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
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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.
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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
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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
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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
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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
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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.
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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
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Commodity type Disease agent requiring risk management
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.
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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
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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
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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
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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
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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
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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.
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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
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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
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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
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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
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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.
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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).
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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.
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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.
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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
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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
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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
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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.
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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),
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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.
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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
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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%)
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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
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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).
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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
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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
Kewagama Research (2002) estimated that around 4.7% of respondents throughout all states in the country
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)
Kewagama Research (2002) estimated that around 14.8% of respondents throughout all states in the country
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
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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
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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)
Kewagama Research (2002) estimated that around 29.5% of respondents throughout all states in the country
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
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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
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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
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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
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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
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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
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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
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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
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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.
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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
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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 � �
___
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____
___
____
___
___
____
___
____
___
____
___
___
____
___
____
___
___
____
___
____
___
____
___
___
____
___
____
____
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_
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
_________________________________________________________________________________________________________________________________________________________________________________
FRDC Project No. 2009/072, Final Report July 2011
52
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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).
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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
_________________________________________________________________________________________________________________________________________________________________________________
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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
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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).
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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
Low risk Moderate risk
High risk Extreme risk
Negligible risk
Negligible risk
Very low risk
Low risk Moderate risk
High risk
Negligible risk
Negligible risk
Negligible risk
Very low risk
Low risk Moderate 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
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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.
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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
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epor
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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
.
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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.
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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
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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.
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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.
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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
be considered further in this RA.
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
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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
be considered further in this RA.
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
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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
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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) � � � �
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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) � � � � �
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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).
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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).
5.1.5 Release assessment
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.
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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
5.1.6 Exposure assessment
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
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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.
5.1.7 Consequence assessment
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
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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.
5.1.8 Risk estimation
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
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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).
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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).
5.2.5 Release assessment
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.
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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
5.2.6 Exposure assessment
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.
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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.
5.2.7 Consequence assessment
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.
5.2.8 Risk estimation
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.
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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
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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.
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5.3.5 Release assessment
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
5.3.6 Exposure assessment
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
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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.
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5.3.7 Consequence assessment
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.
5.3.8 Risk estimation
The unrestricted risk associated with nodavirus 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
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
likelihood of release
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
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5.4 Emergence of a new previously unknown virus of finfish
5.4.1 Aetiologic agent: A previously undescribed virus that infects finfish.
5.4.2 OIE List: No NACA List: No
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.
5.4.5 Release assessment
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
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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
5.4.6 Exposure assessment
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.
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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.
5.4.7 Consequence assessment
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.
5.4.8 Risk estimation
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.
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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
likelihood of release
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
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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.
5.5.2 OIE List: No NACA List: No
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).
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5.5.5 Release assessment
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
commodities are listed below.
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
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5.5.6 Exposure assessment
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
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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.
5.5.7 Consequence assessment
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.
5.5.8 Risk estimation
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.
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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
likelihood of release
and exposure
Moderate Moderate Low Low Low Low
Consequences of
establishment
Low Low Low Low Low Low
Risk estimation Low Risk Low Risk Very Low
Risk
Very Low
Risk
Very Low
Risk
Very Low
Risk
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5.6 Infection of salmonids with Lactococcus garvieae
5.6.1 Aetiologic agent: Lactococcus garvieae, a gram positive bacterium in the Family Streptococcaceae.
5.6.2 OIE List: No NACA List: No
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).
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5.6.5 Release assessment
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
commodities are listed below.
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
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5.6.6 Exposure assessment
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.
5.6.7 Consequence assessment
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
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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.
5.6.8 Risk estimation
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
Low Low Low Low Low Low
Risk estimation Very low
risk
Very low
risk
Very low
risk
Negligible
risk
Negligible
risk
Very low
risk
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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.
5.7.2 OIE List: No NACA List: No
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).
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5.7.5 Release assessment
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
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5.7.6 Exposure assessment
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.
5.7.7 Consequence assessment
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
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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.
5.7.8 Risk estimation
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 estimation Very low
risk
Negligible
risk
Negligible
risk
Negligible
risk
Negligible
risk
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5.8 Infection of finfish with Yersinia ruckeri (Yersiniosis)
5.8.1 Aetiologic agent: Yersinia ruckeri, a gram negative bacterium within the family Enterobacteriaceae.
5.8.2 OIE List: No NACA List: No
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).
5.8.5 Release assessment
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
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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
5.8.6 Exposure assessment
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
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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.
5.8.7 Consequence assessment
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
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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.
