006

, 200

9)

Pen

aeu

s se

mis

ulc

atu

s M

BV

, A

mes

on s

pp., A

gm

aso

ma s

pp., T

hel

oh

an

ia sp

p.,

Aca

ntho

cep

hala

, C

esto

da,

Gre

garin

es,

Nem

ato

da

O’D

onog

hue

and

Adl

ard

(20

00),

Bio

secu

rity

Aus

tra

lia (

2006

, 200

9)

Pen

aeu

s m

erg

uie

nsi

s LP

V,

SM

V,

Ha

emoc

ytic

rod

sha

ped

viru

s, M

BV

, G

AV

/LO

V,

HPV

, Th

elo

ha

nia s

pp., A

mes

on

spp,

Aca

ntho

cep

hala

, C

esto

da,

Ba

cter

ial d

isea

se,

fung

al d

isea

se,

Gre

garin

es,N

ema

toda

R

ouba

l et

a

l. (1

989a

),

O’D

onog

hue

and

A

dla

rd (

2000

), L

a F

auc

e et

al.

(200

7),

La

Fa

uce

and

Ow

ens

(200

7, 2

008)

P

ena

eus

ple

bej

us

Ple

bej

us b

acu

lovi

rus/

MB

V,

Cili

opho

ra

Lest

er e

t a

l. (1

987)

, O

’Don

oghu

e a

nd A

dla

rd

(200

0)

Mel

icer

tus (

Pen

aeu

s) la

tisu

lca

tus

MB

V,

BM

NV

-like

, di

gene

an

met

ace

rca

riae,

A

gm

aso

ma

pen

aei, T

hel

oh

ani

a spp

., ot

her

mic

rosp

orid

ia,

Aca

ntho

cep

hala

, C

esto

da,

Gre

garin

es,

Nem

ato

da

O’D

onog

hue

and

Adl

ard

(20

00),

Bio

secu

rity

Aus

tra

lia (

2006

, 200

9)

Pen

aeu

s spp

. M

BV

, G

AV

/LO

V ,

HP

V,

MoV

, A

cant

hoce

pha

la,

Ba

cter

ial d

isease

, C

esto

da,

Cili

opho

ra,

Cru

sta

cea

, fu

nga

l dis

ease

, G

rega

rines

, M

icro

spor

idia

, Ne

ma

toda

S

pann

et

a

l. (1

997b

),

O’D

onog

hue

and

A

dla

rd (

2000

), H

udso

n et

al.

(200

1),

Sel

lars

et

a

l. (2

005)

, B

iose

curit

y A

ustr

alia

(2

006,

20

09)

C

rabs

Ca

rcin

us

ma

enu

s N

ema

toda

, C

esto

da

Gur

ney

et a

l. (2

004)

, Zet

lmei

sl e

t a

l. (2

011)

L

epto

gra

psu

s spp

., P

ara

gra

psu

s spp

.

Dat

a ga

p

Mic

tyri

s lo

ngi

carp

us,

M.

pla

tych

eles

spp.

Dat

a ga

p

Ocy

po

de s

pp.

D

ata

gap

Pla

gu

sia

ch

ab

rus,

Pla

gu

sia

spp.

Dat

a ga

p

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

29

3

w

ww

.dig

sfis

h.co

m

Sci

entif

ic n

ame

D

isea

se a

gen

t R

efer

ence

s P

ort

un

us

pel

ag

icu

s H

ema

tod

iniu

m sp

p., S

acc

ulin

a spp

., Car

cin

on

emer

tes

mits

uku

rii

, H

PV

, Am

eso

n spp

., T

hel

oh

an

ia sp

p., C

opep

oda

, C

rust

ace

a, C

esto

da,

Cili

opho

ra, o

ther

Mic

rosp

orid

ia, T

urbe

llaria

, O

cto

lasm

is sp

., Ca

rcin

on

emer

tes s

pp.

Shi

elds

(1

992)

, S

hiel

ds

and

W

ood

(199

3),

O’D

onog

hue

and

Adl

ard

(20

00),

La

Fa

uce

and

Ow

ens

(200

7)

Scy

lla s

erra

ta H

ema

tod

iniu

m sp

p.,

HP

V, S

cylla

ba

culo

viru

s (S

BV

), u

nide

ntifi

ed t

hra

usto

chyt

rid,

Cru

sta

cea

, C

esto

da,

Cili

opho

ra, Oct

ola

smis s

p.

And

erso

n a

nd

Prio

r (1

992)

, H

udso

n a

nd

Lest

er

(199

4),

O’D

onog

hue

and

A

dla

rd

(200

0),

Kvi

nged

al

et a

l. (

2006

), L

a F

auc

e a

nd O

wen

s (2

007)

, O

wen

s et

al.

