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Supplementary Import Risk Analysis: Frozen fish and cephalopod molluscs for fish bait

Approved for IHS development

Prepared for Ministry for Primary Industries By the Animal Risk Assessment Team, Risk Assessment Group, Animal and Plant Health Directorate, Biosecurity New Zealand, Ministry for Primary Industries ISBN No:978-1-004381-9 (online) November 2020

Disclaimer

While every effort has been made to ensure the information in this publication is accurate, the Ministry for Primary Industries does not accept any responsibility or liability for error of fact, omission, interpretation or opinion that may be present, nor for the consequences of any decisions based on this information. Cover photo: Atlantic herring (Clupea harengus) (Photo credit: NOAA, Wikimedia Commons) Recommended citation: MPI (2020) Supplementary Import Risk Analysis: Frozen fish and cephalopod molluscs for fish bait. Ministry for Primary Industries, New Zealand. This publication is available on the Ministry for Primary Industries website at http://www.mpi.govt.nz/news-and-resources/publications/ © Crown Copyright - Ministry for Primary Industries

http://www.mpi.govt.nz/news-and-resources/publications/

Supplementary Import risk analysis: Frozen fish and cephalopod molluscs for fish bait

Version 4.0

19 November 2020

Approved for IHS development

Enrico Perotti

Associate Director Animal and Plant Health, Risk Assessment Group,

Biosecurity New Zealand

ii Import risk analysis: Frozen fish and cephalopod molluscs for fish bait

Version information

Version number Comments Date of release

1.0 First draft version October 2019

1.1 Internally peer reviewed version January 2020

2.0 Peer-reviewed version March 2020

3.0 Revised version for second peer review May 2020

3.1 Revised version – Extended frozen storage September 2020

4.0 Approved for IHS development November 2020

New Zealand is a member of the World Trade Organisation and a signatory to the Agreement on the Application of Sanitary and Phytosanitary Measures (The Agreement). Under the Agreement, countries must base their measures on an International Standard or an assessment of the biological risks to plant, animal or human health.

This document extends and supplements a previous risk analysis (eviscerated or trunked fish for human consumption) (Blackwell 2019), to examine the risks associated with frozen fish and cephalopod molluscs imported for use as fish bait in commercial and recreational fishing. It provides a scientific analysis of the likelihood of entry, exposure, establishment and spread of various pathogens and diseases in the commodity and assesses the consequences of their establishment in New Zealand. The document has been internally and externally peer reviewed.

Import risk analysis: Frozen fish and cephalopod molluscs for fish bait iii

Contributors to this risk analysis

The following people provided significant input into the development of this risk analysis:

Primary contributor

Ron Blackwell Senior Adviser, Animal Risk Assessment Team, Risk Assessment Group

Biosecurity New Zealand, Ministry for Primary Industries, Wellington, New Zealand

Internal Peer Review

Sudharma Leelawardana (Sue)

Manager, Animal Risk Assessment Team. Risk Assessment Group


Don Leelawardana Senior Adviser, Animal Risk Assessment Team, Risk Assessment Group


Shahid Haneef Senior Adviser, Animal Risk Assessment Team, Risk Assessment Group


Dan Kluza Principal Adviser, Risk Assessment Group


Vicki Melville

Manager, Animal Imports Team 1, Risk Assessment Group


Nasser Ahmed Senior Adviser, Animal Imports Team 1, Risk Assessment Group


External Scientific Review

Ramesh Perera Biosecurity consultant Griffith, ACT, Australia

iv Import risk analysis: Frozen fish and cephalopod molluscs for fish bait

Table of Contents

Index of Tables and Figures vi

Acronyms and abbreviations vii

Executive summary 1

Introduction 3 2.1 The importance of fish bait in New Zealand 5 2.2 The importance of aquaculture 7

Scope, commodity definition and assumptions 9 3.1 Scope 9 3.2 Commodity definition 10 3.3 Assumptions 12

Risk analysis methodology 13 4.1 General procedures 13

Hazard identification 16 5.1 Hazard list for risk assessment 35

General considerations 35 6.1 Risk assessment considerations 35 6.2 Risk management considerations 39

Birnaviridae: Marine aquabirnavirus 43 7.1 Technical review 43 7.2 Risk assessment 47 7.3 Risk management 48

Iridoviridae: Megalocytivirus (red sea bream iridovirus - RSIV) and associated viruses 50 8.1 Technical review 50 8.2 Risk assessment 52 8.3 Risk management 54

Iridoviridae: Erythrocytic necrosis virus (ENV) 55 9.1 Technical review 55 9.2 Risk assessment 57 9.3 Risk management 59

Nodaviridae: Nervous necrosis virus (NNV) 60 10.1 Technical review 60 10.2 Risk assessment 64 10.3 Risk management 66

Orthomyxoviridae: Infectious salmon anaemia virus (ISAV) 67 11.1 Technical review 67 11.2 Risk assessment 70 11.3 Risk management 71

Reoviridae: Piscine aquareovirus (PRV) and associated aquareoviruses 72 12.1 Technical review 72 12.2 Risk assessment 74 12.3 Risk management 75

Rhabdoviridae: Infectious haematopoietic necrosis virus (IHNV) 75 13.1 Technical review 75 13.2 Risk assessment 77 13.3 Risk management 78

Rhabdoviridae: Viral haemorrhagic septicaemia virus (VHSV) 80 14.1 Technical review 80 14.2 Risk assessment 83 14.3 Risk management 85

Import risk analysis: Frozen fish and cephalopod molluscs for fish bait v

Edwardsiella spp. 86 15.1 Technical review 86 15.2 Risk assessment 89 15.3 Risk management 91

Francisella spp. 92 16.1 Technical review 92 16.2 Risk assessment 94 16.3 Risk management 94

Pseudomonas anguilliseptica 94 17.1 Technical review 94 17.2 Risk assessment 97 17.3 Risk management 97

Streptococcus spp. (S. agalactiae serotype III: 283, S. iniae) 97 18.1 Technical review 97 18.2 Risk assessment 101 18.3 Risk management 103

Myxozoan pathogens 104 19.1 Technical review 104 19.2 Risk assessment 110 19.3 Risk management 112

References 113

Appendices 156

vi Import risk analysis: Frozen fish and cephalopod molluscs for fish bait

Index of Tables and Figures

Table 1. Estimated total number of hooks per year set in commercial long lining, by vessel size group, for fishing years (1 Oct to 30 Sept) 2002-03, 2012-13 and 2018-19 5

Table 2. Estimated number of recreational fishing trips using fish bait during 2017-18 6

Table 3. Estimated volume (t) and source location of finfish imported as fish bait, 2018 8

Table 4. Estimated volume (t) of coleoid cephalopod molluscs imported as fish bait, 2018 9

Table 5. Finfish and coleoid cephalopod mollusc families and species considered as fish bait 12

Table 6. Hazard Identification table for specified1 wild marine finfish and coleoid molluscs used as fish bait17

Table 7. Fish bait species susceptible to marine aquabirnavirus (MABV) 45

Table 8. Fish bait species susceptible to red sea bream iridovirus (RSIV), infectious spleen and kidney necrosis virus (ISKNV) and associated iridoviruses 51

Table 9. Fish bait species susceptible to erythrocytic necrosis virus (ENV) 56

Table 10. Fish and coleoid cephalopod mollusc species susceptible to nervous necrosis virus (NNV) 63

Table 11. Fish bait species susceptible to strains of infectious salmon anaemia virus (ISAV) 69

Table 12. Fish bait species susceptible to piscine aquareovirus and related strains 73

Table 13. Fish bait species susceptible to infectious haematopoietic necrosis virus (IHNV) 76

Table 14. Fish bait species susceptible to viral haemorrhagic septicaemia virus (VHSV) 81

Table 15. Fish bait species susceptible to Edwardsiella spp. 88

Table 16. Fish bait species susceptible to Francisella spp. 93

Table 17. Fish bait species susceptible to Pseudomonas anguilliseptica 95

Table 18. Fish bait species susceptible to Streptococcus agalactiae III: 283 99

Table 19. Fish bait species susceptible to Streptococcus iniae 99

Table 20. Fish species susceptible to exotic myxozoan pathogens Enteromyxum leei, Kudoa clupeidae, K. iwatai. K. nova. K. thyrsites 107

Table 21. Taxonomy and geographical distribution of finfish imported as fish bait, 2008-2017 156

Table 22. Classification and distribution of coleoid cephalopod molluscs imported, 2008-2017 158

Table 23. Imports of coleoid cephalopod molluscs, 2013-2015 160

Table 24. Risk organisms and associated fish bait species where specific risk management measures are proposed 161

Table 25. Assumed reduction in pathogen load associated with risk management options 164

Figure 1. The general risk analysis process 14

Figure 2. Potential distribution pathways for frozen fish or coleoid cephalopod mollusc fish bait 38

Import risk analysis: Frozen fish and cephalopod molluscs for fish bait vii

Acronyms and abbreviations

Term/Acronym Definition

Bait fish Whole, frozen, small sized, generally schooling pelagic finfish (e.g. herring, mackerel) imported for use as fish bait.

Berley Minced fish and fish offal that is scattered on the water or on the seabed to attract fish. Berley may be combined with bread, flour and additional attractants such as fish oil. Berley is also known as chum, or ground-bait.

Biosecurity New Zealand

A business unit of the Ministry for Primary Industries, New Zealand.

CFU Colony forming units of bacteria.

CIHEAM International Centre for Advanced Mediterranean Agronomic Studies.

Commercially prepared and packaged

A product that has been manufactured in a commercial manner by a commercial enterprise and is packaged in tamper-proof packaging (MPI 2011).

Competent Authority Government veterinary authority or other Government authority having the responsibility and competence for ensuring or supervising the implementation of animal health and welfare measures, international veterinary certification and other standards and recommendations in the OIE Code and Aquatic Code.

Derived from marine waters

Organisms derived from marine waters only (i.e. excluding species that spend some, or all of their life cycle in fresh water, such as salmonids).

EFSA European Food Safety Authority.

Epizootic An outbreak of a rapidly spreading disease that is temporarily prevalent and widespread in a population.

ESR Environmental Science and Research, Crown Research Institute Ltd., New Zealand.

Eviscerated finfish Bony fish of Family Actinopterygii which have been processed to remove the viscera (internal organs) but where the head and/or gills may be attached.

FAO Food and Agriculture Organisation of the United Nations, Rome, Italy.

Farm sourced All organisms held in confinement at any point in their life cycle, including stock collected from the wild and then on-grown or held in confinement.

Finfish Poikilothermic vertebrates that breathe by gills, including bony fish of Family Actinopterygii and cartilaginous fish of Class Chondrichthyes.

Fish A general term which includes finfish, as well as other vertebrates and invertebrate organisms.

Fish bait Whole, frozen marine bait fish (see above), small tuna, coleoid cephalopod molluscs (squid, cuttlefish and octopus) imported for use as bait. Fish bait includes eviscerated fish or coleoid cephalopod molluscs originally imported for use for human consumption but downgraded for use as bait.

FishBase The FishBase Project: World online database for fish species (2019).

Free of disease Fish and coleoid molluscs with no external signs of infection, which have not been harvested following an epizootic incident, or as disease control.

viii Import risk analysis: Frozen fish and cephalopod molluscs for fish bait

Term/Acronym Definition

IHS Import Health Standard.

Marine fish Any species of fish that does not spend any part of its life cycle in fresh water. Salmon, which spawn in fresh water, are NOT a marine fish. Brackish water is considered to be part of the marine environment (MPI 2011).

MPI Ministry for Primary Industries, New Zealand.

Meltwater Water released from the melting of ice, including frozen fish bait.

NIWA National Institute for Water and Atmospheric Research, Crown Research Institute, New Zealand.

Non-viable Organism is not capable of living, growing or reproducing

Not cause significant disease

Pathogen is benign, causing no clinical signs of disease in the host species and is not a reservoir host for another pathogenic organism.

Opportunistic pathogen

Only causes clinical disease where the host is weakened or stressed, or infection only occurs as a result of physical injury.

OIE World Organisation for Animal Health, Paris, France.

RMP Risk management proposal.

SeaLifeBase The SeaLife Project. World database for non-fish marine species.

Shelf-stable Not requiring refrigeration or freezing before opening (MPI 2011).

SPS Agreement Agreement on the Application of Sanitary and Phytosanitary Measures.

Trunked cartilaginous fish

Processed fish of Class Chondrichthyes, where the viscera, gills and head have been removed.

Wild-sourced Organisms collected from the wild. This excludes finfish and cephalopods that have been farmed or that have otherwise spent any part of their life cycle in confinement.

WoRMS World online Register of Marine Species (2019).

WTO World Trade Organisation, Geneva, Switzerland.

Import risk analysis: Frozen fish and cephalopod molluscs for fish bait 1

Executive summary

This supplementary import risk analysis (SRA) reviews the biosecurity risks associated with

imported frozen fish bait. This commodity includes non-viable frozen, whole (uneviscerated)

wild-caught marine finfish (Class Actinopterygii) and molluscs (Subclass Coloidea, Class

Cephalopoda - cuttlefish, octopus and squid). It is restricted to specified species that were

imported for use as fish bait in 2018, as listed in Table 5.

Fish bait is a non-human food item, caught, stored and traded as a bulk commodity. The current

import requirements are defined in the Import Health Standard (IHS) for “Fish bait and fish

food” (FISFOOIC.ALL) (MPI 2011). This commodity must be wild caught, non-viable marine

finfish, either frozen at -18 °C for 168 hours (7 days) or treated with ionising radiation (at 25

kGyA). As cuttlefish, octopus and squid are not included in the definition of finfish in

FISFOOIC.ALL, they are imported as a food grade item, which is then on-sold as fish bait.

This SRA supplements an import risk analysis (IRA) completed in 2019, that reviewed trunked

elasmobranch and eviscerated teleost finfish imported for human consumption. As this IRA did

not include whole (uneviscerated) finfish or coleoid molluscs, the SRA assesses the biosecurity

risks associated with these commodities where imported for use as bait for commercial and

recreational fishing. Further, this definition is restricted to the finfish and coleoid cephalopod

mollusc species imported during 2018 (see Table 5) from estimated volumes and location of

capture data provided by the fish bait industry. All other fish bait species are ruled out of scope

of this risk analysis. Any other fish bait species not included in Table 5 should be subject to

specific risk analysis when requested by the Biosecurity New Zealand (BNZ) Animal Imports

team.

The preliminary hazard list for this SRA was largely derived from the earlier finfish IRA. It was

supplemented by a literature review to identify the preliminary hazards associated with coleoid

cephalopod molluscs, as no previous risk analysis has been completed on these species.

The combined preliminary hazard list identified 164 organisms or organism groups. Of these, 19

organisms (9 viruses, 5 bacteria and 5 myxozoan pathogens) were retained for risk assessment

after the hazard identification step. After risk assessment, 14 organisms (6 viruses, 3 bacteria and

5 myxozoan pathogens) were identified as representing non-negligible risk, as described in the

following table. Of these, only nervous necrosis virus (NNV) was associated with both finfish

and squid species. No other risk organisms were associated with coleoid cephalopods.

Group Pathogen

Virus Birnaviridae: marine aquabirnavirus (MABV)

Iridoviridae: Megalocytivirus (red sea bream iridovirus RSIV/Infectious skin and kidney necrosis virus (ISKNV))

Iridoviridae: erythrocytic necrosis virus (ENV)

Nodaviridae: nervous necrosis virus (NNV)

Rhabdoviridae: infectious haematopoietic necrosis virus (IHNV)

Rhabdoviridae: viral haemorrhagic septicaemia virus (VHSV)

Bacteria Edwardsiella spp.

Exotic Streptococcus species complex (S. agalactiae (serotype 3:283) and S. iniae)

Myxozoa Enteromyxum leei, Kudoa clupeidae, K, nova, K, iwatai, K, thyrsites

A Irradiation with 25 kGy is equivalent to a dosage of 2.5 Mrads (MPI 2011).

2 Import risk analysis: Frozen fish and cephalopod molluscs for fish bait

This SRA proposes general and specific risk management options. The general risk management

measures include:

• Frozen storage (pre-export or post-arrival storage in a transitional facility, at a temperature

range from -18 °C to -20 °C, for at least 4 months.

• Health certification from the relevant Competent Authority declaring:

o the species included in the commodity;

o the region/country of origin;

o that the commodity is sourced from wild stock;

o that the commodity has not been harvested from populations experiencing an epizootic

disease;

o has no visible signs of disease;

o labelling commodity as “For bait use only, unfit for human consumption”.

Extension of frozen storage (from the 7 days defined in FISFOOIC.ALL to 4 months)

substantially or completely denatures some bacteria (Edwardsiella spp.) and myxozoans

(Enteromyxum leei, Kudoa clupeidae, K. iwatai. K. nova and K. thyrsites). Therefore, no specific

risk management measures are considered necessary for these species.

For the remaining risk organisms listed in the table above, additional risk management measures

are considered necessary. The species-specific risk management options proposed include:

• Sourcing fish bait from species not associated with pathogens of concern.

• Sourcing fish bait from regions/countries recognised by BNZ as being free of corresponding

pathogens of concern, through the BNZ Country Approval Procedures.

• Pre-export or post-arrival batch testing of fish bait shipments to ensure absence of the

corresponding pathogens of concern, through the BNZ Country Approval Procedures.

• Pre-export or post-arrival irradiation (ionising radiation up to 50 kGyB).

The species-specific risk management options are described in the relevant chapter of this SRA

and summarised in Appendix 2. An assessment of the likely effects on pathogen occurrence in

the commodity associated with the general and specific risk management options is provided in

Appendix 3.

B Irradiation at 25 kGy denatures most bacterial, fungal and metazoan pathogens, but not all viruses (DAFF, 2012).

A dose of 50 kGy will denature all risk organisms associated with the commodity and is consistent with Australian

biosecurity guidelines (DAFF, 2013). It is acknowledged that no suitable irradiation facilities exist in New Zealand.

Further, a lower irradiation dose may be appropriate to denature the risk organisms associated with a particular fish

bait species. This may be determined on a case-by-case basis, as necessary.