5.8.8 Risk estimation
The unrestricted risk associated with Yersinia ruckeri 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
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
likelihood of release
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
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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.
5.9.2 OIE List: Yes NACA List: Yes
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
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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).
5.9.5 Release assessment
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
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5.9.6 Exposure assessment
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.
5.9.7 Consequence assessment
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
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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.
5.9.8 Risk estimation
The unrestricted risk associated with Aphanomyces invadans 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 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
likelihood of release
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
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5.10 Infection of finfish with Microsporidians
5.10.1 Aetiologic agent: Intracellular protozoan parasites of fishes from the Class Microsporea.
5.10.2 OIE List: No NACA List: No
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).
5.10.5 Release assessment
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,
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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
5.10.6 Exposure assessment
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
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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.
5.10.7 Consequence assessment
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.
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5.10.8 Risk estimation
The unrestricted risk associated with microsporidian infection 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 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
Moderate Moderate Low Low Low Low
Consequences of
establishment
Low Low Low Low Low Low
Risk estimation Low Risk Low Risk Very Low
Risk
Very Low
Risk
Very Low
Risk
Very Low
Risk
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5.11 Systemic amoebic infections of goldfish
5.11.1 Aetiologic agent: Amoebae-like protozoa that infect goldfish and other finfish.
5.11.2 OIE List: No NACA List: No
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).
5.11.5 Release assessment
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
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prevalence of the disease agent, the likelihood estimations for the occurrence of amoebae in these
commodities are listed below.
Release assessment for systemic amoebic infections of goldfish
Commodity type Live
goldfish
Whole fresh
dead goldfish
Likelihood of release High Moderate
5.11.6 Exposure assessment
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.
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5.12 Infection of finfish with scuticociliates (including Uronema spp.)
5.12.1 Aetiologic agent: Scuticociliates (Protozoa: Ciliophora) that infect finfish.
5.12.2 OIE List: No NACA List: No
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.
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5.12.5 Release assessment
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
commodities are listed below.
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
5.12.6 Exposure assessment
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
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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.
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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).
5.13.5 Release assessment
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
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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
5.13.6 Exposure assessment
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
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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
considered to be Low.
5.13.7 Consequence assessment
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.
5.13.8 Risk estimation
The unrestricted risk associated with monogeneans 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
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
likelihood of release
and exposure
Low Very low Negligible Negligible Negligible Negligible
Consequences of
establishment
Low Low Low Low Low Low
Risk estimation Very
low risk
Negligible
risk
Negligible
risk
Negligible
risk
Negligible
risk
Negligible
risk
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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.
5.14.2 OIE List: No NACA List: No
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).
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5.14.5 Release assessment
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
commodities are listed below.
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
High Very low Negligible Negligible Negligible Negligible
5.14.6 Exposure assessment
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
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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.
5.14.7 Consequence assessment
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
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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.
5.14.8 Risk estimation
The unrestricted risk associated with digeneans 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
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
likelihood of release
and exposure
Low Very Low Negligible Negligible Negligible Negligible
Consequences of
establishment
Moderate Moderate Moderate Moderate Moderate Moderate
Risk estimation Low risk Negligible
risk
Negligible
risk
Negligible
risk
Negligible
risk
Negligible
risk
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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.
5.15.2 OIE List: No NACA List: No
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
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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.
5.15.5 Release assessment
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.
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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
High Very low Negligible Negligible Negligible Negligible
5.15.6 Exposure assessment
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
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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.
5.15.7 Consequence assessment
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.
5.15.8 Risk estimation
The unrestricted risk associated with nematodes and cestodes 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 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.
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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
Low Very Low Negligible Negligible Negligible Negligible
Consequences of
establishment
Moderate Moderate Moderate Moderate Moderate Moderate
Risk estimation Low risk Negligible
risk
Negligible
risk
Negligible
risk
Negligible
risk
Negligible
risk
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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.
5.16.2 OIE List: No NACA List: No
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).
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5.16.5 Release assessment
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
High Very low Negligible Negligible Negligible Negligible
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5.16.6 Exposure assessment
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.
5.16.7 Consequence assessment
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
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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.
5.16.8 Risk estimation
The unrestricted risk associated with copepods 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
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
likelihood of release
and exposure
Low Very Low Negligible Negligible Negligible Negligible
Consequences of
establishment
Moderate Moderate Moderate Moderate Moderate Moderate
Risk estimation Low risk Negligible
risk
Negligible
risk
Negligible
risk
Negligible
risk
Negligible
risk
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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.
5.17.2 OIE List: No NACA List: No
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).