(201

0)

S

altw

ater

yab

bies

/ ni

pper

s

Biff

ari

us a

ren

osu

s

D

ata

gap

Ca

llia

nass

a (T

ryp

ea)

au

stra

lien

sis

D

ata

gap

Up

og

ebia

spp.

Dat

a ga

p

Fre

shw

ater

cra

yfis

h

Ch

era

x d

estr

uct

or

Che

rax

dest

ruct

or b

acil

lifo

rm v

iru

s (C

dB

V),

Che

rax

dest

ruct

or s

yst

emic

par

vo

-lik

e v

iru

s (C

dS

PV

), T

hel

oh

an

ia sp

p., P

leis

top

ho

ra sp

p., o

ther

Mic

rosp

orid

ia,

Cili

opho

ra, T

rem

ato

da,

C

esto

da,

Tem

noce

pha

la,

Ost

raco

da, P

olyc

haet

es

Eva

ns e

t a

l. (1

998)

, O

’Don

oghu

e a

nd A

dla

rd

(200

0),

Edg

erto

n et

al.

(200

2)

Ch

era

x

qu

ad

rica

rin

atu

s S

MV

, Che

rax

quad

rica

rin

atu

s b

acil

lifo

rm v

iru

s (C

qB

V),

Che

rax

bacu

lovi

rus

(CB

V)

Gia

rdia

viru

s-lik

e vi

rus,

Reo

viru

s, P

arv

oviru

s (C

qPV

), H

PV

, P

soro

sper

miu

m spp

., Th

elo

han

ia sp

p., o

ther

Mic

rosp

orid

ia, Vib

rio

mim

icu

s, E

sch

eric

hia

co

li, E

nter

ob

act

er in

term

ediu

m,

Aer

om

on

as

hyd

rop

hila

, C

itro

ba

cter

freu

nd

ii, C

oxi

ella

ch

era

xi, R

icke

ttsi

a-li

ke o

rga

nism

s,

Cili

opho

ra, T

rem

ato

da,

Ces

toda

, Tem

noce

pha

la

Her

ber

t (1

987)

, A

nder

son

and

Prio

r (1

992)

, E

ave

s a

nd

Ket

tere

r (1

994)

, E

vans

et

a

l. (1

998)

, O

wen

s a

nd

McE

lnea

(2

000)

, O

’Don

oghu

e a

nd A

dla

rd (

2000

), E

dger

ton

et

al.

(200

2),

Bow

ate

r et

al.

(200

2),

La F

auc

e a

nd O

wen

s (2

007)

C

her

ax

ten

uim

an

us, C

. ca

inii

Ba

cilli

form

viru

ses,

Pso

rosp

erm

ium s

pp, T

hel

oh

an

ia sp

p., V

avr

aia

spp

., o

ther

Mic

rosp

orid

ia,

Pso

rosp

erm

ium s

pp .,

Tre

ma

toda

, N

ema

toda

, O

stra

coda

, Tem

noce

pha

la

Eva

ns e

t a

l. (1

998)

, Edg

erto

n et

al.

(200

2)

Ch

era

x spp

. C

iliop

hora

, Th

elo

ha

nia

spp.

, Va

vra

ia s

pp .,

oth

er M

icro

spor

idia

, Pso

rosp

erm

ium s

pp.,

Cili

opho

ra, T

rem

ato

da,

Nem

ato

da,

Olig

ocha

etes

, O

stra

cod

a,

Pol

ycha

etes

, T

emno

cep

hala

E

vans

et

al.

(199

8),

O’D

onog

hue

and

Adl

ard

(2

000)

, E

dger

ton

et a

l. (2

002)

, E

ng

aeu

s spp

. C

iliop

hora

, Tem

noce

pha

la

Jone

s a

nd L

este

r (1

993)

, O

’Don

oghu

e a

nd

Adl

ard

(20

00)

Eu

ast

acu

s spp

. T

emno

cep

hala

S

ewel

l and

Ca

nnon

(19

98)

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

29

4

w

ww

.dig

sfis

h.co

m

Sci

entif

ic n

ame

D

isea

se a

gen

t R

efer

ence

s S

altw

ater

cr

ayfis

h/lo

bste

rs

P

an

ulir

us

cyg

nu

s B

act

era

l dis

ease

, fu

nga

l dis

ease

, M

icro

spor

idia

, T

rem

ato

da

O’D

onog

hue

and

Adl

ard

(20

00),

Shi

elds

et

al.

(200

6)

Pa

nu

liru

s sp

p.