Introduction

This supplementary import risk analysis (SRA) examines the biosecurity risks associated with

whole (uneviscerated) frozen marine species of finfish (class Actinopterygii) and cephalopod

molluscs (squid, cuttlefish and octopus - Subclass Coleoidea of Class Cephalopoda). These

finfish and cephalopod species were sourced from wild stocks and were imported for use as fish

bait in 2018 (Table 5). The SRA supplements a previous import risk analysis (IRA) that

examined risks associated with imported non-viable fresh, chilled and frozen eviscerated bony

fish (Osteichththyes) and trunked cartilaginous fish (Chondrichthyes). These may be sourced

from marine, fresh or brackish waters and are intended for human consumption (Blackwell,

2019). The 2019 IRA did not assess the risk to New Zealand from the importation of whole

(uneviscerated) finfish, or of cephalopod molluscs, so the relevant species have been assessed in

this SRA.

Finfish and cephalopod fish bait species, other than Octopus spp., generally occur in large

schools. Some, including herring (Clupea spp.) and sprat (Sprattus spp.) commonly occur in

mixed schools based on fish size (Maes & Ollevier, 2020). These fish bait species are caught for

human consumption and also support competing large-scale industrial fisheries for fish meal,

fish food and fish oil (Merino et al., 2014).

The knowledge on the population dynamics and epidemiology (including reservoir species,

modes of transmission, prevalence, morbidity and mortality) of the major risk organisms in

regard to lower value marine species such as fish bait, is complex and poorly understood (Munro

et al., 1983; Suttle, 2007; Dunn et al., 2012; Engelhard et al., 2014). Pathogen-associated

mortality in wild fish is generally un-noticed and rarely reported (Lafferty et al., 2015).

Translocated fish bait species represent a direct pathway for the spread of aquatic animal

diseases into the aquatic environment (Hine & MacDiarmid, 1997; Athanassopoulou & Roberts,

2004; Suttle, 2007; Herve-Claude et al., 2008; Dunn et al., 2012; Phelps et al., 2013; Pearce et

al., 2014; Oidtmann et al., 2013, 2017).

Pathogen introduction and establishment through the commodity has resulted in epizootics in

naïve wild fish host populations (including conspecifics separated by geography), causing

population level changes (Dunn et al., 2012). Examples include:

• Pilchard herpesvirus (PHV), which was introduced into the pilchard (Sardinops sagax)

wild fisheries in Australia in 1995 and New Zealand during 1996. This was reported to

have occurred through the use of infected frozen bait fish imported from California as

aquaculture feed and fish bait in Australia, and as fish bait in New Zealand (Smith et al.,

1996; Ward et al., 2001; Gaughan, 2002; Diggles, 2011). Recreational and commercial

fishers in New Zealand were observed collecting moribund and dead pilchards for later use

as fish bait in 1996 (Smith et al., 1996; Diggles, 2011). This introduction resulted in a 70-

75% decrease in spawning biomass in South Australian and New Zealand pilchard stocks

(Smith et al., 1996; Jones et al., 1997; Ward et al., 2001).

• A previous epizootic incident had occurred in Australian pilchards during 1988-89

(Whittington et al., 2008; Diggles, 2011). The lack of mortalities in New Zealand pilchards

during this period was attributed to a temporary ban on frozen pilchards imported from

Australia during the entire course of this first epizootic (Diggles, 2011).

https://en.wikipedia.org/wiki/Coleoidea


• White spot syndrome virus (WSSV), which affected wild and farmed wild prawns

(Penaeus spp.) in Australia. This was reported to have occurred through the use of

imported infected frozen prawns as fish bait by recreational fishers (McColl et al., 2004a;

Biosecurity Queensland, 2017).

• Epizootic haematopoietic necrosis virus (EHNV), which has been spread across the inland

waterways of Northern Australia through infected frozen redfin perch (Perca fluviatilis)

used as bait (Whittington et al., 2010; Diggles, 2011).

• Spring viraemia of carp virus (SVCV), which may be spread from infected farmed or wild

cyprinid finfish to wild carp by recreational fishers (Goodwin et al., 2004; Diggles, 2011).

The inter-connectedness of the aquatic environment means that pathogen transfer between wild

and farmed stocks can also occur (Harvell et al., 1999; Gaughan, 2002; Dunn et al., 2012;

Oidtmann et al. 2013, 2017; Georgiades et al., 2016; Groner et al., 2016). Exposure may be

direct, where infected fish bait is used as fish feed. It may also be indirect, where susceptible

wild stocks are initially infected and then these stocks act as reservoirs for infection of farmed

species (Diamant et al., 2007). Examples include:

• Pilchard orthomyxovirus (POMV), a disease of wild pilchards (Sardinops sagax) that

has transferred to farmed Atlantic salmon (Salmo salar) in Tasmanian aquaculture (Mohr et

al., 2020).

• Viral haemorrhagic septicaemia virus (VHSV) (Rhabdoviridae), which first became

established in the Mediterranean Sea during 2004 following the use of infected sprats

(Sprattus sp.) and herring (Clupea harengus) as feed for marine farmed rainbow trout

(Oncorhynchus mykiss). Epizootics then occurred in other wild and farmed species, including

farmed European seabass (Dicentrarchus labrax), a previously unknown host for VHSV

(Dixon, 1999; Skall et al., 2005a, 2005b).

• VHSV was introduced into the North America Great Lakes through use of live fish as

fish bait (Diggles, 2011).

• Crayfish plague (Aphanomyces astaci), which was introduced into European waters

through frozen fish tissue containing A. astaci. This pathogen remains infective unless the fish

is frozen for at least 72 hours (Oidtmann et al. 2002; Diggles, 2011; OIE, 2019b).

If an introduced marine pathogen with a wide host range (such as VHSV) becomes endemic, its

eradication may be difficult or impossible (Myers et al., 1992; Amos et al., 1998, 2010;

Hershberger et al., 1999; Gaughan, 2002; Hedrick et al., 2003; Arkush et al., 2006).

The process of disease establishment in a new host population is not however automatic. The

presence of a pathogen in a commodity in itself does not result in disease establishment.

Successful establishment depends on many factors involving the pathogen, the host and the

environment (Stevens, 1960; Salama & Rabe, 2013).

This SRA examines the risks associated with aquatic animal diseases exotic to New Zealand.

These include both the aquatic animal diseases listed in the OIE Aquatic Code (OIE, 2019a) and

other significant aquatic animal diseases (Blackwell, 2019). These diseases could have a major

effect on the economy, people (including society and culture) and environment of New Zealand.


2.1 The importance of fish bait in New Zealand

New Zealand is geographically isolated. It has the world’s fourth largest Exclusive Economic

Zone, comprising 1.3 million square kilometres, with a 15,100 km long coastline. New Zealand’s

marine fisheries (consisting of commercial, customary and recreational sectors) play an

important economic and societal role. Of the 16,000 marine species identified in New Zealand

waters, 130 species are commercially fished (FAO, 2020a).

2.1.1 Fish bait use in New Zealand commercial fisheries

The New Zealand commercial fisheries were valued at NZ$ 4.18 billion (annually on average

during 2000 to 2015) (BERL, 2017). In this period, seafood was the fifth largest export

commodity, by value, representing (on average) 3.2% of total annual exports. In this period, the

seafood sector directly employed 4,305 full time workers, with a total employment of 13,468

people, representing 0.7% of New Zealand’s total employment (BERL, 2017). New Zealand

fishery exports in 2018 were collectively valued at NZ$ 1,817 million (Seafood New Zealand,

2019).

Fish bait is an important component of commercial and recreational fishing operations.

Commercial fishing operations using fish bait caught an estimated NZ$345 million worth of fish

in New Zealand during 2010–2015 (BERL, 2017; MPI, 2018a, 2018b, 2018c). Fish bait is used

to attract marine fish in line fishing, pot fishing and purse seining methods. Baited hooks may be

set in offshore waters to attract pelagic fish (those that inhabit the water column) or discharged in

quantity as an attractant to hold an entire school of pelagic fish in position prior to capture by

purse seine. Baited hooks may also be used in offshore and inshore waters to attract demersal

(bottom dwelling) fish to baited hooks or discharged in quantity as ground bait (FAO, 2018,

2019).

The total number of baited hooks used in New Zealand waters has declined, from 65 million in

2002-03, to 43 million in 2018-19 (Table 1).

Fish bait is also used to attract and retain fish to baited pots used by commercial and recreational

fishers in coastal inshore waters (MPI 2018a, 2018b, 2018c). During the 2017-18 fishing year,

2,083,276 commercial pot fishing sets were conducted (J. Moriarty, BNZ, pers. comm., 2020).

Table 1. Estimated total number of hooks per year set in commercial long lining, by vessel size group, for fishing years (1 Oct to 30 Sept) 2002-03, 2012-13 and 2018-19

Fishing year Number of hooks per year (thousands) Vessels < 20 m Vessels 20-30 m Vessels > 34 m Total

2002-03 27,115,490 1,887,569 36,278,908 65,281,967

2012-13 32,525,000 9,992 5.635,005 38,169,997

2018-19 19,834,717 6,011,654 18,007,719 43,854,090

Source: 2000-01 to 2012-13 from Pierre et al., (2014). 2018-19 from Fisheries New Zealand, (2020; Moriarty, J, BNZ, pers. comm., 2020).

2.1.2 Fish bait use in New Zealand recreational fishing

Recreational marine fishing activity primarily uses fish bait. The contribution of recreational

marine fishing activity to the New Zealand GDP (Gross Domestic Product) is estimated at

NZ$ 638 million annually (Southwick et al., 2018). Data from the BNZ Recreational fishing

survey (Table 2) indicates that over 1.8 million fishing trips using fish bait were carried out


during 2017-18. Recreational fishing mainly occurs in coastal inshore and shallow shelf areas.

Recreational marine fishing often occurs adjacent to marine aquaculture structures, as sea cages

act as fish aggregation devices (P. Lamb, pers. comm., 2019).

Table 2. Estimated number of recreational fishing trips using fish bait during 2017-18

Recreational fishing method

Rod/line Longline/Kontiki Pot Total

Trailer motorboat 880,018 26,501 21,165 927,684

Large Motor launch 207,711 4,103 6,389 218,203

Trailer yacht 3,565 150 164 3,879

Large yacht 13,960 525 0 14,485

Kayak/rowboat 75,634 3,220 2,681 81,535

Off land 464,505 84,402 3,468 552,375

Other 8,457 207 475 9,139

Total 1,653,850 119,108 34,342 1,807,300

Source: National Recreational Fishing Panel Survey, 2017-18 (Fisheries New Zealand 2020b).

2.1.3 Bait loss from commercial and recreational fishing

The proportion of fish baits in recreational and commercial line fishing lost to the environment

may reach 50% (Skud et al., 1978). Bait loss may result from initial mis-hooking of the bait, loss

through mechanical abrasion during setting and retrieval, loss when baits hit the water surface, or

loss when passing through the water column. Bait loss may also occur where baits are partially

consumed by fish or invertebrates including shrimps and crabs.

The automatic baiting systems used on larger commercial vessels use thousands of hooks (Table

1) and the overall number of lost baits may be considerable (Skud et al., 1978; High, 1980;

Smith, 2001; Ward & Myers, 2007). Unwanted fish bait is commonly discarded at sea by

commercial and recreational fishers (High 1980; Lokkeborg et al., 2010, 2014; Kumar et al.,

2016; Wynn-Jones et al., 2019). It is possible that fish bait may be discarded by recreational

fishers adjacent to marine sea cages.

2.1.4 Finfish bait imports, 2018

Prior to 2008, New Zealand was essentially self-sufficient in fish bait. During 2008 – 2017, a

variety of fish and coleoid mollusc species have been imported (Appendix 1).

Available import data for frozen fish bait do not provide the location of capture. Industry-

sourced data on species, volume of trade and location of capture of fish bait imported in 2018

(Table 3) indicate this mainly comprised of species of families Clupeidae (58%), Scombridae

(20%) and Mugilidae (13%). In contrast to previous years (see Appendix 1), mackerel species

(Carangidae) were not imported in 2018. A minor amount (< 1 t) of flying fish (family

Exocoetidae) were imported as a trial in 2018 (B. Burney, pers. comm., 2019). This is considered

insignificant and flying fish are not considered further.

2.1.5 Coleoid cephalopod bait imports, 2018

Molluscs are not currently included in the definition of marine fish imported as frozen fish bait

(MPI, 2011). Import commodity data do not separate frozen coleoid molluscs used as fish bait

from imports for human consumption. Industry-sourced data on species, volume of trade and


location of capture of coleoid molluscs in 2018 (Table 4) indicate most (99%) cephalopod fish

bait consisted of squid (family Ommastrephidae).

2.2 The importance of aquaculture

The aquaculture sector employed 3000 people in 2015, with a contribution of $NZ 500 million to

the New Zealand economy. Of this total, $NZ 338.1 million was generated in exports

(Aquaculture, 2020). The New Zealand Government Aquaculture Strategy projects an increase

of this sector to reach $NZ 3 billion in annual sales by 2035 (Aquaculture, 2020). New Zealand

aquaculture is dependent upon access to clean marine and fresh waters (Lafferty et al., 2015;

Georgiades et al., 2016; Haenen, 2017).

Salmonid aquaculture in New Zealand is mainly focussed on Chinook (king) salmon

(Oncorhynchus tshawytscha), where hatchery-raised smolt are on-grown in fresh-water

aquaculture, or in sea-cages. Chinook salmon aquaculture was valued at $NZ 77 million in 2018

(Aquaculture New Zealand, 2019). Other developing marine finfish aquaculture species farmed

for food include yellowtail kingfish (Seriola lalandi), snapper (Sparus aurata), turbot (Colistium

nudipinnis) and grouper/bass (Polyprion oxygeneios) (NIWA, 2017a; Plant & Food, 2016).

While salmonid aquaculture uses purpose-designed pellet fish food, the aquaculture of non-

salmonid species may utilise fish bait species as aquaculture feed, as practiced in the Australian

aquaculture for Southern bluefin tuna (Thunnus maccoyii) (Ellis, 2016).

In fresh waters grass carp (Ctenopharyngodon idella) and silver carp (Hypophthalmichthys

molitrix) are also farmed for weed control purposes (Clayton & Wells, 1999; Blackwell, 2019).


Table 3. Estimated volume (t) and source location of finfish imported as fish bait, 2018

Finfish family Common name Scientific name Volume 1 (t)

Volume 1

(t) by family

Reported location2 of capture

Clupeidae Herring, pilchard, sardine, sprat 600 Spotted sardinella Amblygaster (=Sardinella)

sirm 390 Malaysia

Round sardinella Sardinella aurita 100 China South American

pilchard Sardinops sagax 100 China, Japan,

South Africa Indian oil sardine Sardinella longiceps 10 Indonesia

Scombridae Tuna, mackerel 208 Blue mackerel Scomber australasicus 90 Fiji Skipjack tuna Katsuwonas pelamis 55 China, Indonesia Little tunny Euthynnus alletteratus 28 China, Indonesia Atlantic mackerel Scomber scombrus 25 Spain Chub mackerel Scomber japonicus 10 China

Mugilidae Mullet Flathead grey mullet Mugil cephalus 140 140 Australia,

Indonesia

Scomberesocidae Saury Pacific saury (Samna) Cololabis saira 57 57 China, Chinese

Taipei

Engraulidae Anchovy Californian anchovy Engraulis mordax 33 33 United States

Hemiramphidae Halfbeak Ballyhoo halfbeak Hemiramphus (Esox)

brasiliensis 4 4 Indonesia

Exocoetidae Flying fish a Japanese flying fish Cheilopogon agoo <1 Japan Tropical two-wing

flying fish Exocoetus volitans <1 Japan

Total 1042 Source New Zealand fish bait industry: B. Burney, pers. comm., 2019; R. Clark, pers. comm., 2019; D. Rutherford, pers. comm., 2019; Spencer, pers. comm., 2019; C. Williams, pers. comm., 2019.

Notes 1. Estimated total weight provided by fish bait industry. 2. Reported location of capture provided by fish bait industry.


Table 4. Estimated volume (t) of coleoid cephalopod molluscs imported as fish bait, 2018

Family Species (current scientific name)

Common name Previous scientific name

Volume 1

(t) Volume1

(t) by family

Location of capture2

Ommastrephidae

Subfamily Illicinae

Illex argentinus Argentine shortfin squid

Ommastrephes argentinus

276 Argentina, China

Illex coindetii Broadtail shortfin squid

Illex illicibrosus 80 United States

Subfamily Todarodinae

Todarodes pacificus Japanese flying squid

Ommastrephes pacificus

350 706 China

Loliginidae Doryteuthis opalescens

Opalescent inshore squid

Loligo opalescens

80 United States

Loligo spp. Common squid Loligo spp. 1 81 Chinese Taipei, United States

Uroteuthis duvaucelli Indian squid Loligo duvaucelii 03 Malaysia3

Sepiidae Sepia recurvirostra Curvespine cuttlefish

Sepia recurvirostra

2 Malaysia

Sepia spp. Cuttlefish Sepia spp. 1 3 Malaysia

Octopodidae Octopus spp. Octopus Octopus spp. 1 1 China, Japan

Source New Zealand fish bait industry: B. Burney, pers. comm. 2019; R. Clark, pers. comm., 2019.

Notes 1. Estimated weight (t) provided by fish bait industry

2. Location of capture as provided by fish bait industry

3. Uroteuthis duvaucelii pre-ordered in 2018 to be caught during 2018-2019.

Scope, commodity definition and assumptions

3.1 Scope

This SRA assesses the biosecurity risks associated with imported frozen, whole, wild-caught

marine fish (Class Actinopterygii) and coleoid cephalopod molluscs (Class Cephalopoda,

subclass Coleoidea), ‘imported for use as fish bait’ under the Biosecurity Act 1993C.

This risk analysis includes whole coleoid cephalopod molluscs, which are imported for human

consumption and redirected into the fish bait pathway. However, other imported aquatic animal

products may become unfit for human consumption due to post-border product spoilage

(including freezer burn, incorrect frozen storage, or contamination). Most rejected product is

discarded, or processed into fish meal, but some may be re-directed into the fish bait pathway

(Blackwell, 2019). As this occurs in an ad-hoc manner, the volume of transferred product is

unknown, but likely to be small. While this latter pathway is not considered further, the findings

in this SRA should be considered in any future review of aquatic animal products for human

consumption.

Processed “consumer-ready aquatic animal products” imported for human consumption (MPI,

2001, 2008), fish bait products derived from imported ornamental fish, live or preserved fish or

molluscs for display, zoos or for biological research purpose (Hine & Diggles, 2005), or fish

The Biosecurity Act 1993 and Fisheries Act 1996 are available online from the Parliamentary Counsel Office of the New Zealand Government at: http://www.legislation.govt.nz/act.


food derived from any other plant or animal sources (Cobb, 2008) are outside the scope of this

analysis.