5.17.5 Release assessment
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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/
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5.17.6 Exposure assessment
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.
5.17.7 Consequence assessment
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
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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.
5.17.8 Risk estimation
The unrestricted risk associated with myxosporeans 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
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
Moderate Moderate Low Low Low Low
Consequences of
establishment
Low Low Low Low Low Low
Risk estimation Low risk Low risk Very low
risk
Very low
risk
Very low
risk
Very low
risk
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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).
5.18.2 OIE List: No NACA List: No
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).
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5.18.5 Release assessment
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
5.18.6 Exposure assessment
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
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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.
5.18.7 Consequence assessment
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
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introduction of crayfish viruses into different parts of the Australian environment via use of infected bait
would likely be Moderate.
5.18.8 Risk estimation
The unrestricted risk associated with freshwater crayfish viruses 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 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
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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.
5.19.2 OIE List: Yes NACA List: Yes
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
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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).
5.19.5 Release assessment
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
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 (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
commodities are listed below.
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
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5.19.6 Exposure assessment
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.
5.19.7 Consequence assessment
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
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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.
5.19.8 Risk estimation
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
Moderate Moderate Moderate Moderate Moderate
Risk estimation Low risk Low risk Very low risk Very low
risk
Very low risk
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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).
5.20.2 OIE List: No NACA List: No
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).
5.20.5 Release assessment
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
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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
5.20.6 Exposure assessment
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
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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.
5.20.7 Consequence assessment
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.
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5.20.8 Risk estimation
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 estimation Very low
risk
Very low risk Very low risk Very low
risk
Very low risk
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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).
5.21.2 OIE List: Yes NACA List: Yes
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
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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.
5.21.5 Release assessment
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
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 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
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5.21.6 Exposure assessment
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.
5.21.7 Consequence assessment
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
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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.
5.21.8 Risk estimation
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 estimation Very low
risk
Very low risk Very low risk Very low
risk
Very low risk
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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.
5.22.2 OIE List: No NACA List: No
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,
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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.
5.22.5 Release assessment
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
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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
5.22.6 Exposure assessment
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
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 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
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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.
5.22.7 Consequence assessment
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.
5.22.8 Risk estimation
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.
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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 estimation Negligible
risk
Negligible risk Negligible risk Negligible
risk
Negligible risk
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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).
5.23.2 OIE List: No NACA List: No
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
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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.
5.23.5 Release assessment
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
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). 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
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5.23.6 Exposure assessment
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.
5.23.7 Consequence assessment
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.
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5.23.8 Risk estimation
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
Very low Very low Very low Very low Very low
Risk estimation Negligible
risk
Negligible risk Negligible risk Negligible
risk
Negligible risk
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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).
5.24.2 OIE List: No NACA List: No
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
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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.
5.24.5 Release assessment
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
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
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). 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
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5.24.6 Exposure assessment
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
particles were introduced into an area where susceptible prawns were present under suitable environmental
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
considered to be Low.
5.24.7 Consequence assessment
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
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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.
5.24.8 Risk estimation
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
Moderate Moderate Moderate Moderate Moderate
Risk estimation Low risk Low risk Very low risk Very low
risk
Very low risk
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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).
5.25.2 OIE List: Yes NACA List: Yes
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).
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5.25.5 Release assessment
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
5.25.6 Exposure assessment
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.
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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.
5.25.7 Consequence assessment
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.
5.25.8 Risk estimation
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.
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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
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
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5.26 Infection of crustaceans with rickettsia-like organisms (RLOs)
5.26.1 Aetiologic agent: Rickettsia –like and chlamydia-like organisms that infect crustaceans.
5.26.2 OIE List: Yes NACA List: Yes
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
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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).
5.26.5 Release assessment
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
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. 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
5.26.6 Exposure assessment
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
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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.
5.26.7 Consequence assessment
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.
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5.26.8 Risk estimation
The unrestricted risk associated with crustacean RLOs 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 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
Low Low Very low Very low Very low
Consequences of
establishment
Very low Very low Very low Very low Very low
Risk estimation Negligible
risk
Negligible risk Negligible risk Negligible
risk
Negligible risk
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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).
5.27.2 OIE List: No NACA List: No
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
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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.
5.27.5 Release assessment
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
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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
5.27.6 Exposure assessment
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,
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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.
5.27.7 Consequence assessment
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
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of introduction of Hematodinium spp. into different parts of the Australian environment via use of infected
bait would likely be Moderate.