Ba

cter

al d

isea

se,

Ces

toda

, Cop

epod

a,

Fun

gal d

isea

se,

Mic

rosp

orid

ia,T

rem

ato

da

O’D

onog

hue

and

Adl

ard

(20

00),

Shi

elds

et

al.

(200

6)

Jasu

s ed

wa

rdsi

i B

act

era

l dis

ease

, C

opep

oda

, F

unga

l dis

ease

Shi

elds

et

al.

(200

6)

Jasu

s (S

ag

ma

ria

sus)

verr

eau

xi

Ba

cter

al d

isea

se,

Cop

epod

a,

Fun

gal d

isea

se

Shi

elds

et

al.

(200

6)

M

OLL

US

CS

Gas

trop

ods

C

ella

na

spp.

Dat

a ga

p H

alio

tis la

evig

ata

A

balo

ne V

iral G

ang

lione

uriti

s,

Per

kin

sus

ols

eni, R

icke

ttsi

a-li

ke o

rga

nism

s,

Po

lyd

ora

spp.

, B

occ

ard

ia s

pp.,

Ba

cter

ial d

isea

se,

Cili

opho

ra, F

unga

l dis

ease

, T

rem

ato

da

O’D

onog

hue

and

Adl

ard

(20

00),

Ha

ndlin

ger

et a

l. (2

006)

, H

oop

er e

t a

l (20

07)

Ha

liotis

ru

bra

A

balo

ne V

iral G

ang

lione

uriti

s, Per

kin

sus

ols

eni, R

icke

ttsi

a-li

ke o

rga

nism

s,

Po

lyd

ora

spp.

, B

occ

ard

ia s

pp.,

Ba

cter

ial d

isea

se,

Cili

opho

ra, F

unga

l dis

ease

, T

rem

ato

da

O’D

onog

hue

and

Adl

ard

(20

00),

Ha

ndlin

ger

et a

l. (2

006)

, H

oop

er e

t a

l. (2

007)

H

alio

tis s

pp.

Per

kin

sus

ols

eni, R

icke

ttsi

a-l

ike

orga

nism

s,

Po

lyd

ora

spp.

, Bo

cca

rdia

spp.

,Ba

cter

ial d

isea

se,

Cili

opho

ra, F

unga

l dis

ease

, T

rem

ato

da

O’D

onog

hue

and

Adl

ard

(20

00),

Ha

ndlin

ger

et a

l. (2

006)

Biv

alve

s

Am

usi

um

spp.

, Pec

ten

spp.

P

erki

nsu

s spp

., D

iges

tive

epith

elia

l viro

sis,

Ces

toda

, Tre

ma

toda

O

’D

onog

hue

and

Adl

ard

(20

00),

AF

FA

(2

002)

C

rass

ost

rea

gig

as

Po

lyd

ora

spp.

, Bo

cca

rdia

spp.

, Ric

kett

sia

-like

org

ani

sms,

Uni

dent

ified

mic

roce

ll, V

iral

gam

etoc

ytic

hyp

ertr

ophy

, B

act

eria

l inf

ectio

ns,

Cop

epod

a,

Cili

opho

ra

Lest

er (

1989

), D

iggl

es (

2003

)

F. H

yriid

ae

Dat

a ga

p K

ate

lysi

a spp

., A

na

da

ra s

pp.

Per

kin

sus s

pp.

O’D

onog

hue

and

A

dla

rd (

2000

), H

ine

and

T

horn

e (2

000)

M

ytilu

s ed

ulis

. C

opep

oda

, T

rem

ato

da,

Ste

inh

ausi

a m

ytilo

vum, P

oly

do

ra sp

p., B

occ

ard

ia sp

p., P

inn

oth

eres

spp.

P

rege

nzer

(19

83),

Jon

es a

nd C

reep

er (

2006

) P

inn

a b

ico

lor

Viru

s-lik

e in

clus

ions

, R

icke

ttsi

a-li

ke o

rga

nism

s, C

esto

da,

Cili

opho

ra

Hin

e a

nd T

horn

e (2

000)

P

leb

ido

na

x d

elto

ides

Non

e fo

und

B

ott

et a

l. (2

005)

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

29

5

w

ww

.dig

sfis

h.co

m

Sci

entif

ic n

ame

D

isea

se a

gen

t R

efer

ence

s S

acc

ost

rea

cu

ccu

lata

Ha

plos

por

idia

, Ma

rtei

lia s

p, V

irus-

like

incl

usio

ns, Per

kin

sus s

p., R

icke

ttsi

a-li

ke o

rga

nism

s,

Cili

opho

ra, T

rem

ato

da,

Nem

ato

da,

greg

arin

e H

ine

and

Tho

rne

(200

0, 2

002)

, B

earh

am

et

al.