Fish bait caught in fresh, brackish or estuarine waters is also outside the scope of this risk

analysis. Euryhaline species such as flathead grey mullet (Mugil cephalus) are considered in-

scope if caught in the marine environment, but not if caught in fresh or brackish waters.

Fish bait may also be imported as a “user-ready shelf-stable” product that has been chemically or

heat-processed (MPI, 2011). This commodity is outside the scope of this risk analysis.

3.2 Commodity definition

For the purposes of this risk analysis, the commodity definition is restricted to non-viable finfish

of Class Actinopterygii and coleoid cephalopod molluscs of Class Cephalopoda: Subclass

Coleoidea (squid, cuttlefish and octopus) currently imported into New Zealand for use as fish

bait.

The commodity definition for finfish (Class Actinopterygii) used in this SRA is consistent with

the definition used in the IRA for eviscerated or trunked fish for human consumption (Blackwell,

2019).

Coleoid cephalopod molluscs were not included in the IRA for eviscerated or trunked fish for

human consumption (Blackwell, 2019) and no previous risk analysis has been completed for this

commodity. For the purposes of this risk analysis, the definition of coleoid cephalopod molluscs

is consistent with the Fisheries Act, 1996 as:

“All species of the phylum Echinodermata and phylum Mollusca and all species of the

class Crustacea at any stage of their life history, whether living or dead.”

Not all molluscs are used as fish bait (B. Burney, pers. comm., 2019). For the purposes of this

risk analysis, molluscan fish bait is restricted to members of sub-class Coleoidea (squid,

cuttlefish and octopus). This excludes the shelled cephalopods of genera Allonautilus and

Nautilus (Subclass Nautiloidea) (WoRMS, 2019) as these are not used as bait.

Cephalopod molluscs are defined (Jereb et al., 2010) as:

“The class within the Phylum Mollusca, characterized by bilateral symmetry, internal

‘shell’ or absence of shell (except nautiluses), anterior head, appendages and funnel,

posterior mantle, mantle cavity with organs, and shell and fins when present.”

Sub-class Coleoidea is defined (MolluscaBase, 2019; WoRMS, 2019) as:

“molluscs within the Class Cephalopoda including squid, (families Loliginidae and

Ommastrephidae), cuttlefish (family Sepiidae) and octopus (family Octopodidae) but

excluding nautiloid cephalopods of Subclass Nautiloidea.”

Fish bait is a non-human food item currently imported as a whole (uneviscerated) bulk

commodity, under Section 7.2.4 of the Import Health Standard (IHS) for fish bait and fish food


(FISFOOIC.ALL) (MPI, 2011). Under this IHS, fish bait is restricted to marine finfish (Class

Actinopterygii) which is:

• Labelled with species and country of capture

• Non-viable and either:

o Frozen (to below -18 °C for at least 18 hours); or,

o Irradiated with 2.5 MradsD.

The IHS (FISFOOIC.ALL) is restricted to marine finfish. For the purposes of this risk analysis,

the definition of fish bait is extended to include coleoid molluscs (Class Cephalopoda, subclass

Coleoidea). Further, this definition is restricted to the finfish and coleoid cephalopod mollusc

species listed in Table 5. This has been determined from import data from 2018 and information

provided by the fish bait industry (B. Burney, pers. comm., 2019; R. Clark, pers. comm., 2019; P.

Lamb, pers. comm., 2019; M. Lyford, pers. comm., 2020; C. Williams, pers. comm., 2019).

All other fish bait species are ruled out of scope of this risk analysis. Any other fish bait species

not included in Table 5 would be subject to specific risk analysis when requested by the

Biosecurity New Zealand Animal Imports team.

This commodity includes fish bait that is sorted to species and frozen (at sea or on shore) into

blocks of varying size (typically around 20 kg). The blocks are each placed into cardboard

cartons and these are stored in frozen containers, ready for export (B. Burney, pers. comm.,

2020). Imported fish may be used in the whole form as fish bait or be further processed into

berley.

Frozen fish bait is transported in refrigerated containers. The standard refrigerated containers used

in international trade can be maintained within a temperature range from -20 °C to + 10 °C (Sea

Containers, 2020). In their frozen storage mode these containers are maintained within a

temperature range from -18 °C, to -20 °C (B. Burney, pers. comm., 2019; T. Leighton, BidFood

Ltd., Wellington, pers. comm., 2020).

D Irradiation with 2.5 Mrads is equivalent to a dosage of 25 kGy. It is suggested this be increased to 50 kGy, consistent with Australian Government biosecurity guidelines (DAFF, 2013).


Table 5. Finfish and coleoid cephalopod mollusc families and species considered as fish bait

Category Family Species

Finfish Carangidae Trachurus japonicus, T. murphi, T. symmetricus, T. trachurus

Clupeidae Ambylygaster sirm, Clupea harengus, Clupea pallasii pallasii, Sardinella aurita, Sardinella lemuru, Sardinella longiceps, Sardinops sagax

Engraulidae Engraulis mordax, E. ringens

Hemiramphidae Hemiramphus balao, H. brasiliensis, H. dussumieri, H. lutkei

Mugilidae Chelon auratus, C. labrosus1, C. ramada, Mugil cephalus

Scomberosocidae Cololabias saira

Scombridae Auxis rochei, A. thazard, Euthynnus alletteratus, Katsuwonas pelamis, Sarda chilensis, S. lineolata, S. sarda, Scomber australasicus, S. japonicus, S. scombrus, Thunnus obesus

Cephalopods Loliginidae Doryteuthis opalescens, Loligo spp., Uroteuthis duvaucelli

Ommastrephidae Illex argentinus, I. coindetti, Todarodes pacificus

Sepiidae Sepia recurvirostrata, Sepia spp.

Octopodidae Octopus spp.

Notes

1 The thicklip grey mullet Chelon (Mugil) labrosus is included here because of potential confusion with the flathead grey mullet (Mugil cephalus). Source

New Zealand fish bait importers, Animal Imports team, Biosecurity New Zealand

3.3 Assumptions

For the purposes of this risk analysis, the following general assumptions are made:

• Susceptibility determined experimentally (where the infection process closely follows a

likely route of natural infection e.g. per os or cohabitation) represents a potential exposure

pathway.

• Pathogen epidemiology observed in aquaria or experiment follows similar processes to

disease in wild stocks of fish and coleoid molluscs.

• Organisms that have never been reported in the scientific literature or technical databases

(such as the New Zealand Organism Register (NZOR, 2017)) are assumed to be absent from

New Zealand (exotic), following the principles of Chapter 1.4 of the OIE Aquatic Code

(OIE, 2019a).

• Frozen storage (to at least -18 °C for at least 168 hours (7 days)) is assumed to denature

fungi, protozoa, nematodes, monogenean, digenean and cestode helminths, as well as most

metazoan parasites present in the commodity (Blackwell, 2019; USDA, 2019). These

pathogens are not considered further.

• Viral, bacterial and myxozoan pathogens may remain viable in fish bait frozen (to -18 °C

or -20 °C for 168 hours (7 days)) (Hine & MacDiarmid, 1997; Johnston, 2008).

• Host range may be wider than reported in the literature, particularly for low value species

such as fish bait (Blackwell, 2019; OIE, 2019a). As the scope of the commodity is confined

to listed fish bait species (Table 5), the rule that applied in Blackwell (2019) has been

modified, following the principles of Chapter 1.5 of the OIE Aquatic Code (OIE 2019a), as

follows:

o If only one species in a genus is susceptible to an identified risk organism, then only

that species is excluded in the species declaration option

o If two or more species in a genus are susceptible to an identified risk organism, then all

listed fish bait species in that genus are excluded in the species declaration option


o If two or more genera in a family are susceptible to an identified risk organism, then all

listed fish bait species in the family (or sub-family) are excluded in the species

declaration option.

Risk analysis methodology

4.1 General procedures

The methodology used in this risk analysis is guided by the Biosecurity New Zealand Risk

Analysis Procedures – Version 1 (Biosecurity New Zealand, 2006), the Handbook on Import

Risk Analysis for animals and animal products (OIE, 2010) and Chapter 2 of the Aquatic Animal

Health Code (OIE, 2019a). The risk analysis process comprises several steps: hazard

identification, risk assessment and risk management (Figure 1). It also includes risk

communication.

4.1.1 Hazard identification

A list of organisms of concern associated with the commodity (the preliminary hazard list) is

compiled from the OIE list of aquatic animal diseases (OIE, 2019a), published BNZ risk

analyses and from the relevant published scientific literature. For each organism on this list,

several steps are completed: formal identification (taxonomic classification), OIE status, New

Zealand status, together with a determination of whether or not the organism meets key

biological and epidemiological characteristics to be considered as a hazard.

Hazard identification concludes with an assessment of whether or not the organism is identified

as a hazard in the commodity. The results of the hazard identification are commonly summarised

as a table. All organisms deemed to be hazards are subjected to further risk assessment.

4.1.2 Risk assessment

Risk assessment (Figure 1) consists of four steps applied to each hazard: entry assessment,

exposure (and establishment) assessment, consequence assessment and risk estimation. At each

of the first two steps a qualitative assessment is made of the likelihood (of entry, exposure and

establishment, respectively), based on the available epidemiological information. The

consequence assessment is undertaken to determine the likely direct and indirect impacts of

entry, exposure and establishment of a risk organism in New Zealand. This includes effects on

people (including societal and cultural), the New Zealand environment and the New Zealand

economy (Biosecurity Act 1993, Section 23).


Figure 1. The general risk analysis process

Organisms of concern

HAZARD IDENTIFICATION

Can the organism remain viable inthe commodity ?

Does transmission require a vector?

Is the vector present

in New Zealand?

Is the pathogen present in New Zealand

Are there different

strains overseas?

Not identified as a hazard in this

risk analysis

Identified as ahazard in thisrisk analysis

RISK ASSESSMENT

Entry AssessmentLikelihood of hazard entering New Zealand

on the pathway

Exposure AssessmentLikelihood of exposure and establishment in

New Zealand

Consequence AssessmentLikely impacts on economy,

environment andhuman health in New Zealand

Risk Estimation

Organism is assessed

to be a risk

Risk Estimation

Not assessed to

be a risk

RISK MANAGEMENT

What options are available to managethe risk ?

What is the effect of each measure on the

level of risk?

yes

yes

yes

yes

no

no

no

no

no

no

yes

negligible

negligible

negligible

-

non negligible

-non negligible

non negligible

Is the organism present in theCommodity ?

yes

RISK COMMUNICATION

In addition to using the terms negligible and non-negligible to describe risks, the entry, exposure

and consequence assessments also use qualitative descriptors (ranging from very low to very

high) to describe the comparative levels of likelihood (in the case of entry and exposure

assessments) or impact (in the case of consequence assessment) (Biosecurity New Zealand,

2006).

The Biosecurity New Zealand Risk Analysis Procedures (Biosecurity New Zealand, 2006)

provides a more detailed explanation of the terminology as follows:

Risk attributes Description

Negligible Not worth considering, insignificant

Non-negligible Worth considering, significant

Risk descriptors

Very low Close to insignificant

Low Less than average, coming below the normal level

Medium Around the normal or average level

High Extending above the normal or average level

Very high Well above the normal or average level

These qualitative levels of likelihood of entry, exposure and establishment and the levels of

impact (economic, social, and environmental) were combined to assess whether the associated

risk is negligible or non-negligible.


If, either the likelihood of entry or the likelihood of exposure of a pathogen is assessed as

negligible, then the overall risk is deemed negligible and the remaining steps, being redundant, are

not undertaken.

If the expected likelihood of entry or exposure are assessed to be non-negligible, but the impacts of

establishment are assessed as negligible, then the overall risk is also negligible, and no further

analysis is necessary.

Those pathogens considered to pose a non-negligible risk are deemed to require risk

management. The relative magnitude of the risk (very low to very high) assists in determining

the stringency of risk management measures that would be needed to reduce the risk to an

acceptable level.

4.1.3 Risk management

Risk management identifies the options available for managing non-negligible risks, based on

the epidemiology of the risk organism. Where the OIE Code (OIE, 2019a) lists recommendations

for the management of a risk, these are described alongside additional risk management options,

where available, from the scientific literature.

In addition to the options presented, prohibition may also be considered. Recommendations for

the appropriate sanitary measures to achieve the effective management of risks are not made in

this document. These will be determined when an Import Health Standard (IHS) is drafted.

As obliged under Article 3.1 of the World Trade Organisation (WTO) SPS Agreement (WTO,

2017), the measures adopted in IHSs will be based on international standards, guidelines and

recommendations where they exist, except as otherwise provided for under Article 3.3 of the

SPS Agreement. That is, measures providing a higher level of protection than international

standards can be applied if there is scientific justification, or if there is a level of protection that

the member country considers is more appropriate. These additional measures must be based on

a scientific risk analysis.

4.1.4 Risk communication

After a draft import risk analysis has been written, BNZ analyses the options available and

proposes draft measures for the effective management of the identified risks. These are then

presented in a draft IHS that is released for public comment, together with a risk management

proposal (RMP) that summarises the options analysis, the rationale for the proposed measures

and provides a link to the draft risk analysis.

Not every risk organism identified in the risk analysis may be associated with a particular

imported fish bait species and require risk management in an IHS. The RMP will take into

account specific information that would affect the need for risk management measures. For

instance, factors considered in an RMP would include (but not be limited to) the country of

origin of the commodity, the presence or absence of risk organisms in that country, the species

from which the commodity is derived, and any manufacturing processes that inactivate risk

organisms.

The document package (draft IHS, RMP and risk analysis) is then released for stakeholder

consultation. Stakeholder submissions in relation to these documents are reviewed and

published, including any supplementary risk analyses that may be required, before a final IHS is

issued.


Hazard identification

Previous risk analyses have been completed for fish bait and fish food (Cobb, 2009) and for

eviscerated/trunked fish for human consumption (Blackwell, 2019). No previous risk assessment

has been completed for coleoid cephalopod molluscs (squid, cuttlefish, and octopus) imported

for any end use. The preliminary hazard list was compiled using Cobb (2009) and Blackwell

(2019), the OIE list of aquatic animal diseases (OIE, 2019a), the peer-reviewed scientific

literature, health databases, previous BNZ risk assessments and information provided by experts

and interested parties. Each organism was assessed in terms of eight key criteria:

• OIE status

• New Zealand status

• Viability in fish bait (frozen to -20 °C for 168 hours (7 days))

• Cause significant disease

• Presence of more virulent exotic strains

• Necessity for a vector or intermediate host in the life cycle

• Zoonotic potential

• Potential hosts in New Zealand.

Not all the organisms identified as hazards in the previous risk analyses are relevant to the

commodities considered here. Some organisms were not considered as hazards, Blackwell

(2019) because they were not present in eviscerated or trunked fish. These, however, may be

present in whole frozen fish or coleoid cephalopods.

Only the pathogens reported from the fish and coleoid cephalopod host species listed in Table 5

are considered for this analysis. To establish in New Zealand, a pathogen must be present and

viable in the commodity.

The findings of the Hazard Identification for pathogens derived from finfish and coleoid

cephalopod molluscs are presented in Table 6. All organisms identified as hazards were

subjected to risk assessment.


Table 6. Hazard Identification table for specified1 wild marine finfish and coleoid molluscs used as fish bait

Pathogen Fish host family Species known to be susceptible

Host distribution OIE listed disease

Reported from New Zealand

Viable in frozen fish bait

Causes significant disease

Virulent exotic strains

Vector/ intermediate host required

Zoonotic potential

Potential hosts in New Zealand

Retained for risk assessment

Reference

Viruses

Astroviridae

Eastern sea garfish astrovirus

Hemiramphidae Eastern sea garfish (Hyporhamphus australis)

Southwest Pacific, Australia

N N Y N N/A N N Y N Geoghegan et al., 2018

Birnaviridae

Marine aquabirnavirus (MABV)

Infectious pancreatic necrosis virus-like (IPNV-like strains) including viral deformity virus (VDV) and yellowtail ascites virus (YAV)

Carangidae Japanese amberjack (Seriola quinqueradiata), Japanese jack mackerel (Trachurus japonicus), Mediterranean horse mackerel (Trachurus mediterraneus). yellowtail amberjack (Seriola lalandi)

Widespread Western Atlantic (Gulf of Cadiz), Mediterranean Sea, Japan, North-central Pacific, Australia

N Y Y Y Y N N Y Y Sorimachi & Hara, 1985; Castric, 1997,;Hanlon & Forsythe, 1990; Nakajima et al.,1993; Diggles, 2004a, 2004b; Isshiki et al., 2004; Tisdall & Phipps, 1987; Davies et al., 2010;; Munro & Midtlyng, 2011;; Ogut & Altuntas, 2014,; Diggles, 2016

MABV (cont.) Clupeidae Atlantic herring (Clupea harengus), sardine (Sardinops spp.), Cllupeidae (wide host range)

West Atlantic (Gulf of Cadiz)

N N Y Y N/A N N Y Y Nakajima et al., 1998; Crane et al., 2000; Crane & Williams, 2008; Wallace et al., 2008; McColl et al., 2009; Crane & Hyatt, 2011; Diggles, 2011 2016; Fish Base, 2019

MABV (cont.) Engraulidae European anchovy (Engraulis encrasiciolus)

West Atlantic (Gulf of Cadiz), Mediterranean Sea

N N Y Y N/A N N Y Y Wallace et al., 2008; Ogut & Altuntas, 2014

MABV (cont.) Scombridae Atlantic mackerel (Scomber scombrus)

West Atlantic (Gulf of Cadiz)

N N Y Y N/A N N Y Y Wallace et al., 2008; Moreno et al., 2014

Bunyaviridae

Eastern sea garfish bunya-like virus




Hepnaviridae

Eastern sea garfish hepatitis B virus




Herpesviridae

Malacoherpesvirus (unclassified)

Octopodidae Octopus vulgaris United States N N Y N N/A N N Y N Prado-Alvarez & Garcia-Fernandez, 2019

Pilchard herpesvirus Clupeidae South American pilchard (Sardinops sagax) (= S. neopilchardus)

Southwest Pacific (Australia, New Zealand)

N Y Y Y N N N Y N Crockford et al. 2005; Hanson et al., 2011









Zoonotic potential



Reference

Iridoviridae

Subfamily Megalocytivirus

Red sea bream iridovirus (RSIV), infectious spleen and kidney necrosis virus ISKNV, heart and skeletal muscle inflammation (HSMI) and associated viruses