5.27.8 Risk estimation
The unrestricted risk associated with Hematodinium spp. 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 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
Low Low Very low Very low Very low
Consequences of
establishment
Moderate Moderate Moderate Moderate Moderate
Risk estimation Low Risk Low Risk Negligible risk Negligible
risk
Negligible risk
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5.28 Infection of crustaceans with Microsporidians
5.28.1 Aetiologic agent: Thelohania spp., and other parasites of the Phylum Microsporea that infect
crustaceans.
5.28.2 OIE List: No NACA List: No
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).
5.28.5 Release assessment
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
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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
commodities are listed below.
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
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5.28.6 Exposure assessment
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.
5.28.7 Consequence assessment
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
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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.
5.28.8 Risk estimation
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 estimation Low risk Low risk Very low risk Very low
risk
Very low risk
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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.
5.29.2 OIE List: No NACA List: No
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).
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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.
5.29.5 Release assessment
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
5.29.6 Exposure assessment
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
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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.
5.29.7 Consequence assessment
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.
5.29.8 Risk estimation
The unrestricted risk associated with rhizocephalan parasites 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 rhizocephalan parasites exceeds the ALOP only for live crabs, suggesting that additional risk
management for these disease agents is required for these commodities.
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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
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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).
5.30.2 OIE List: Yes NACA List: Yes
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
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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.
5.30.5 Release assessment
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
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5.30.6 Exposure assessment
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.
5.30.7 Consequence assessment
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
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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.
5.30.8 Risk estimation
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
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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.
5.31.2 OIE List: Yes NACA List: Yes
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.
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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).
5.31.5 Release assessment
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
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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
commodities are listed below.
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
5.31.6 Exposure assessment
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.
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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.
5.31.7 Consequence assessment
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.
5.31.8 Risk estimation
The unrestricted risk associated with Bonamia spp. 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
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.
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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
Moderate Moderate Moderate Moderate Moderate
Risk estimation Moderate
risk
Low risk Negligible
risk
Negligible
risk
Negligible
risk
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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.
5.32.2 OIE List: No NACA List: No
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
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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).
5.32.5 Release assessment
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.
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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
5.32.6 Exposure assessment
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
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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.
5.32.7 Consequence assessment
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
environment via use of infected bait would likely be Moderate.
5.32.8 Risk estimation
The unrestricted risk associated with haplosporidians 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
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
Moderate Moderate Moderate Moderate Moderate
Risk estimation Low risk Very low
risk
Negligible
risk
Negligible
risk
Negligible
risk
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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.
5.33.2 OIE List: No NACA List: No
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
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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).
5.33.5 Release assessment
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.
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.)
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
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5.33.6 Exposure assessment
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.
5.33.7 Consequence assessment
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
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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.
5.33.8 Risk estimation
The unrestricted risk associated with M. sydneyi 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
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
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5.34 Infection of molluscs with Perkinsus olseni
5.34.1 Aetiologic agent: Perkinsus olseni and other parasitic protozoa within the Family Perkinsidae.
5.34.2 OIE List: Yes NACA List: Yes
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
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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).
5.34.5 Release assessment
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.
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.). 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
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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
5.34.6 Exposure assessment
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
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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.
5.34.7 Consequence assessment
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
environment via use of infected bait would likely be Moderate.
5.34.8 Risk estimation
The unrestricted risk associated with Perkinsus olseni 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
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
Moderate Moderate Moderate Moderate Moderate
Risk estimation Moderate Risk Moderate Risk Moderate Risk Moderate Risk Negligible
risk
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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.
5.35.2 OIE List: No NACA List: No
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
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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).
5.35.5 Release assessment
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
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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
5.35.6 Exposure assessment
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
_________________________________________________________________________________________________________________________________________________________________________________
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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.
5.35.7 Consequence assessment
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.
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5.35.8 Risk estimation
The unrestricted risk associated with spionid mudworm infections 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 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
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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.
___
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___
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___
____
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___
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___
____
___
___
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_
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
_________________________________________________________________________________________________________________________________________________________________________________
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Table 12. 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 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
_________________________________________________________________________________________________________________________________________________________________________________
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Commodity type Disease agent requiring risk management
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.
_________________________________________________________________________________________________________________________________________________________________________________
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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.
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ma
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., C
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___
___
____
___
____
___
____
___
___
____
___
____
___
____
___
___
____
___
____
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____
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____
___
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___
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____
___
____
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___
___
____
___
____
___
____
___
___
____
___
____
__
___
___
____
___
___
____
___
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F
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Fin
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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
_________________________________________________________________________________________________________________________________________________________________________________
FRDC Project No. 2009/072, Final Report July 2011
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.