2008

a, 2

008c

, 20

09

Sa

cco

stre

a g

lom

era

ta M

art

eilia

syd

ney

i, Ma

rtei

lioid

es b

ran

cha

lis, P

erki

nsu

s sp.

, Bo

na

mia

ro

ug

hley

i, Po

lyd

ora

h

op

lura

, Ste

inh

ausi

a-like

mic

rosp

orid

ian,

Ric

kett

sia

-like

org

ani

sms,

Ces

toda

, C

iliop

hora

, H

apl

osp

orid

ia,

Tre

ma

toda

Lest

er (

1989

), A

nder

son

et a

l. (1

995)

, H

ine

and

Tho

rne

(200

0),

O’D

onog

hue

and

Adl

ard

(2

000)

, C

oche

nnec

–La

urea

u et

al.

(200

3)

Cep

halo

pods

Lo

ligo

spp.

Dat

a ga

p N

oto

da

rus

go

uld

i

Dat

a ga

p N

oto

da

rus s

pp.

D

ata

gap

Oct

op

us

au

stra

lis

Dat

a ga

p O

cto

pu

s cy

an

ea

Dat

a ga

p O

cto

pu

s m

ao

rum

D

ata

gap

Oct

op

us

pa

llid

us

D

ata

gap

Oct

op

us t

etri

cus

D

ata

gap

Oct

op

us s

pp

D

ata

gap

Sep

ia a

pa

ma

D

ata

gap

Sep

ia r

oze

lla

D

ata

gap

Sep

ia sp

p

Dat

a ga

p S

epio

teu

this

au

stra

lis

Dat

a ga

p S

epio

teu

this

less

on

iana

D

ata

gap

A

NN

ELI

DS

O

ligoc

haet

es

F

. Aca

ntho

drili

dae

Dat

a ga

p E

isen

ia f

oet

ida

D

ata

gap

Lu

mb

ricu

s re

bel

lus

D

ata

gap

F. L

umbr

icid

ae

D

ata

gap

F

. Meg

asc

olec

ida

e

Dat

a ga

p F

. Oct

ocha

etid

ae

D

ata

gap

Per

ion

yx e

xca

vatu

s

Dat

a ga

p

___

___

____

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

___

___

____

___

____

___

____

___

___

____

___

____

____

___

___

____

___

____

___

____

___

___

____

___

____

__

___

___

____

___

___

____

___

_

F

RD

C P

roje

ct N

o. 2

009/

072,

Fin

al R

epor

t Ju

ly 2

011

29

6

w

ww

.dig

sfis

h.co

m

Sci

entif

ic n

ame

D

isea

se a

gen

t R

efer

ence

s P

oly

chae

tes

A

ust

ralo

ner

eis

ehle

rsi

D

ata

gap

Au

stra

lon

up

his

pa

rate

res

D

ata

gap

Au

stra

lon

up

his

tere

s

Dat

a ga

p D

iop

atr

a d

enta

ta,

D.

aci

cula

ta

D

ata

gap

F. E

unic

ida

e

Dat

a ga

p G

lyce

ra o

vig

era

D

ata

gap

F. G

lyce

rida

e

Dat

a ga

p M

arp

hys

a s

an

gu

inea

D

ata

gap

Nep

hty

s a

ust

ralie

nsi

s, P

erin

erei

s n

un

tia M

art

eilia

syd

ney

i A

dla

rd a

nd N

ola

n (2

009)

, C

ribb

(201

0)

F. N

epht

yida

e M

art

eilia

syd

ney

i A

dla

rd a

nd N

ola

n (2

009)

, C

ribb

(201

0)

F. N

erei

dida

e

Dat

a ga

p H

irsu

ton

up

his

gyg

is

Dat

a ga

p H

irsu

ton

up

his

ma

ria

hir

stu

a

D

ata

gap

On

up

his

taen

iata

D

ata

gap

E

CH

INO

DE

RM

S

C

entr

ost

eph

an

us

rod

ger

sii

D

ata

gap

Hel

ico

cid

ari

s sp

p.

Dat

a ga

p H

olo

pn

eust

es p

ycn

otil

us

D

ata

gap

Tri

pn

eust

es g

ratil

la

D

ata

gap

A

SC

IDE

AN

S

P

yura

sto

lon

ifera

D

ata

gap

_________________________________________________________________________________________________________________________________________________________________________________


297

www.digsfish.com

Appendix 2. Example of educational posters employed by Michigan Department of Natural Resources as a form of risk mitigation to help control spread of VHS virus in the great lakes region of the USA.

Date post:	28-Dec-2021
Category:	Documents
Upload:	others
View:	3 times
Download:	1 times

RISK ANALYSIS – AQUATIC ANIMAL DISEASES ASSOCIATED …

Documents