Carangidae Japanese jack mackerel (Trachurus japonicus), bigeye trevally (Caranx sexfasciens), Japanese scad (Decapterus maruadsi), shrimp scad (Alepes djedaba (= Carynx calla)), yellowstripe scad (Selaroides leptolepis), barred queenfish (Scomberoides tala (= Chorinemus hainanensis)). double spotted queenfish (Scomberoides lysan (= Chorinemus moadetta)

Northwest Pacific (China, Korea, Japan, southeast Asia)

Y N Y Y N/A N N Y Y Wang et al., 2007; Anon., 2017a; OIE, 2019a; Rimmer et al., 2015, 2017; CIFA, 2019

RSIV (cont.) Clupeidae Dotted gizzard shad (Konosirus (Clupanodon) punctatus), Pacific herring (Clupea pallasii pallasii)

Northwest Pacific, (China, Korea, Japan, southeast Asia)

Y N Y Y N/A N N Y Y Wang et al., 2007

RSIV (cont.) Mugilidae Flathead mullet (Mugil cephalus), longarm mullet ((Osteomugil cunnesius (= Mugil (Osteomugil) ophuyseni))

Widespread Indo-Pacific

Y N Y Y N/A N N Y Y Gibson-Kueh et al., 2004, Wang et al., 2007; OIE, 2019a; Rimmer et al., 2015, 2017

RSIV (cont.) Scombridae Chub mackerel (Scomber japonicus), greater amberjack (Seriola dummerili), yellowtail amberjack (Seriola lalandi), Japanese amberjack (S. quinqueradiata), Japanese Spanish mackerel (Scomberomorus niphonius), northern bluefin tuna (Thunnus thynnus)

Widespread Indo-Pacific, anti-tropical

Y N Y Y N/A N N Y Y Wang et al., 2007; Rimmer et al., 2015, 2017; Anon., 2017a; OIE, 2019a; CIFA., 2019

Viral erythrocytic necrosis virus (VEN or ENV) and associated viruses (Iridoviridae)

Clupeidae Atlantic herring (Clupea harengus), Pacific herring (C. pallasii pallasii)

Atlantic and Pacific Salish Sea, Alaska

N N Y Y N/A N N Y Y Reno et al., 1985; Emmenegger et al., 2014

VEN (Cont.). Engraulidae Californian anchovy (Engraulis mordax)

Northeast Pacific N N Y Y N/A N N Y Y Pagowski et al., 2019









Zoonotic potential



Reference

Subfamily Lymphocystivirus

Lymphocystis disease iridovirus (LCDV)

Clupeidae Indian oil sardine (Sardinella longiceps)

Widespread N Y Y N N/A N N Y N Matsusato, 1975; Leong & Colorni, 2002; Borrego et al., 2017; Vijapoopathy et al., 2016

Lymphocystis iridovirus (cont,)

Mugilidae Flathead grey mullet (Mugil cephalus)

Widespread N Y Y N N/A N N Y N Alexandrawicz,1951; Ovcharenko, 2016

Iridoviridae (unclassified)

Octopodidae, Sepiidae

Octopus vulgaris Western Atlantic N N Y N (not in wild stocks)

N/A N N Y N Rungger et al., 1971; Farley, 1978

Nodaviridae: Betanodavivus

Korean shellfish nervous necrosis virus (KSNNV) (Nodaviridae)

Sepiidae Sepia spp. Korea, China N N Y N N/A N N Y N Kim et al., 2018; Bandin & Souto, 2020

Viral encephalopathy and retinopathy (VER) and nervous necrosis virus (VNN) (NNV) Striped Jack nervous necrosis virus (SJNNV)

Carangidae Atlantic horse mackerel (Trachurus trachurus), greater amberjack (Seriola dummerili), Japanese jack mackerel (T. japonicus), permit (yellow-wax pompano) (Trachinotus falcatus), snub-nose pompano (Trachinotus blochii), white trevally (striped jack) (Pseudocaranx dentex), yellowtail amberjack (S. quinqueradiata)

Widespread N N Y Y N/A N N Y Y Mori et al., 1991, 1992; Danayadol et al., 1995; Nguyen et al., 1996; Tubbs et al., 2007; Gomez, 2010; Crane & Hyatt, 2011; Forrest et al., 2011; IDAAD, 2019; OIE 2019a

NNV (cont.) Clupeidae European pilchard (Sardina pilchardus)

Widespread N N Y Y N/A N N Y Y Panzarin et al., 2012

NNV (cont.) Engraulidae Japanese anchovy (Engraulis japonicus)

Northwest Pacific (Japan)

N N Y Y N/A Y N Y Y Gomez et al., 2006

NNV (cont.) Mugilidae Flathead grey mullet (Mugil cephalus), golden grey mullet (Chelon (Mugil) auratus) (= Liza aurata), leaping mullet (Chelon (=Liza, Mugil) saliens), thicklip grey mullet (Chelon (=Mugil) labrosus)

Mediterranean, Caspian Sea, Indo-Pacific (Israel, Iran)

N N Y Y N/A N N Y Y Ucko et al., 2004; Panzarin et al., 2012; Zorriehzahra et al., 2005, 2016

NNV (cont.) Octopodidae Octopus vulgaris Italy N N Y Y N/A N N Y Y Fichi et al., 2015; Bandin & Souto, 2020

NNV (cont.) Scombridae Chub mackerel (Scomber japonicus)

Korea N N Y Y N/A N N Y Y Gomez et al., 2008

NNV (cont.) Todaronidae Todarodes pacificus Widespread N N Y Y N/A N N Y Y Ford et al., 1986; Gomez, 2010; Fiorito et al., 2015; OIE, 2019b









Zoonotic potential



Reference

Betanodavirus (unclassified strains 1 and 2)

Octopodidae Octopus vulgaris Europe N N Y N N/A N N N N Fichi et al., 2015; Fiorito et al., 2015; Prado-Alvarez & Garcia-Fernandez, 2019

Orthomyxoviridae

Isavirus

Infectious salmon anaemia virus (ISAV) (HPR0 ISAV, HPR-deleted ISAV)

Clupeidae Atlantic herring (Clupea harengus)

Widespread Y N Y Y N/A N N Y Y Nylund et al., 2002; Hine & Diggles, 2005; Tubbs et al., 2007; Diggles, 2011; OIE, 2019a

Orthomyxovirus

Pilchard orthomyxovirus (POMV)

Clupeidae South American pilchard (Sardinops sagax) (=S. neopilchardus)

Southwest Pacific (Australia, New Zealand)

N Y Y N N N N Y N SCAAH, 2015; Diggles, 2016; PROMED, 2017; NIWA, 2018; Blackwell, 2019

Picornaviridae

Eastern sea garfish picornavirus


Southwest Pacific (Australia)


Reoviridae: Aquareovirus

Piscine aquareovirus (PRV) and associated viruses

Carangidae Atlantic horse mackerel (Trachurus trachurus)

Widespread N N Y Y N/A N N Y Y Fauquet et al., 2005; Cobb, 2008; King et al., 2011; Garseth et al., 2012; Roberts, 2012; Wiik-Nielsen et al., 2012; Carlile et al., 2014

PRV (cont.) Clupeidae Atlantic herring (Clupea harengus)

Northeast Atlantic N N Y Y N/A N N Y Y Wiik-Nielsen et al., 2012

Reoviridae (unclassified)

Sepiidae Sepia officianalis Europe N N Y N N N N Y N Devauchelle & Vago, 1971

Rhabdoviridae

Eastern sea garfish rhabdovirus




Infectious haematopoietic necrosis virus (IHNV)

Clupeidae Pacific herring (Clupea pallasii pallasii)

East, West Pacific, Northeast Atlantic

Y N Y Y N/A N N Y Y Hart et al., 2011; Dixon et al., 2016; OIE, 2019a

Viral haemorrhagic septicaemia virus (VHSV) (Rhabdoviridae)

Carangidae Mediterranean horse mackerel (Trachurus mediterraneus), Japanese amberjack (Seriola quinqueradiata)

Black Sea, Europe, Northwest Pacific (Japan, S. Korea)

Y N Y Y N/A N N Y Y Ito et al., 2004; Kim et al., 2013a; OIE, 2019a









Zoonotic potential



Reference

VHSV (cont.). Clupeidae Atlantic herring (Clupea harengus)

Black Sea, Europe, Widespread, East and West Atlantic

Y N Y Y N/A N N Y Y Meyers et al., 1994; Traxler & Kieser, 1994; Kocan et al., 1997; Dixon et al., 1997; Marty et al., 1998; Mortensen et al., 1999; Traxler et al., 1999; Kocan et al., 2001; Biosecurity Australia, 2002; Hedrick et al., 2003; Goodwin et al., 2004; OIE, 2019a

VHSV (cont.). Clupeidae Pacific herring (Clupea pallasii pallasii)

Pacific Ocean Y N Y Y N/A N N Y Y Meyers et al., 1994; Hershberger et al., 1999; Meyers & Winton, 1995; Marty et al.,1998; Biosecurity Australia, 2002; Purcell et al., 2012; OIE, 2019a

VHSV (cont.). Clupeidae European sprat (Sprattus sprattus)

Northeast Pacific (Alaska, Canada)

Y N Y Y N/A N N Y Y Skall et al., 2005a, 2005b; Crane & Hyatt, 2011; Diggles, 2011; Ogut & Altunas, 2014; Sandlund et al., 2014; CIFA, 2019

VHSV (cont.). Clupeidae European pilchard (Sardinia pilchardus)

East Atlantic Y N Y Y N/A N N Y Y Herve-Claude et al., 2008

VHSV (cont.) Clupeidae South American pilchard (Sardinops sagax)

Pacific Y N Y Y N/A N N Y Y Biosecurity Australia, 2002; OIE, 2019a

VHSV (cont.) Engraulidae European anchovy (Engraulis encrasicolus)

Black Sea, Northeast Atlantic (Europe) Widespread

Y N Y Y N/A N N Y Y OIE, 2019a

VHSV (cont.) Mugilidae Flathead grey mullet (Mugil cephalus)

Northwest Pacific (S. Korea), Northeast Pacific (California)

Y N Y Y N/A N N Y Y Biosecurity Australia, 2002; Lee et al., 2007; Kim & Faisal, 2010; Kim et al., 2013a; OIE, 2019a

VHSV (cont.). Scombridae Chub mackerel (Scomber japonicus)

Northeast Pacific (California)

Y N Y Y N/A N N Y Y Hedrick et al., 2003; Kim et al., 2013a; Anon., 2017a; OIE, 2019a

Unassigned virus Octopodidae Macroctopus maorum, Octopus vulgaris

Widespread, New Zealand

N Y Y Y N N N Y N Prado-Alvarez & Garcia-Fernandez, 2019

Unassigned virus (possibly Ostreid herpesvirus (OsHV-1)

Loliginae Loligo pealei Northwest Atlantic, Northeast Pacific

N Y Y N N N N N N Hanlon & Forsyth, 1990; Prado-Alvarez & Garcia-Fernandez, 2019

Bacterial pathogens

Alphaproteobacteriaceae

Rickettsia-like organisms (RLO)

Octopodidae Octopus vulgaris Northwest Atlantic, Mediterranean

N Y Y Y N Y N Y N Gestal et al., 1988; Fiorito et al., 2015; Diggles, 2011









Zoonotic potential



Reference

Bacillaceae

Bacillus spp. Octopodidae Octopus vulgaris Mediterranean Sea, West Atlantic

N Y Y N N N Y Y N Ford et al., 1986; Farto et al., 2014; NZ Fungi, 2019

Betaproteobacteriaceae

Acinetobacter anitratus (= A. baumanii)

Octopodidae Enteroctopus dofleini Europe N Y N Y N N Y Y N Stoskopf et al., 1987; Fiorito et al., 2015; Anon., 2017b

Achromobacter sp. Mugilidae Flathead grey mullet (Mugil cephalus), Longarm mullet (Osteomugil (Mugil) cunnesius)


N Y Y N N N Y Y N Almedia et al., 1968; Paperna & Overstreet, 1981; Ovcharenko, 2016

Enterobacteriaceae

Aeromonas caviae (= A. punctata) (Aeromonas hydrophila species complex)

Loliginidae Loligo forbesi, Loligo sp. Indo-Pacific (Philippines)

N Y Y (20 days at -20⁰C)

Y N N Y Y N Baldrias & Alvero, 1999; Joseph et al., 2013; Fiorito et al., 2015

Aeromonas hydrophila (cont.)

Carangidae Mediterranean horse mackerel (Trachurus mediterraneus)

Mediterranean Sea (Turkey)

N Y Y Y N N Y Y N Boran et al., 2013; Ozturk & Altinok, 2014; DermNet NZ, 2014




N Y Y Y N N Y Y N Paperna & Overstreet, 1981; Ovcharenko, 2016


Octopodidae Octopus joubini Widespread N Y Y Y N N Y Y N Fiorito et al., 2015; NZ Fungi, 2019


Octopodidae Octopus spp. Widespread N Y Y Y (Zoonotic) N N Y Y N DermNet NZ, 2014; Gestal et al., 2019

Aeromonas salmonicida var. salmonicida (atypical strains)

Clupeidae Atlantic herring (Clupea harengus), Pacific herring (C. pallasii pallasii)

Widespread N Y Y Y N N N Y N Anderson et al., 1994; Evelyn, 2001; Keeling et al., 2013; Brosnahan et al., 2018b

Aeromonas spp. Loliginidae Lollicuncula brevis Northeast Pacific N Y Y Y N N Y Y N Ford et al., 1986; Sherrill et al., 2000; Fiorito et al., 2015

Aeromonas spp. (cont.)

Sepiidae Sepia offinianalis Northeast Pacific N Y Y Y N N Y Y N Hanlon et al., 1984; Ford et al., 1986; Sherrill et al., 2000; Fiorito et al., 2015

Aeromonas spp. (cont.)

Octopodidae Octopus sp. Northeast Pacific N Y Y Y N N Y Y N Hanlon et al., 1984; Ford et al.,1986; Sherrill et al., 2000; Fiorito et al., 2015

Citrobacter freundii Sepiidae Sepia officinalis Europe N Y Y Y N N Y Y N Sherrill et al., 2000; DermNet NZ, 2014

Edwardsiella species complex


India N N Y Y N/A N Y Y Y Kumar et al., 2016









Zoonotic potential



Reference

Edwardsiella species complex (cont.)


Widespread N N Y Y N/A N Y Y Y Kusuda et al., 1976; Miyazaki & Egusa, 1976; Evans et al., 2011; Plumb & Hansen, 2011; Park et al., 2012; Kluzik & Woodford, 2016; Haenen, 2017; Reichley et al., 2017

Enterobacter cloacae Mugilidae Flathead grey mullet (Mugil cephalus)


N Y Y Y N N Y Y N Sekar et al., 2008; Ozturk & Altinok, 2014; Clark et al., 2018; Yow et al., 2018

Escherichia sp. Mugilidae Longarm mullet (Osteomugil (Mugil) cunnesius)

Indo-Pacific (Kuwait)

N Y Y Y N N Y Y N Almeida et al., 1968; Ovcharenko, 2016; NZFungi, 2019

Eubacterium tarantellae



N Y (sea lions)

Y Y N N Y N N Austin & Austin, 2007; Buller, 2014; Austin, 2005

Enterococcaceae

Lactococcus garviae (= Enterococcus seriolicida)

Loliginidae, Octopodidae

Squid (Loligo sp.) Octopus (Octopus vulgaris)

Northeast Pacific N Y Y Y N N Y Y N Wang et al., 2007; Farto et al., 2014; DermNet NZ, 2014

Lactococcus garviae (cont.)


Northwest Pacific (Chinese Taipei)

N Y Y Y (zoonotic) N N Y Y N Chen et al., 2002; Wang et al., 2007; NZOR, 2019

Chlamydiaceae

Epitheliocystis (Chlamydia-like intracellular disease)

Carangidae Greater amberjack (Seriola dummerili), greenback horse mackerel (Trachurus declivus), yellowtail amberjack (Seriola lalandi), Japanese amberjack (S. quinqueradiata)

Northwest Pacific (Australia, Japan), Southeast Pacific (South America)

N Y Y N N N N Y N Nowak & LaPatra, 2006; Stride & Nowak, 2013; Georgiades et al., 2016

Epitheliocystis spp. (cont.)

Clupeidae Pacific herring (Clupea pallasii pallasii), South American pilchard (Sardinops sagax)

Central Pacific (Australia)

N Y Y N N N N Y N Nowak & LaPatra, 2006; Stride & Nowak, 2013; Georgiades et al., 2016

Bacteroidetes: Flavobacteriaceae

Flavobacterium sp. Loliginidae Lolliguncula (Loligo) brevis Europe N Y Y N N N Y N N Ford et al., 1986; Farto et al., 2014; DermNet NZ, 2014

Tenacibaculum maritimum (Flexibacter maritimus, Cytophaga marina)

Clupeidae Pacific sardine (Sardinops sagax)

Mediterranean (Turkey), Northwest Pacific (Japan), North east Pacific (California)

N Y Y Y N N Y Y N Chen et al., 1995; Anderson, 1996; McGladdery, 1999; Santos et al., 1999; Brosnahan et al., 2017

Tenacibaculum maritimum (cont.)

Engraulidae Anchovy (Engraulis sp.), northern anchovy (E. mordax)

Northeast Atlantic (Europe), South Pacific

N Y Y Y N N Y Y N Diggles, 2011; Ozturk & Altinok, 2014; Diggles, 2016; Brosnahan et al., 2018a









Zoonotic potential



Reference

Gammaproteobacteria: Francisellaceae

Francisella spp. Scombridae Atlantic mackerel (Scomber scombrus)

Widespread N N Y Y N N/A Y Y Y Johnston, 2008; Colquhoun & Duodu, 2011

Gammaproteobacteria: Halomonadaceae

Halomonas aquamarina (= Achromobacter aquamarines), H. superficialis

Muglidae Flathead grey mullet (Mugil cephalus)

West Atlantic (USA, Gulf of Mexico)

N N Y N N N/A N Y N Almeida et al., 1968; Paperna & Overstreet, 1981; NZFungi, 2019

Shewanella spp. Octopodidae Octopus vulgaris Northwest Atlantic, Mediterranean

N Y Y N N N Y Y N Farto et al., 2014

Micrococcinaceae: Micrococcaceae

Micrococcus spp. Loliginidae Loligo forbesii, Sepioteuthis lessoniana

Northwest Atlantic, Mediterraneam

N Y Y N N N Y Y N Fiorito et al., 2015; DermNet NZ, 2014

Staphylococcus sp. Loliginidae Lolliguncula brevis Northwest Atlantic, Mediterranean

N Y Y N N N Y Y N Ford et al., 1986; Farto et al., 2014

Renibacterium salmoninarum

Clupeidae Atlantic herring (Clupea harengus) (by experimental infection, not present in wild stocks)

Widespread N N Y Y N N/A N Y N Evelyn, 1993; Wiens, 2011; Hershberger et al., 2013

Cornyebacteriaceae: Mycobacteriaceae

Mycobacterium (=Myxobacterium sp., Mycobacterium marinum, M. fortuitum))

Loliginidae Doryteuthis (Amerigo) pealeii (=Loligo pealei)

Widespread, Europe, Mediterranean

N Y Y N N N Y Y N Fiorito et al., 2015; DermNet NZ, 2014

Mycobacterium spp. (cont.)


Northeast Atlantic (Europe), Mediterranean

N Y Y Y (zoonotic)

N N Y Y N Paperna & Overstreet, 1981; Ngan et al., 2005; Varello et al., 2014; Ovcharenko, 2016; NZ Fungi, 2019

Spirochaetales: Leptospira

Leptospira icterohaemorhagiae

Muglidae Flathead grey mullet (Mugil cephalus)

West Atlantic (Gulf of Mexico)

N Y Y N N N Y Y N Paperna & Overstreet, 1981; Ovcharenko, 2016; Anon., 2012; MOH, 2019

Moritellaceae

Moritella (Vibrio) viscosa (winter ulcer disease)


Widespread N N Y N N N/A N Y N Austin & Austin, 2007; Tubbs et al., 2007; Björnsdóttir, 2011; Buller, 2014









Zoonotic potential



Reference

Pseudomonadaceae

Pseudomonas anguilliseptica (Serotype 2 isolates)

Clupeidae Atlantic (Baltic) herring (Clupea harengus membras)

Widespread N N Y Y N N/A Y Y Y Lonnstrom et al., 1994

Pseudomonas stutzeri Octopodidae Octopus (Octopus briareus, O. joubini)

Northwest Atlantic, Mediterranean

N Y Y Y N N Y Y N Ford et al., 1986; Sherrill et al., 2000

P. stutzeri (cont.) Sepiidae Sepia officinalis Northwest Atlantic, Mediterranean

N Y Y Y N N Y Y N Ford et al., 1986; Sherrill et al., 2000

Pseudomonas sp. Mugilidae Flathead grey mullet (Mugil cephalus)

Northeast Atlantic (Europe)

N Y Y Y N N Y Y N Lewis et al., 1970; Paperna & Overstreet, 1981; Ovcharenko, 2016; NZFungi, 2019

Pseudomonas sp. (cont.)

Octopodidae Octopus bimaculoides, Enteroctopus dofleini


N Y Y N N N Y Y N Stoskopf et al., 1987; Fiorito et al., 2015; NZFungi, 2019

Pseudomonas sp. (cont.)

Sepiidae Loligo forbesii, Lolliguncula brevis, Sepioteuthis sp., Lessoniana sp.


N Y Y N N N Y Y N Hanlon et al., 1984; Ford et al., 1986; Sherrill et al., 2000; Fiorito et al., 2015

Streptococcaceae

Streptococcus agalactiae (Group B type 283)

Clupeidae Pacific sardine (Sardinops sagax)

Indo-Pacific (Singapore, Japan, Kuwait)

N N Y Y Y N/A Y Y NZOR, 2017; Odhiambo et al., 2018

Streptococcus agalactiae (Group B type 283) (cont.)

Mugilidae Klunzinger's mullet (Liza klunzingeri), flathead grey mullet (Mugil cephalus)

Indo-Pacific (Singapore, Japan, Kuwait)

N N Y Y Y N/A Y Y Y Bunch & Bejerano, 1997; Evans et al., 2002; NZOR, 2017

Streptococcus iniae Clupeidae Sardine (Sardinops sagax (= S. melanostictus))

Widespread N N Y Y N N/A Y Y Y Minami et al.,1979; Salati, 2011

S. iniae (cont.) Engraulidae Anchovy (Engraulis japonicus) (= E. japonica)


N N Y Y N N/A Y Y Y Minami et al.,1979

S. iniae (cont.) Mugilidae Flathead grey mullet (Mugil cephalus)

Widespread N N Y Y N N/A Y Y Y Johnston, 2008; Kluzik & Woodford, 2016

Streptococcus sp. Mugilidae Flathead grey mullet (Mugil cephalus), thinlip grey mullet (Chelon (Liza) ramada)

Northwest Atlantic (United States), Mediterranean, Northwest Pacific (Chinese Taipei)

N Y Y Y N N Y Y N Plumb et al., 1974; Wang et al., 2001; Chen et al., 2002; Ovcharenko, 2016

Vibrionaceae: Photobacteriaceae

Photobacterium damsela (= P. damsela piscida)

Carangidae Mediterranean horse mackerel (Trachurus mediterraneus)

Mediterranean (Turkey) Widespread

N Y Y Y N N Y Y N Tanrikul & Cagirgan, 2001; Ozturk & Altinok, 2014; NZ Fungi, 2017









Zoonotic potential



Reference

Photobacterium damsela (= P. damsela piscida) (cont.)


Mediterranean (Turkey) Widespread

N Y Y Y N N Y Y N Lewis et al., 1970; Paperna & Overstreet. 1981; Black, 1997; Ozturk & Altinok, 2014; NZ Fungi, 2017

Vibrionaceae: Vibrio

Vibrio alginolyticus, V. carchariae, V. harveyi, V. parahaemolyticus, V. vulnificus

Carangidae Mediterranean horse mackerel (Trachurus mediterraneus), Atlantic horse mackerel (T. trachurus)

Widespread N Y Y Y N N Y Y N Ozturk & Altinok, 2014; NZOR, 2017

Vibrio spp. (cont.) Mugilidae Flathead grey mullet (Mugil cephalus)

Widespread N Y Y Y N N Y Y N Paperna & Overstreet, 1981

Vibrio spp., V. anguillarum, V. lentus, V. pelagicus, V. splendidus, V. neptunius

Mugilidae Flathead grey mullet (Mugil cephalus), Lebranche mullet (Mugil liza)

Widespread N Y Y Y N N Y Y N Burke & Rodgers, 1981; Paperna & Overstreet, 1981; De Sousa et al., 1999

Vibrio spp. (cont.) Sepiidae Sepia officinalis, Sepia spp., Sepioteuthis lessoniana, Loligo forbesii, Lolliguncula brevis


N Y Y N N N Y Y N Reimschuessel et al., 1990; Sherrill et al., 2000; Farto et al., 2003, 2014; Fiorito et al., 2015; Fichi et al., 2015

Vibrio spp. (cont.) Octopodidae Octopus bimaculoides, O. O. briareus, O. maya, O. joubini, O. vulgaris


N Y Y N N N Y Y N Reimschuessel et al., 1990; Sherrill et al., 2000; Farto et al., 2003, 2014; Fiorito et al., 2015; Fichi et al., 2015

Cnidaria: Myxozoa

Alataspora serenum Carangidae European horse mackerel (Trachurus trachurus)

Northeast Atlantic, Irish Sea

N N Y N (coelozoic) N/A Y N Y N Campbell, 2005

Alataspora solomoni Carangidae European horse mackerel (Trachurus trachurus), Mediterranean horse mackerel (T. mediterraneus)

Black Sea, Mediterranean

N N Y N (coelozoic) N/A Y N Y N Iurakhno, 1988; Campbell, 2005

Alataspora sp. Mugilidae Thinlip grey mullet (Chelon (Liza) ramada)

Mediterranean N N Y N (coelozoic) N/A Y N Y N Campbell, 2005; Yurakhno & Ovcharenko, 2014

Ceratomyxa auerbachi Clupeidae Atlantic herring (Clupea harengus), Pacific herring (C. pallasi pallasi)

Widespread N N Y N (coelozoic) N/A Y N Y N Rahimian, 2007; Koie et al., 2008

Ceratomyxa australis Carangidae Cape horse mackerel (Trachurus capensis)

Southeast Atlantic (Namibia)

N N Y N (coelozoic) N/A Y N Y N Gaevskaya & Kovaleva, 1979; Campbell, 2005; Sarkar, 2010

Ceratomyxa sardinellae


Indo-Pacific (Bay of Bengal)

N N Y N (coelozoic) N/A Y N Y N Sarkar, 2010

Chloromyxum kotorensis

Mugilidae Golden grey mullet (Chelon (Mugil) auratus (= Liza aurata)

Mediterranean, Black Sea

N N Y N (coelozoic) N/A Y N Y N Lubat et al., 1989









Zoonotic potential



Reference

Chloromyxum marinum

Hemiramphidae Japanese halfbeak Hyporhamphus (Hemirhamphus) sajori


N N Y N (coelozoic) N/A Y N Y N Shulman, 1966

Davisia donecae Carangidae Atlantic horse mackerel (Trachurus trachurus), Cape horse mackerel (T. capensis)

Southeast Atlantic, (Namibia)

N N Y N (coelozoic) N/A Y N Y N Gaevskaya & Kovaleva, 1979; Campbell, 2005

Enteromyxum leei (=Myxidium leei)

Mugilidae Flathead grey mullet (Mugil cephalus) and 46 other species from 16 genera

East Atlantic, Mediterranean, West Pacific

N N Y Y N/A N N Y Y Montero et al., 2007

Henneguya ouakamensis


East Atlantic (Senegal),

N N Y N N/A Y N Y N Kpatcha et al., 1997; Whipps et al., 2003

Henneguya species 3 Mugilidae Flathead grey mullet (Mugil cephalus)

Southeast Atlantic (Senegal)

N N Y N N/A Y N Y N Faye et al., 1997

Kudoa azevedoi Carangidae Horse mackerel (Trachurus trachurus)

Northeast Atlantic, Mediterranean (Tunisia)

N N Y N N/A Y N Y N Moran et al., 1999a; Eiras et al., 2005; Mansour et al., 2014

Kudoa bora (= Chloromyxum bora)

Mugilidae Flathead grey mullet (Mugil cephalus) (= M. japonicus), keeled mullet (Liza carinata)

West Pacific (Chinese Taipei)

N N Y N N/A Y N Y N Fujita, 1930; Eiras et al., 2005; Ovcharenko, 2016

Kudoa caudata Scombridae Chub mackerel (Scomber japonicus)

Pacific Ocean N N Y N N/A Y N Y N Kovaleva & Gayevskaya, 1983; Li et al., 2013

Kudoa (= Chloromyxum) clupeidae

Clupeidae Alewife (Alosa pseudoharengus), Atlantic menhaden (Brevoortia tyrannus), European herring (Clupea harengus) and 9 other species

Northwest, Southwest Atlantic

N N Y Y N/A Y N Y Y Hahn, 1917; Meglitsch, 1947; Reimschuessel et al., 2003; Langdon, 2007

Kudoa haridasae Carangidae European horse mackerel (Trachurus trachurus)


N N Y N (coelozoic) N/A Y N Y N Sarkar & Ghosh, 1991; Campbell, 2005

Kudoa haridasae (cont.)

Mugilidae Goldspot mullet (Liza parisa)


N N Y N (coelozoic) N/A Y N Y N Sarkar & Ghosh, 1991; Campbell, 2005

Kudoa hexapunctata Scombridae Pacific bluefin tuna (Thunnus orientalis)


N N Y N N/A Y N Y N Quiazon, 2015; Kasai et al., 2017

Kudoa intestinalis Mugilidae Flathead grey mullet (Mugil cephalus)


N N Y N N/A Y N Y N Maeno et al., 1993; Moran et al., 1999a

Kudoa iwatai Mugilidae Flathead grey mullet (Mugil cephalus) and 19 other species

Mediterranean N N Y N N/A Y N Y Y Egusa & Shiomitsu, 1983; Diamant et al., 2005; Ovcharenko, 2016

Kudoa nova (= Kudoa quadratum)

Carangidae European horse mackerel (Trachurus trachurus) and 20 other species

North Atlantic (Norway, Bay of Biscay), Mediterranean

N N Y Y N/A Y N Y Y Lom & Dykova, 1992; Longshaw et al., 2004; Campbell, 2005









Zoonotic potential



Reference

Kudoa prunusi (Occurs in farmed fish, but rare in wild stocks)

Scombridae Pacific bluefin tuna (Thunnus orientalis)


N N Y N N/A Y N Y N Meng et al., 2011

Kudoa shiomitsui Scombridae Pacific bluefin tuna (Thunnus orientalis)


N N Y N NA Y N Y N Zhang et al., 2010

Kudoa tetraspora Mugilidae Flathead grey mullet (Mugil cephalus)

Indo-Pacific (India)

N N Y N N/A Y N Y N Narasimhamurti & Kalavati, 1979

Kudoa thyrsites Clupeidae South African pilchard (Sardinops sagax ocellatus), S. sagax neopilchardus (Western Australia), Sardinella lemuruand 37 other species in 18 families

Southwest, Northwest Pacific (Australia, Japan), South Atlantic (Africa), Northeast Atlantic (England, Europe), Southeast Pacific (Chile), Northeast Pacific (Canada, Alaska)

N N Y Y N/A Y N Y Y Munday et al., 1998; Whipps & Kent, 2006; Henning et al., 2013; Levsen, 2015

Kudoa thyrsites (cont.) Engraulidae Japanese anchovy (Engraulis japonicus) and 18 other hosts

Pacific (Australia, Japan)

N Y Y Y N Y N Y Y Stehr & Whittaker, 1986; Moran & Kent, 1992; Hine et al., 2000; Diggles, 2011

Kudoa trachuri Carangidae Japanese jack mackerel (Trachurus japonicus)

Northwest Pacific N N Y N N/A Y N Y N Matsukane et al., 2011

Kudoa trifolia Mugilidae Flathead grey mullet (Mugil cephalus), thinlip grey mullet (Chelon (Mugil) ramada), Golden grey mullet (C. (Mugil) auratus (= Liza aurata)

Indo-Pacific, Mediterranean

N N Y N N/A Y N Y N Holzer et al., 2006; Ovcharenko, 2016

Kudoa unicapsula Mugilidae Golden grey mullet (Chelon (Mugil) auratus =Liza aurata)

Mediterranean N N Y N N/A Y N Y N Yurakhno et al., 2007

Kudoa yasunagi Scombridae Pacific bluefin tuna (Thunnus orientalis)


N N Y N N/A Y N Y N Zhang et al., 2010

Kudoa sp. Carangidae Horse mackerel (Trachurus trachurus)

East Atlantic, Mediterranean

N N Y N N/A Y N Y N Campbell, 2005

Kudoa sp. Mugilidae Flathead grey mullet (Mugil cephalus), fringelip mullet (Crenimugil (Mugil) crenilabis)

Widespread, West Atlantic, Red Sea

N N Y N (coelozoic) N/A Y N Y N Paperna & Overstreet, 1981

Kudoa sp. (cont.) Octopodidae Paroctopus dofleini Japan N N Y N N/A Y N Y N Yokoyama & Masuda, 2001; Lom & Dykova, 2006

Microspora sp. (Microsporaceae)

Sepiidae Sepia sp. Widespread N Y N N N N N Y N Hochberg, 1983; Novis, 2004









Zoonotic potential



Reference

Myxidium incurvatum Mugilidae Flathead grey mullet (Mugil cephalus)

West Atlantic (USA), Northwest Pacific

N Y Y N N/A Y N Y N Jaysari & Hoffman, 1982; Paperna & Overstreet, 1981; Cairns et al., 2009

Myxobolus adeli Mugilidae Golden grey mullet (Chelon (Mugil) auratus (= Liza) aurata)

Mediterranean, Black Sea, Azov Sea

N N Y N (coelozoic) N Y N Y N Yurakhno & Ovcharenko, 2014

Myxobolus bizerti (= M. hannensis), M. goensis, M. nile


Indo-Pacific, Mediterranean

N N Y N N Y N Y N Bahri & Marques, 1996; Eiras & D’Sousa, 2004; Sharon et al., 2019

Myxobolus branchialis (= M. branchiale, Myxosoma branchialis)

Mugilidae Flathead grey mullet (Mugil cephalus), golden grey mullet (Chelon (Mugil) auratus (= Liza aurata), leaping mullet (C. (Liza, Mugil) saliens)

Black, Caspian Seas, Northwest Pacific

N N Y N (coelozoic)

N Y N Y N Shulman, 1966; Paperna & Overstreet, 1981; Ibragimov, 1987; Iskov, 1989; Yurakhno & Ovcharenko, 2014

Myxobolus cephalus (= M cephalis)


Atlantic Ocean, Gulf of Mexico

N N Y N N Y N Y N Lom & Dykova, 1992; Yurakhno & Ovcharenko, 2014

Myxobolus cheni Mugilidae Flathead grey mullet (Mugil cephalus), So-iuy mullet (Planiliza haematocheila (= M.l haematocheilus, M. soiuy))

Black Sea, Northwest Pacific (China, Japan, Russia)

N N Y N (coelozoic) N Y N Y N Shulman, 1966; Paperna & Overstreet, 1981; Yurakhno & Ovcharenko, 2014

Myxobolus chiungchowensis


Northwest Pacific, China

N N Y N (coelozoic) N Y N Y N Eiras et al., 2005

Myxobolus episquamalis


East Atlantic, Mediterranean, Pacific (Australia, Korea, Chinese Taipei)

N N Y N N Y N Y N Bahri et al., 2003; Rothwell et al., 1997; Kim et al., 2013b; Lane et al., 2014; Yurakhno & Ovcharenko, 2014

Myxobolus exiguus (=Myxosporidium mugilis)

Mugilidae Golden grey mullet (Chelon (Mugil) auratus) (= Liza aurata), flathead grey mullet (M.l cephalus), thicklip grey mullet (C. labrosus (= M. l chelo)), thinlip grey mullet (C. (Mugil) ramada)

Black Sea, Russia, Mediterranean

N N Y N N Y N Y N Paperna & Overstreet, 1981; Bahri et al., 2003

Myxobolus goensis Mugilidae Flathead grey mullet (Mugil cephalus)

Indo-Pacific (Indian Ocean)

N N Y N N Y N Y N Eiras & D’Sousa, 2004; Eiras et al., 2005

Myxobolus goreensis Mugilidae Flathead grey mullet (Mugil cephalus)

Indo-Pacific (Indian Ocean)

N N Y N N/A Y N Y N Fall et al., 1997; Yurakhno & Ovcharenko, 2014

Myxobolus improvisus Mugilidae Golden grey mullet (Chelon (Liza) auratus)

Mediterranean, Black Sea, Azov Sea

N N Y N (coelozoic) N/A Y N Y N Yurakhno & Ovcharenko, 2014









Zoonotic potential



Reference

Myxobolus incurvatum, M. rotundus


North, South Pacific (California, New Zealand)

N Y Y N N Y N Y N Jaysari & Hoffman, 1982; Hine & Diggles, 2000; Yurakhno & Ovcharenko, 2014

Myxobolus mugauratus (= M. nile)

Mugilidae Golden grey mullet (Chelon (Mugil) auratus (= Liza aurata))


N N Y N N/A Y N Y N Eiras et al., 2005; Yurakhno & Ovcharenko, 2014

Myxobolus mugcephalus



N N Y N N/A Y N Y N Eiras et al., 2005; Yurakhno & Ovcharenko, 2014

Myxobolus mugilis (= M. mugili)

Mugilidae Flathead grey mullet (Mugil cephalus), golden grey mullet (Chelon (Mugil) auratus (= Liza aurata)), thicklip grey mullet (C. labrosus (= Mugil chelo))

Mediterranean (Greece, Egypt), Indian Ocean

N N Y N N/A Y N Y N Parenzan, 1966; Paperna & Overstreet, 1981; Eiras et al., 2005

Myxobolus muelleri Mugilidae Flathead grey mullet (Mugil cephalus), golden grey mullet (Chelon (Mugil) auratus (= Liza aurata)), thinlip grey mullet (C. (Mugil) ramada)

Black Sea, Russia, Mediterranean (Italy, Tunisia), Southeast Atlantic (Senegal)

N N Y N N/A Y N Y N Paperna & Overstreet, 1981; Bahri et al., 2003; Yurakhno & Ovcharenko, 2014; Sharon et al., 2019

Myxobolus mugchelo Mugilidae Thicklip grey mullet (Chelon labrosus (= Mugil chelo)

Mediterranean N N Y N N/A Y N Y N Eiras et al., 2005; Yurakhno & Ovcharenko, 2014

Myxobolus parvus Mugilidae Flathead grey mullet (Mugil cephalus), golden grey mullet (Chelon (Mugil) auratus = Liza aurata)), so-iuy mullet (Planiliza (Liza) haematocheila (= M. l haematocheilus, M. soiuy))

Widespread N N Y N N/A Y N Y N Shulman, 1966; Paperna & Overstreet, 1981; Ovcharenko, 2015

Myxobolus raibauti Mugilidae Flathead grey mullet (Mugil cephalus)

Southeast Atlantic (Senegal)

N N Y N (coelozoic) N/A Y N Y N Eiras et al., 2005

Myxobolus rhodei Mugilidae Flathead grey mullet (Mugil cephalus)


N N Y N (coelozoic) N/A Y N Y N Eiras et al., 2005

Myxobolus rotundus Mugilidae Flathead grey mullet (Mugil cephalus), golden grey mullet (Chelon (Mugil) auratus =Liza aurata))

Black Sea, Pacific Ocean (California, New Zealand)

N Y Y N (coelozoic) N Y N Y N Chernova, 1967; Jaysari & Hoffman, 1982; Hine & Diggles, 2000; Yurakhno & Ovcharenko, 2014

Myxobolus saranai (= M. brachialis)

Mugilidae Golden grey mullet (Chelon (Mugil) auratus = Liza aurata)

Black Sea N N Y N N/A Y N Y N Paperna & Overstreet, 1981; Ovcharenko, 2015

Myxobolus spinacurvatura

Carangidae Horse mackerel (Trachurus trachurus)


N N Y N (coelozoic) N/A Y N Y N Bahri et al., 2003; Campbell, 2005

Myxobolus (= Myxosoma) sp.

Mugilidae Flathead grey mullet (Mugil cephalus), squaretail mullet (Ellochelon vaigiensis (= M.l waigensis))

Indo-Pacific (India), East Atlantic, Mediterranean

N N Y N (coelozoic) N/A Y N Y N Paperna & Overstreet, 1981









Zoonotic potential



Reference

Myxodavisia (Davisia) donecae

Carangidae Atlantic horse mackerel (Trachurus trachurus), Cape horse mackerel (T. capensis)

Southeast Atlantic (Namibia)

N N Y N (coelozoic) N/A Y N Y N Gaevskaya & Kovaleva,1979; Campbell, 2005

Myxodavisia (Davisia) haldare

Clupeidae Sardinella longiceps Indo-Pacific N N Y N (coelozoic) N/A Y N Y N Sarkar, 2010

Myxosoma branchialis Mugilidae Flathead grey mullet (Mugil cephalus)

Northwest Pacific N N Y N (coelozoic) N/A Y N Y N Paperna & Overstreet, 1981

Ortholinea (Sphaerospora) divergens


Black Sea N N Y N (coelozoic) N/A Y N Y N Yurakhno & Ovcharenko, 2014; Ozer et al., 2015a

Ortholinea orientalis Clupeidae Atlantic herring (Clupea harengus), Pacific herring (C. pallasii pallasii), sprat (Sprattus sprattus)

East Atlantic (Denmark) Northeast Pacific (Alaska)

N N Y N (coelozoic) N/A Y N Y N Marty et al., 1998; Karlsbakk & Koie, 2011

Parvicapsula minibicornis

Clupea European sprat (Sprattus sprattus), European herring (Clupea harengus)

Northeast Pacific N N Y N (coelozoic) N/A Y N Y N Jones et al., 2003; Atkinson et al., 2011; Jorgensen et al., 2011; Koie et al., 2013

Polysporoplasma mugilis

Mugilidae Golden grey mullet (Chelon (Mugil) auratus (= Liza aurata)), thinlip grey mullet (C (Mugil) ramada), thicklip grey mullet (C. (Mugil) labrosus)

Mediterranean, Black Sea

N N Y N (coelozoic) N/A Y N Y N Sitja-Bobadilla & Alvarez-Pellitero, 1995; Yurakhno, 2011; Yurakhno & Ovcharenko, 2014; Ovcharenko, 2016

Pseudalataspora pontica


Black Sea N N Y N (coelozoic) N/A Y N Y N Dmitrieva & Gaevskaya, 2001

Sphaeromyxa argentinensis

Engraulidae Argentine anchovy (Engraulis anchoita)

Southwest Atlantic (Argentina, Uruguay)

N N Y N (coelozoic) N/A Y N Y N Timi & Sardella, 1998

Sphaeromyxa balbianii Clupeidae Allis shad (Alosa sardina), European pilchard (Sardina (Clupea) pilchardus)

Widespread N N Y N (coelozoic) N/A Y N Y N Lom, 2004

Sphaeromyxa bonaerensis

Engraulidae Argentine anchovy (Engraulis anchoita)

Southwest Pacific (Argentina, Uruguay)

N N Y N (coelozoic) N/A Y N Y N Timi & Sardella, 1998

Sphaeromyxa parva Scomberosocidae Pacific saury (Cololabias saira)


N N Y N (coelozoic) N/A Y N Y N Lom, 2004

Sphaeromyxa (= Sphaeromyxum) sabrazesi

Mugilidae Flathead grey mullet (Mugil cephalus), golden grey mullet (Chelon (Mugil) auratus (= Liza aurata))

Black Sea, Western Atlantic, Mediterranean (Spain)

N N Y N (coelozoic) N/A Y N Y N Dmitrieva & Gaevskaya, 2001; Yurakhno & Ovcharenko, 2014









Zoonotic potential



Reference

Sphaerospora dicentrachi (=S. mugili)

Mugilidae Flathead grey mullet (Mugil cephalus), golden grey mullet (Chelon (Mugil) auratus (= Liza aurata)), leaping mullet (C. (Liza, Mugil) saliens), so-iuy mullet (Planiliza (Liza) haematocheila (= Liza haematocheilus, M. soiuyi)), thicklip grey mullet (C. (Mugil) labrosus), thinlip grey mullet (C. (Liza ramada)

East Atlantic, Mediterranean Black Sea

N N Y N (coelozoic) N/A Y N Y N Quaglio et al., 2002; Yurakhno & Ovcharenko, 2014

Sphaerospora testicularis

Clupea European sprat (Sprattus sprattus), European herring (Clupea harengus)

Northeast Atlantic, Mediterranean

N N Y N (coelozoic) N/A Y N Y N Atkinson et al., 2011; Koie et al., 2013

Zschokkella admiranda


Black, Mediterranean Seas

N N Y N N/A Y N Y N Yurakhno & Ovcharenko, 2014

Zschokkella dogieli Mugilidae Flathead grey mullet (Mugil cephalus), fringelip mullet (Crenimugil crenilabis)

Indo-Pacific (Japan to Australia)

N N Y N N/A Y N Y N Paperna & Overstreet, 1981

Zschokkella mugilis Mugilidae Flathead grey mullet (Mugil. cephalus), leaping mullet (Chelon (Liza, Mugil) saliens), (thinlip grey mullet (C. (Liza, Mugil) ramada), thicklip grey mullet (C. (Mugil) labrosus)

Mediterranean (Italy, Spain)

N N Y N (coelozoic) N/A Y N Y N Paperna & Overstreet, 1981; Lubat et al., 1989; Qualigo et al., 2002; Yurakhno & Ovcharenko, 2014

Ascomycetes

Cladosporium sp. Octopodidae Eledone cirrhosa Widespread N Y N N N N N Y N Polglase et al., 1984; Simeca & Oestmann, 1995; Harms et al., 2006; Fiorito et al., 2015; NZ Fungi, 2019; Polglase, 2019

Cladosporium sp. (cont.)

Sepiidae European cuttlefish (Sepia officinalis)

Widespread N Y N N N N N Y N Polglase et al., 1984; Simeca & Oestmann, 1995; Harms et al., 2006; Fiorito et al., 2015; NZ Fungi, 2019; Polglase, 2019

Labyrinthula sp. Marine slime mould

Octopodidae Octopus vulgaris Northwest Atlantic, Mediterranean

N Y N N (in aquaria)

N N N Y N Fiorito et al., 2015; Armiger, 1964

Ulkenia amoeboidea Octopodidae Eledone cirrhosa, Octopus vulgaris

Widespread N N N N N/A N N Y N Polglase, 2019; Fiorito et al., 2015; NZFungi, 2019









Zoonotic potential



Reference

Schizochytrium sp. Illicinidae Illex illicibrosus Canada N Y N N N N N Y N Jones, 1981; Jones & O’Dor, 1983; NZ Fungi, 2019; Polglase, 2019

Ichthyobodo necator Octopodidae Octopus sp. Northeast Pacific N N N N N/A N Y Y N Forsythe et al., 1991; Poynton et al., 2009

Chromidina coronata Octopodidae Octopus vulgaris, Eledone cirrhosa, Scaeurgus unicirrhus

Mediterranean, Northeastern Atlantic

N N N Y N/A Y N Y N Hochberg, 1983; Souidenne & Furuya, 2019

C. coronate (cont.) Sepiidae Sepiola rondeletii Mediterranean, Northeast Atlantic


C. coronate (cont.) Ommastrephidae Illex coindetiii Mediterranean, Northeast Atlantic


Chromidina elegans Octopodidae Octopus salutii, Mediterranean, Northeast Atlantic


C. elegans (cont.) Sepiidae Sepia elegans, S. orbignyana, Ilex coindetti,

North Pacific N N N Y N/A Y N Y N Hochberg, 1983; Souidenne & Furuya, 2019

C. elegans (cont.) Ilicinidae Dosidicus spp., Toderodes saggitatus

North Pacific N N N Y N/A Y N Y N Hochberg, 1983; Souidenne & Furuya, 2019

Opalinopsis sepiolae (= Opalinopsis octopi)

Octopodidae Octopus macropus, O. tetracirrhus

Mediterranean, Northeast Atlantic

N N N N N/A Y N Y N Hochberg, 1983; Souidenne & Furuya, 2019

O. sepiolae (cont.) Sepiidae Rossia macrosoma, Sepietta oweniana, Sepia rondeletii, S. elegans, Sepiola atlantica

Mediterranean, Notheast Atlantic

N N N N N/A Y N Y N Hochberg, 1983; Souidenne & Furuya, 2019

Conocyema polymorpha

Octopodidae Octopus vulgaris (= O. sinensis)


N N N N N/A Y N Y N Furuya & Souidenne, 2019

Aggregata andresi Octopodidae Martialia hyadesi Southwest Atlantic

N N N Y N/A Y N Y N Gestal et al., 2005

Aggregata octopiana, A. spinosa, Aggregata spp.


Mediterranean, Northwest Atlantic

N N N N N/A Y N Y N Gestal et al., 2002a, 2002b, 2007; Castellanos-Martinez et al., 2013, 2019

Aggregata bathytherma

Octopodidae Muusoctopus hydrothermalis (=Vulcanoctopus hydrothermalis)

Northeast Pacific N N N N N/A Y N Y N Gestal et al., 2010

Aggregata dobelli (Not in WoRMS 2019)

Octopodidae Enteroctopus dofleini Northeast Pacific N N N N N/A Y N Y N Poynton et al., 2002

Aggregata eberthi Sepiidae Sepia officinalis Mediterranean, Northeast Atlantic

N N N N N/A Y N Y N Castellanos-Martinez et al., 2013

Aggregata kudoi Sepiidae Sepia elliptica Northwest Indian Ocean

N N N N N/A Y N Y N Narasimhamurti, 1979

Aggregata millerorum (Not in WoRMS 2019)

Octopodidae Octopus bimaculoides Northeast Pacific N N N N N/A Y N Y N Poynton et al., 1992









Zoonotic potential



Reference

Aggregata saggitatus, Aggregata sp.

Ommastrephidae Todarodes sagittatus Northeast and Western Atlantic, Caribbean

N Y N N N Y N Y N Hochberg & Couch, 1971; Simeca & Oestmann, 1995; Gestal et al.,2002a, 2002b; Diggles et al., 2002; Castellanos-Martinez et al., 2013, 2019

Aggregata spp. Octopodidae Patagonian octopus (Octopus tehuelchus), Southern red octopus (Enteroctopus megalocyathus)

Southeast Atlantic N N N N N/A N N Y N Sardella et al., 2000

Dicyema paradoxum, D. typus, Dicyemennea lameerei


Mediterranean, Northwest Atlantic

N N N N N/A N N Y N Furuya & Souidenne, 2019; Gleadill 2016

Pleodicyema delamaeri

Octopodidae Bathypolypus sponsalis Mediterranean, Northwest Atlantic

N N N N N/A N N Y N Furuya & Souidenne, 2019

Dicyemennea eledones (=Dicyemodeca eledones), D. lameerei

Octopodidae Eledone cirrhosa Mediterranean, Northeast Atlantic

N N N N N/A N N Y N Souidenne et al., 2016; Furuya & Souidenne, 2019

D. moschatum, D. eledones (cont.)

Octopodidae Eledone (Octopus) moschata



D. macrocephalum Octopodidae Macrotritopus defilippi (=Octopus defilippi)



D. macrocephalum (cont.)

Sepiidae Rondeletiola minor, Sepietta obscura, S. oweniana, Sepiola steenstrupiana, Sepia elegans



D. paradoxum Octopodidae Octopus macropus Mediterranean, Northeast Atlantic


D. rondaletiolae Sepiidae Sepietta neglecta, S. oweniana



D. banyulensis, D. benedeni, D. eledones

Octopodidae Octopus salutii Mediterranean, Northeast Atlantic


Dicyemennea gracile, Pseudicyema truncatum, D. whitmani, Microcyema vespa

Sepiidae Rossia macrosoma, Sepia officinalis, S. orbignyana



Symbols N/A Not applicable Shaded pathogens are retained for risk analysis Viable in frozen fish bait as defined in the current IHS for fish bait (MPI, 2011)

http://www.marinespecies.org/aphia.php?p=taxdetails&id=534126

http://www.marinespecies.org/aphia.php?p=taxdetails&id=534126

Import risk analysis: Frozen fish and coleoid cephalopod molluscs for bait 35

5.1 Hazard list for risk assessment

As per the findings of the hazard identification in Table 6 the following organisms were

identified as hazards associated with fish bait. Only nervous necrosis virus (NNV) was

associated with coleoid cephalopod molluscs. All organisms identified as hazards were subject to

a risk assessment.

Viral pathogens

Birnaviridae: Marine aquabirnavirus (MABV) including ‘IPNV-like’ strains

Iridoviridae: Red sea bream iridovirus (RSIV), Infectious spleen and kidney necrosis virus

(ISKNV), heart and skeletal muscle inflammation (HSMI) and associated iridoviruses;

Erythrocytic necrosis virus (ENV)/Viral erythrocytic necrosis (VEN).

Nodaviridae-Betanodavirus: nervous necrosis virus (NNV)/ viral nervous necrosis (VNN)/

viral encephalopathy and retinopathy (VER)

Orthomyxoviridae-Isavirus: Infectious salmon anaemia virus (ISAV)

Reoviridae-Aquareovirus: Piscine aquareovirus (PRV)

Rhabdoviridae: Infectious haematopoietic necrosis virus (IHNV), Viral haemorrhagic

septicaemia virus (VHSV)

Bacterial pathogens

Enterobacteriaceae: Edwardsiella spp.

Gammaproteobacteriaceae: Francisella spp.

Pseudomonadaceae: Pseudomonas anguilliseptica

Streptococcaceae: Streptococcus agalacteae, Streptococcus iniae

Myxozoan pathogens

Enteromyxum leei, Kudoa clupeidae, K. iwatai, K. nova, K. thyrsites

General considerations

6.1 Risk assessment considerations

The following presents general, non-disease specific information that was taken into

consideration in the risk assessment.

For the purposes of this risk analysis, fish bait is non-viable. It has either been frozen (to at least

-18 °C or -20 °C, for at least 168 hours (7 days)) consistent with (Johnston, 2008; USDA, 2019);

or, irradiated with 25 kGy (=2.5 Mrads) consistent with MP (2011). It is wild-sourced, caught in

marine waters (MPI, 2011) and limited to the species defined in the Commodity Definition

(Table 5).


Fish bait has been derived from wild populations of marine finfish and cephalopods, which may

harbour pathogens. This commodity is recognised as an open or poorly regulated pathway for

pathogens through international trade (Biosecurity Australia, 2002; Dalton, 2004; Goodwin et

al., 2004; Arkush et al., 2006; Oidtmann et al., 2011; Phelps et al., 2013, 2014). As a traded

commodity, fish bait is caught, transported and stored in bulk, with no procedural steps (no

international standards) in the supply chain targeting pathogen reduction (Phelps et al., 2013,

2014; OIE, 2019b). Similarly, fish bait is imported in bulk and is not subject to any pathogen

reduction steps in the supply chain before entry into the aquatic environment.

Little information is available on the prevalence of most aquatic animal diseases in wild marine

fish stocks (Peeler et al., 2007; Peeler & Taylor, 2011), particularly for low value species such as

most fish bait (Dunn et al., 2012).

6.1.1 Entry assessment

A qualitative assessment of the likelihood of pathogen entry through the imported commodity is

provided for each identified hazard. Indicators may include, but are not limited to, the volume of

imports of the host species, the geographical distribution of the pathogen and the host species, as

well as the prevalence and persistence of infection in the host. The influence of climate change in

determining and extending the range of both pathogen and host is poorly understood (Harvell et

al., 2009; Burge et al., 2014).

6.1.2 Exposure assessment

Bait fish and coleoid cephalopods are schooling species (Maes & Ollevier, 2003; Merino et al.,

2014). They may be more susceptible to disease transmitted through close contact than solitary

fish species (Burge et al., 2014), but factors affecting the establishment of exotic pathogenic

organisms in susceptible aquatic hosts are poorly known (Munro et al., 1983; Peeler et al., 2007;

Jereb et al., 2010; Peeler & Thomas, 2011; Dunn et al., 2012 Engelhard et al., 2014).

Indicators affecting the assessment of likely exposure, establishment and spread include, but are

not limited to, persistence of the pathogen in the aquatic environment, infective dose, routes of

infection and methods of transmission (including vectors), host range (including conspecifics

and reservoir organisms), host density and geographical distribution, susceptible host life stages

and presence of obligate intermediate hosts (Phelps et al., 2013).

Exposure may also be influenced by natural risk mitigation features. These include abiotic

factors such as water temperature, dilution (of pathogen concentration below the infectious dose)

and distribution of the infective agent by ocean current hydrodynamics (Phelps et al., 2013;

Burge et al., 2014), while climate driven temperature changes may influence the host immune

response and affect the frequency of disease (Bowden, 2008; Burge et al., 2014).

While marine waters do provide a significant dilution factor, the distribution of both fish hosts

and pathogenic agents is not random (Suttle, 2007; Oidtmann et al. 2017). The fish hosts have

evolved behavioural responses to a complex set of coastal currents (Hume et al., 1992). Fishing

activity is also not random but is strongly directed to maximise contact with fish species (FAO,

2018). It is reasonable to assume that with the high number of fishhook sets, with an estimated

bait-loss of up to 50% (Skud et al., 1978) and the use of berley as a fish attractant (Wynn-Jones

et al., 2019), at least some infected fish bait may contact a suitable host species and allow a

pathogen to complete its life cycle.


Biotic factors that may provide natural mitigation are complex. Available data for many marine

pathogens are usually confined to reported presence in a host. Comparatively little is known

about the epidemiology of aquatic infectious diseases compared to terrestrial vertebrate diseases

(Kermack & McKendrick, 1991; Burge et al., 2014). The processes leading to epizootics are

poorly understood (Bidegain et al., 2016).

Available data on host susceptibility, morbidity, mortality, or infectious dose have generally

been determined from experimental studies. Where studies have used unnatural infection

pathways (such as intra-peritoneal injection) the results must be treated with caution because

such data may not reflect the likely natural exposure potential in marine waters. Where study

methods reasonably simulate natural exposure pathways (such as infection per os),

epidemiological data (such as susceptibility of host species) the results are more reliable (OIE,

2019a).

Laboratory-based estimates of pathogen virulence and survival should be interpreted as

maximum estimates. They do not adequately take into consideration the influence of the aquatic

environment, including dilution factors, or predation. For example, viral and bacterial pathogens

may be consumed by protozoans in marine waters while infected host fish may be consumed by

predators before the pathogen life cycle is complete (Burge et al., 2014; Hallett et al., 2015;

Jones et al., 2015). Even where disease establishment occurs more quickly, infections may be

unapparent or unnoticed in wild fish stocks (Sindermann, 1987; Jones et al., 2015). This is most

likely to occur for diseases of lower economic importance, or for lower value host species (Vike

et al., 2014; Burge et al., 2014).

Given these uncertainties, the likelihood of exposure and establishment for pathogens associated

with fish bait has been capped as “medium” to better reflect the associated risk.

The potential distribution pathways whereby viable pathogens associated with fish bait can directly

enter natural waters are shown in Figure 2. In addition to the intended end use as fish bait, other

potential pathways include diversion of imported fish bait to other end uses (e.g. fish food) or

waste from further processing.

A smaller volume is destined for retail sale. This may be part-thawed and re-packaged through the

wholesale and retail distribution process, prior to retail sale. Pathogens may remain viable in liquid

wastes from thawing during this process and enter the aquatic environment. Where disposed of to

landfill, leachates and effluent harbouring pathogens may also enter aquatic environments either

directly or be dispersed through the action of piscivorous and scavenging birds (Blackwell, 2019).

The relative importance assigned to each pathway is shown in Figure 2, categorised as probable,

less significant or unlikely. These pathways are considered where relevant for each identified

pathogen. The likelihood of introduction of significant pathogens through redirection of fish from

the human consumption pathway (Blackwell, 2019) is likely to be very low and is not considered

further in this risk analysis.


Figure 2. Potential distribution pathways for frozen fish or coleoid cephalopod mollusc fish bait

New Zealand marine fish aquaculture is largely focused on Chinook (King) salmon

(Oncorhynchus tshawytscha) (Family Salmonidae) on-grown in sea-cages. Chinook salmon

exports alone were valued at NZ$77 million in 2018 (Seafood New Zealand, 2019). Salmonid

aquaculture in New Zealand exclusively uses formulated fish food (Seafood New Zealand,

2019).

Wild fish species may become infected through imported fish bait. These may enter the sea

cages and be consumed by salmon. Recreational fishing activity commonly occurs immediately

adjacent to these sea cages, as these act as fish aggregation devices (Fisheries New Zealand,

2020). Discarded fish bait may also drift into a marine farm and recreational fishers may discard

their unused fish bait into the fish farm.

Other developing marine aquaculture fish species in New Zealand include snapper (Pagrus

auratus) (Family Sparidae), yellowtail kingfish (Seriola lalandi) (Family Carangidae), hapuku

(Polyprion oxygeneios) (Polyprionidae) (NIWA, 2020; Plant & Food, 2020). These developing

fisheries may use imported fish bait as fish food, as no artificial fish foods have yet been

developed. These activities represent viable risk pathways between wild fisheries and

commercial aquaculture.

6.1.3 Consequence assessment

A qualitative assessment of the consequences of pathogen exposure through the imported

commodity is provided for each identified hazard. This considers the impact on people

(including zoonotic potential, societal values and cultural values), the New Zealand environment

and the New Zealand economy (Biosecurity Act 1993).


The economic consequences are comparatively easy to determine. Direct effects include, but are

not limited to, loss of volume or value of export trade, reduced productivity, increased mortality,

or reduced marketability of infected stocks (Dobson & May 1987). Indirect effects include

restrictions on New Zealand exports and trade access, as well as the costs of mitigation measures

and disease control processes (Biosecurity New Zealand 2006).

Little published information is available to determine the social and environmental consequences

of aquatic pathogen establishment (Munro et al. 1983; Peeler & Taylor 2011; Groner et al.

2016). Social effects include, but are not limited to, the direct consequences for human health of

zoonotic pathogens, the effects on traditional Maori customary fishing, cultural, spiritual

environmental and economic values, other cultural values and perceptions, as well as the effect

on recreational fishing activity. Indirect social effects include changed perceptions or job losses

in associated industries (such as tourism) resulting from the introduction of a risk organism

(Biosecurity New Zealand 2006). Indicators may be from non-market analysis (Southwick et al.

2018) or changes in social perception of amenity values (Biosecurity New Zealand 2006).

Environmental consequences may be direct or indirect. Direct consequences on indigenous fauna

include, but are not limited to, increased mortality and lower productivity, muscle degeneration,

liver dysfunction, interference with nutrition, cardiac disruption, nervous system involvement,

castration or mechanical interference with spawning, weight loss, and gross distortion of the

body (Sindermann 1987). These effects may only be apparent in wild fish stocks during

epizootics, due to high rates of predation on sick fish and scavenging of dead fish.

Indirect environmental effects may result from pathogen-induced changes in food webs. Bait fish

species such as clupeids feed on phytoplankton, zooplankton and detritus and represent a major

link between trophic levels in marine food webs. Bait fish also provide a significant food source

for other fish, seabirds and marine mammals (Dunn et al. 2012).

6.2 Risk management considerations

Risk management measures may be applied to reduce the risk associated with a specific risk

organism. These can be applied either pre-border (offshore) or post-arrival (where the shipment

is held in a transitional facility). Risk management measures are aimed at a particular point of the

risk pathway, typically the likelihood of entry.

Risk management measures may be applied singly or in combination with other measures.

The measures applied need to be commensurate with the perceived risk. Not all risk management

measures proposed may need to be applied at the present level of risk. Other identified measures

may be applied if and when the perceived level of risk changes.

6.2.1 General risk management measures

Identification of the species included in the commodity

This is intended to identify fish bait species that are not associated with any identified risk

organisms.

Declaration of the country of capture

Not all fish bait is caught within the territorial seas of a fishing nation. Some species such as

squid are taken in international waters. The declaration of country of capture is intended to

https://besjournals.onlinelibrary.wiley.com/doi/full/10.1111/j.1365-2664.2005.01043.x#b2


establish that the consignment has been sourced from areas where the risk organism has not

previously been reported.

Declaration that the commodity is sourced from wild stock

Fish bait species are of relatively low value. Species including flathead grey mullet (Mugil

cephalus) are farmed both for human consumption and to supply the international demand for

fish food and fish bait in the Mediterranean, United States and Southeast Asia (FAO, 2020b). It

is likely that the prevalence of infection in farmed fish bait may be higher than the assumed

prevalence of 2% to 4% generally associated with wild fish stocks (Corsin et al., 2009).

Declaration that the commodity has not been harvested from populations experiencing an

epizootic disease.

Fish bait species commonly occur in schools and wild stocks are subject to epizootics where risk

organisms may be endemic. The prevalence of infection during an epizootic may exceed 50%

(Reno et al., 1985; de la Pena et al., 2011). Where the commodity is used for human

consumption, diseased fish may be redirected to the fish bait pathway as a common method of

disposal. It is important to ensure that the commodity should only comprise healthy fish.

Declaration that the commodity has no visible signs of disease.

This implies the fish have been inspected under the oversight of the Competent Authority, where

appropriate standards have been established and followed. This does not mean actual inspection

is necessary.

Declaration of frozen storage

The current IHS for fish bait (MPI. 2011) requires a declaration that the commodity has been

frozen (to below -18 °C to -20 °C for at least 18 hours). The risk analysis for filleted

Oreochromis sp. (Johnston, 2008) proposed a minimum frozen storage period of at least 7 days

(168 hours, at -18°C to -20°C). Quick freezing and frozen storage will denature most metazoan

organisms (Hamed & Elias, 1970; Hauck, 1977; Barlow & Sleigh, 1979; Paperna & Overstreet,

1981; McVicar, 1982; Alderman & Polgase, 1986; Fayer & Nerad, 1996; Morris et al., 1998;

Olson et al., 1999; Shaw et al., 2000; Oidtmann et al., 2002; Racz & Zemankovics, 2002;

Johnston, 2008; Gregg et al., 2012; Leiro et al., 2012; Noga, 2012; Phelps et al., 2013;

Hershberger et al., 2015; Blackwell, 2019).

While the mechanisms of cell death are poorly understood (Pegg, 1987), factors include the

formation of ice crystals within and between cells, cell shrinkage, the destruction of cell

organelles and the concentrations of cellular metabolites. Ice crystals destroy external cell

membranes resulting in the loss of cellular integrity causing cell shrinkage. Ice crystals also

cause damage to internal cell membranes and disrupt organelle function. Ice formation also

dehydrates the cytoplasm, concentrating cell metabolites. These may reach toxic levels resulting

in cell death (Pegg, 1987; OHSU, 2020).

Viability may decrease over time, so extended frozen storage may be necessary to ensure

denaturation (Conlan et al., 2011). For many organisms, it is the freeze-thaw cycle, rather than

the length of freezing that leads to cell death (OHSU, 2020).

Most viruses and many gram-positive bacteria are unaffected by multiple freeze-thaw cycles and

extended frozen storage (Sourek, 1974; Meyers et al., 1999; Gaughan, 2001; Arkush et al., 2006;

Hervé-Claude et al. 2008; OIE, 2019b). Viruses are non-living and generally stable in

temperatures between + 20 °C and -80 °C, without alteration of their molecular structure


(Mortensen et al., 1998). Frozen storage (-20 C to -10°C) has been used for long-term

preservation of viruses in fish tissue (Plumb et al., 1973; Burke & Mulcahy 1983; Plumb &

Zilberg 1999) and for storage of pathogenic bacteria (Sourek, 1974).

The bacterium Edwardsiella spp. and myxozoan pathogens (Enteromyxum leei, Kudoa

clupeidae, K, iwatai, K. nova and K. thyrsites) are only slightly affected by short term (7 days)

frozen storage (Yurakhno. 2017; Castro et al., 2006; Johnston, 2008), but are substantially or

completely denatured following extended (at least 4 months) frozen storage (Brady &

Vinitnantharat, 1990; Arkush et al., 2013; Hoffman & Putz, 2011; Ohnishi et al., 2016; Anon.

2020b).

For the purposes of this risk analysis, it is proposed that the period of frozen storage be extended

to at least 4 months (17 weeks), for all fish bait, to denature Francisella spp., Enteromyxum leei,

Kudoa clupeidae, K. iwatai. K. nova and K. thyrsites. This may be achieved by either:

(a) a declaration that the commodity has been previously frozen for at least 4 months (17

weeks) at a temperature of -18 °C to -20 °C or below, from the date of capture; or,

(b) a requirement that the imported commodity must remain in frozen storage (at -18 °C to

-20 °C) in a BNZ-approved Transitional Facility, for this minimum time period, before

release.

Labelling the commodity as “Bait only, unfit for human consumption”.

While little actual control is possible post-border, the risk to human health can be reduced by

appropriate labelling of the commodity.

6.2.2 Pathogen specific risk management options

Sourcing fish bait from listed species not associated with pathogens of concern

Species prohibition includes identifying host species for identified pathogens of concern. From

the finfish and coleoid cephalopod molluscs identified as fish bait (Table 5) some species are

considered susceptible to identified risk organisms. Other fish bait species are unlikely to

represent a risk in the commodity.

Region/Country freedom

Fish bait may be sourced from regions or countries recognised by BNZ as being free of

corresponding pathogens of concern. Acceptance of region/ country freedom may be provided

through the BNZ Country Approval Procedures. For OIE-listed diseases, a certificate of country

freedom is provided by the Competent Authority of the exporting country. For non OIE-listed

diseases, acceptance of country freedom through the BNZ Country Approval Procedures would

be possible by a Health Certificate provided by the Competent Authority stating that the risk

organism had not been reported from the country of capture.

Pre-export or post-arrival irradiation (ionising radiation to 50 kGy)

Ionising radiation is commonly applied to fresh food commodities to extend shelf life by

denaturing food-spoiling bacteria (DAFF, 2013). Doses in the range of 1–10 kGy are unlikely to

change the nature of the commodity and 10 kGy is the maximum recommended for commodities

intended for human consumption (Anon., 1981, 1984; Crawford & Ruff, 1996). A minimum

dose of 3 kGy is recommended for frozen fish and shellfish product for human consumption

(IAEA, 2000). A dose of 1 kGy is recommended for denaturing microorganisms in fruit and

vegetables, and up to 30 kGy for bacterial decontamination of herbs and spices (FSANZ, 2020).


A dose of 10 kGy is equivalent to heat of energy of 10 J/kg and the heat capacity of water is 4.2

J/kg, therefore a dose of 10 kGy should raise the temperature by 2.4 °C (Loaharanu, 1997). For

food irradiation, a dose of 1 to 5 kGy is estimated to raise the temperature of the fish bait by 0.24

to 1.2 °C (Loaharanu, 1997).

Higher doses to 25 kGy are commonly used for the ‘bacterial sterilisation’ of medical equipment

and pharmaceuticals (DAFF, 2013) and as a biosecurity application to inactivate most bacterial,

fungal and metazoan pathogens likely to be present in imported fish and fish food (MPI, 2011).

Following Loraharanu (1997), a dose of 25 kGy would be expected to raise the product

temperature by 6 °C.

Viral pathogens present in frozen fish bait may require a higher dose to ensure denaturation.

Further, higher doses are generally required to achieve ‘radiation sterilisation’ and ensure viral

denaturation in frozen tissues, as compared to fresh or chilled commodities (DAFF, 2013). A

dose of 50 kGy, or the application of a 25 kGy dose twice is recommended for inactivation of

pathogens of biosecurity concern (DAFF, 2013). This seemingly high dose of radiation may be

suitable for frozen fish bait as the commodity is not intended for human consumption. Use of a

50 kGy dose could increase the product temperature by 12 °C (Loaharanu, 1997). However,

where the commodity is irradiated twice at 25 kGy (DAFF, 2013), the temperature increase

would remain at 6 °C. It is likely that the effect on frozen fish bait would be minimal.

It is acknowledged that no suitable irradiation facility is currently available in New Zealand.

However, an overseas facility may be approved by BNZ to undertake the irradiation of bait (N.

Ahmed, BNZ, pers. comm., 2020). Approved overseas irradiation facilities are used for

biosecurity irradiation by the Australian Government (DAWR, 2016).

Fish bait industry sector representatives (B. Burney, pers. comm., 2020; M. Lyford, pers. comm.,

2020) have expressed concerns that costs of overseas irradiation may make the product cost

prohibitive. Further, there is concern that irradiation may alter the nature of the product by

softening the tissues and reducing the volume of blood and fluids in the thawed product that

attract fish. Additional tests may be necessary to determine whether irradiated bait product

remains fit-for-purpose. Further, a lower irradiation dose may be appropriate to denature the risk

organisms associated with a particular fish bait species. This may be determined on a case-by-

case basis, as necessary. Irradiation, however, may be still be a viable risk management option

for the commodity.

Heat treatment

Heat treatment is a viable risk management option for fish for human consumption (OIE, 2019a;

Blackwell, 2019) or for processed fish food based on fish meal (Cobb, 2009). However, heating

alters the nature of the commodity, making it unfit for use as fish bait (B. Burney, pers. comm.,

2020; R. Clarke, pers. comm., 2020; M. Lyford, pers. comm., 2020). Heat treatment is not a

viable risk management option as the commodity definition is for frozen fish bait. Heat treatment

is not considered any further.

Chemical treatment

Surface disinfectant denature most risk organisms (Bovo et al., 2005; Yoshimizu et al., 2005;

Watanabe & Yoshimizu, 1998; OIE, 2019b). These chemicals have poor penetration and are

unsuitable for treatment of whole fish bait (Fraser et al., 2006; Phelps et al., 2013; Chua et al.,

2019).


Chemical bath processing methods are effective against bacteria and viruses present in whole

fish bait in laboratory trials (Faisal et al., 2012; Phelps et al., 2013; Tang et al., 2016). These

methods are either impractical for commercial-scale use for whole fish bait (N. Phelps, pers.

comm., 2020), or too toxic for general use (A. Pearson, BNZ, pers. comm., 2019). Chemical

processing methods are not viable risk management options and are not considered further.

Pre-export or post-arrival batch testing

OIE-approved batch testing methods (Corsin et al., 2009; OIE, 2019a) and standard

microbiological or molecular testing protocols are available to certify the absence of identified

risk organisms in the commodity. Pre-border testing procedures may be carried out by a testing

laboratory certified by the Competent Authority of the exporting country. Acceptance of a

negative batch test result by BNZ through the BNZ Country Approval Procedures may be a

viable risk management option (N. Ahmed, BNZ, pers. comm., 2020).

These risk management options are evaluated for each risk organism in the appropriate chapter.

6.2.3 Estimated levels of pathogen reduction

Qualitative descriptors are used to categorise estimated levels of reduction in pathogen load

considered likely to result from the application of each risk management option and assist in the

selection of risk management options for each risk organism. The indicative levels are:

Eliminate > 95% reduction in pathogen load

Substantial 71–95% reduction in pathogen load

Moderate 51–70% reduction in pathogen load

Slight < 50% reduction in pathogen load

Birnaviridae: Marine aquabirnavirus

7.1 Technical review

7.1.1 Aetiological agent

Marine aquabirnavirus (MABV) is a double-stranded RNA virus in the Family Birnaviridae,

which includes infectious pancreatic necrosis virus (IPNV) and related strains. Genomic analysis

indicates the strains of IPNV identified from salmonids are clustered into 6 genogroups (MABV

genogroups 1-6). A seventh genogroup includes 28 MABV strains associated with non-salmonid

fish. As IPNV is only associated with salmonids, other related MABV strains are considered as

IPNV-like MABV strains (Blake et al., 2001).

IPNV-like MABV strains associated with fish bait include yellowtail ascites virus (YAV) and

viral deformity virus (VDV) isolated from yellowtail (Seriola quinqueradiata) in Japan

(Sorimachi & Hara, 1985; Nakajima et al., 1998). These strains have low host specificity.

Strains isolated from different non-salmonid hosts and geographical areas are highly homologous

(more than 98.5% identical) (Hosono et al., 1996; Zhang & Suzuki, 2004; (Crane & Hyatt, 2011;

Shin et al. 2011; B. Jones, pers. comm., 2015). Each strain exerts different mortality rates in

different hosts depending on the water quality (temperature, salinity) and stocking levels (Barja,

2004; Crane & Hyatt, 2011).


7.1.2 OIE status

Infection with MABV is not listed by the OIE (OIE, 2019a).

7.1.3 New Zealand status

Infection with IPNV is regarded as an exotic, notifiable disease in New Zealand (Johnston, 2008;

Anon., 2016). Related MABV strains of genogroup 5 have been reported from farmed New

Zealand turbot (Colistium nudipinnis), flounder (Rhombosolea spp.) and from wild Chinook

salmon (Oncorhynchus tshawytscha) (Tisdall & Phipps, 1987; Anderson, 1996; Johnston, 2008).

Other non-salmonid MABV strains reported from fish bait species overseas (including VDV and

YAV) have not been reported from New Zealand (Anderson, 1990; Diggles et al., 2002; Tubbs

et al., 2007).

7.1.4 Zoonotic disease

MABV strains are not zoonotic (CABI, 2019a).

7.1.5 Epidemiology

Distribution and host range

Over 100 species representing more than 30 families of marine finfish are susceptible to IPNV

and IPNV-like MABV strains (Blackwell, 2019). IPNV strains also occur naturally in mussels

(Mytilus galloprovincialis, M. edulis) and oysters (Pinctada fucata), with moderate clinical signs

(Hill & Alderman, 1979) while scallops (Pecten spp.) can be experimentally infected. Shellfish

isolates cause clinical signs of IPNV in rainbow trout (Oncorhynchus mykiss) in experimental

infection (Hill, 1982; Mortensen et al., 1992; Isshiki et al., 2001; Molloy et al., 2013).

MABV strains have low host specificity. The species associated with MABV strains in fish bait

and their geographic/pathogen range are given in Table 7 (Nakajima et al., 1998; Kahn et al.,

1999; Crane et al., 2000; Isshiki et al., 2001; Crane & Williams, 2008; McColl et al. 2009a;

Crane & Hyatt, 2011; Munro & Midtlyng, 2011; Diggles, 2011;Van Beurden et al., 2012).

Prevalence ranges from 0.1% to 12.5% in saithe (Pollachius virens) and flounder (Platichthys

flesus) (Wallace et al., 2008; Diggles, 2011). IPNV prevalence was higher in wild non-salmonid

fish caught less than 5 km from infected marine salmon farms in Scottish waters, suggesting fish

farms may act as a reservoir for infection (Wallace et al., 2008, 2017). MABV strains including

IPNV were not reported in European anchovy (Engraulis encrasiciolus) or Atlantic mackerel

(Scomber scombrus), from a survey of 37 species of marine fish undertaken in coastal Scottish

waters (Wallace et al., 2008).


Table 7. Fish bait species susceptible to marine aquabirnavirus (MABV)

Family Host species Host geographical range1 Pathogen range2

Carangidae Japanese jack mackerel (Trachurus japonicus)

Northwest, Central Pacific West, Northwest Pacific (Asia, Japan, South Korea)

Clupeidae Atlantic herring (Clupea harengus), Bali sardinella (Sardinella lemuru), Indian oil sardine (Sardinella longiceps), Pacific herring (C. pallasii pallasii), round sardinella (Sardinella aurita), South American pilchard (Sardinops sagax), spotted sardinella (Ambylygaster sirm)

West Atlantic (USA to Argentina), Mediterranean, Western Pacific Japan, Indo-Pacific

East, Western Atlantic, Mediterranean, Western Pacific (Asia, Japan, South Korea)

Scombridae Atlantic mackerel (Scomber scombrus)

East Atlantic (Gulf of Cadiz) East Atlantic

Notes 1 Host geographical range (from FishBase, 2019) 2 Pathogen geographical range (from literature)

Potential host species in New Zealand

Potential host species of MABV include farmed Chinook salmon (O. tshawytscha) and

yellowtail kingfish (Seriola lalandi). MABV potentially affects invertebrates including mussels

(Mytilus spp.) and scallops (Pecten spp.) (Tubbs et al., 2007).

Pathogenicity

MABV strains are associated with economically serious diseases affecting wild and farmed fish.

Mortalities of up to 100% occur in salmonids (Oncorhynchus spp., Salmo spp. and Salvelinus

spp.) (Barja, 2004; Ruane et al., 1997; Skjesol et al., 2011).

Mortalities in non-salmonid fisheries are variable. Mortality ranges from 5% to 100% in farmed

eels (Anguilla anguilla) and from 80% to 90% in farmed Japanese amberjack (Seriola

quinqueradiata) (Nakajima et al., 1998; Van Beurden et al., 2012).

Clinical infection is rarely associated with wild fish, but subclinical infection may be endemic

(Van Beurden et al., 2012). Surviving fish are carriers of disease (Kahn et al., 1999).

Pathogenicity varies with season, size and age of fish, where older fish are less susceptible to

clinical infection (Isshiki et al., 2001).

The minimum infectious dose is low (> 1 TCID50 mL-1 (= 0.7 × 101 pfu mL-1) as determined by

bath immersion (4 h at 10 °C) (Smith et al., 2000; Isshiki et al., 2001; Urquhart et al., 2008;

Murray et al., 2005; Bebak & McAllister, 2009; Munro & Midtlyng, 2011; Oidtmann et al.,

2017).

Host specificity is low and cross-infection occurs between host species (Kahn et al., 1999;

Isshiki et al., 2001; Munro & Midtlyng, 2011; Van Beurden et al., 2012). Strains isolated from

Atlantic cod (Gadus morhua) caused 20% mortality in Atlantic salmon (S. salar) (Urquhart et

al., 2009), while strains from Japanese amberjack (Seriola quinqueradiata) were pathogenic to

spotted knifejaw (= gold stripe parrotfish) (Oplegnathus punctatus), barred knifejaw (=Japanese

parrotfish) (O. fasciatus) and yellowtail amberjack (S. lalandi) (= gold stripe amberjack, S.

aureovittata) (Isshiki et al., 2001).


Clinical signs

External clinical signs are variable. Subclinical carriers typically show no external signs.

Clinically infected fish may have a distended abdomen, moderate exophthalmia and

haemorrhages on the ventral skin surface and ventral fin base (Munro & Midtlyng, 2011).

Pathogen survival

MABV strains are unaffected by frozen storage (Kahn et al., 1999). They remain viable in

fomites (shipping containers, processing equipment and bait storage boxes) for at least 8 weeks

(Desautels & MacKelvie, 1975; Munro & Midtlyng, 2011). They remain viable for extended

periods in dead and decaying fish at high titre (> 106.0 pfu g-1) (plaque forming units) (Isshiki et

al., 2001; Munro & Midtlyng, 2011).

MABV strains remain viable in marine coastal waters for up to 35 days (at 10 °C to 15 °C)

(Toranzo et al., 1983; Kitamura & Suzuki, 2000; Kitamura et al., 2004; Oidtmann et al., 2017).

MABV strains also survive passage through piscivorous birds, where viral titre in faeces may

exceed the minimum infectious dose (McAllister & Owens, 1992). They can remain viable in

fresh waters for 17–43 days (at 10 °C to 15 °C) (Desautels & MacKelvie, 1975; Tu et al., 1975;

Ahne, 1982; Raynard et al., 2007; Oidtmann et al., 2017). However, data derived from

experiments may not reflect length of infectivity in natural waters (see section 6.1.2).

Transmission

Transmission is direct. This may be vertical (via infected eggs), or horizontal through

consumption of infected bait, cohabitation (through infected water, fish urine, faeces, and sexual

fluids) or through fomites (vehicles, nets, utensils associated with human activities such as

aquaculture and fish processing) (Munro & Midtlyng, 2011). Surviving fish are life-long latent

carriers and typically shed high levels of viral particles (Urquhart et al., 2008; Munro &

Midtlyng, 2011). Wild fish may function as a reservoir of infection for aquaculture species

(Castric, 1997).

MABV strains may be transported at least 30 km in coastal marine waters, even in slow currents

(2 cm sec-1), at titres exceeding the minimum infectious dose (Smith et al., 2000; Isshiki et al.,

2001; Urquhart et al., 2008; Murray et al., 2005; Bebak & McAllister, 2009; Munro & Midtlyng,

2011; Oidtmann et al., 2017).

MABV strains may be dispersed from fish bait discarded in landfill to waterways by piscivorous

birds (Munro & Midtlyng, 2011). They may also be dispersed between waterways (Desautels &

MacKelvie, 1975; Raynard et al., 2007). They are reported in rivers (at a titre of 1.3 × 104 pfu L-

1) up to 19 km from an infected freshwater site (Smith et al., 2000).

Infection and disease progression

Pathogen entry occurs by ingestion, usually through the intestinal epithelium or across the gill

membranes (Munro & Midtlyng, 2011). All life stages may be infected (DFO, 2017). Infection is

typically focused on tissues of the pancreas, kidney, spleen, pyloric caecae, intestine and liver,

but may also be present in the brain, gonads and gills (McColl et al., 2009; Munro & Midtlyng,

2011; Roberts, 2012).

Infected fish may shed high viral titres (from 101 to 107.5 pfu L-1) in body fluids, depending on

the virus strain, host species and age (Billi & Wolf, 1969; Johansen et al., 2011). However,

factors influencing morbidity appear complex and likely to also include environmental stressors


(e.g. temperature, salinity and oxygen concentration) (Munro & Midtlyng, 1999; Isshiki et al.,

2001).

Inactivation

MABV strains are unaffected by either short (30 days) or long-term (4 months) frozen storage (at

-20 °C) (Kahn et al., 1999). They are inactivated by UV light in seawater (Liltved et al., 2006)

and are likely to be denatured by ionising radiation at 50 kGy in frozen fish (DAFF, 2013).

7.2 Risk assessment


MABV strains are associated with fish bait species (Table 7), including herrings (Clupea sp.,

Sardinella spp. and Sardinops spp.) (Clupeidae) imported in large quantities during 2018 (Table

3). The prevalence of clinical disease is low in wild fish but may be higher adjacent to infected

marine farms (Wallace et al., 2008). Subclinically infected fish remain lifelong carriers and shed

high volumes of virus (Munro & Midtlyng, 2011). Pathogenicity varies with season, size and age

of the host (Isshiki et al., 2001; Munro & Midtlyng, 2011).

MABV strains are likely to remain viable in fish bait. As prevalence of subclinical disease in

wild fish is likely to be low, the likelihood of entry is assessed as low.

7.2.2 Exposure and establishment assessment

MABV strains are environmentally stable and can be spread by fomites, including commercial

processing, storage and transport equipment (Munro & Midtlyng, 2011). They are likely to

remain viable in commercial fish processing and bait storage facilities, in bait discarded to

landfill or through meltwater discharges (Munro & Midtlyng, 2011) for up to 24 weeks

(Desautels & MacKelvie, 1975). They may be dispersed from landfill to waterways and between

waterways by piscivorous birds (McAllister & Owens, 1992).

MABV strains remain viable in dead fish tissues, in moist packaging materials at high titre (>

106.0 pfu g-1) and in water for up to 8 weeks (Desautels & MacKelvie, 2011; Munro & Midtlyng,

2011). The infectious dose is low (Urquhart et al., 2008; Johansen et al., 2011) and all life stages

may be infected (DFO, 2017). Host specificity is low which could allow exposure to occur

through cross-contamination with discarded packaging in the supply chain.

The likelihood of exposure and establishment is assessed as medium.


Economic consequences

MABV strains affect over 30 families of marine fish (Blackwell 2019), including major farmed

and fished species in New Zealand. Wild stocks may act as reservoirs of infection (Raynard et

al., 2007; Wallace et al., 2008; Lafferty et al., 2015). Host specificity is low, while morbidity

and mortality may be high, reaching 100% in farmed fish. All life stages can be infected (DFO,

2017).

Infection with MABV is not an OIE-listed disease, so establishment would not result in

restrictions on trade access for New Zealand. It is likely that establishment would result in

lowered productivity and increased mortality, with serious economic consequences through lost

production in salmonid aquaculture, which was valued at $77 million in 2018 (Aquaculture New


Zealand, 2019). Establishment is also likely to adversely affect other aquaculture species,

including yellowtail kingfish (Seriola lalandi) (NIWA, 2017a) and snapper (Sparus aurata)

(Plant & Food, 2016). Establishment would also have indirect economic effects on businesses

reliant on aquaculture and commercial wild fisheries sectors (MPI, 2018a, 2018b, 2018c).

Social consequences

MABV strains are not zoonotic (CABI, 2019a). They are reported from a wide range of species

including several major inshore recreational target species such as snapper (Pagrus auratus) and

trevally (Pseudocarynx dentax) (Wynn-Jones et al., 2014, 2019). Introduction is likely to reduce

both the productivity and relative abundance of these already stressed inshore fish stocks (MPI,

2018a, 2018b, 2018c), with significant negative social impacts for customary and recreational

fishers.

Recreational fishing is the fifth most popular recreational activity for adults, involving 700,000

marine fishing trips annually, with a contribution of NZ$ 638 million to the GDP (Gross

domestic product) in 2015 (Southwick et al., 2018). The snapper and trevally recreational

fisheries contributed NZ$ 402 million and NZ$ 71 million respectively to the GDP in 2015

(Southwick et al., 2018).

Environmental consequences

MABV strains affect fish and invertebrates including mussels (Mytilus galloprovincialis) and

pearl oysters (Pinctada spp.). While scallops (Pecten spp.) can also be experimentally infected,

the environmental significance of infections in bivalves is unknown (Mortensen et al., 1992;

Isshiki et al., 2001; Molloy et al., 2013). Inshore fish and invertebrate species are considered

likely to encounter and consume discarded fish bait. Clupeids which feed on phytoplankton,

zooplankton and detritus, provide a significant food source for other fish, seabirds and marine

mammals and constitute a major link between trophic levels in marine food webs (Dunn et al.,

2012).

While host specificity and infectious dose are low, little is known of the effects of MABV strains

on wild fish stocks (Lafferty et al., 2015). MABV strains may be dispersed from marine to fresh

waters by piscivorous birds (Johansen et al., 2011) and may cause population level epizootics

that could result in significant environmental effects (Whittington et al., 2008; Paul et al., 2001;

Fish & Game, 2014).

The introduction of MABV strains would have moderately adverse economic, social and

environmental consequences. The consequences of establishment are assessed to be medium.

7.2.4 Risk estimation

The entry, exposure and consequence assessments are low to medium, so the risk estimate for

MABV strains is assessed to be medium. Under the procedures followed in this risk analysis,

risk management measures may be justified.

7.3 Risk management

MABV strains have been assessed to be a risk in fish bait. Infection is not an OIE-notifiable

disease, so the Aquatic Code (OIE 2019a) provides no specific guidance or processing

requirements that would ensure the destruction of the virus.


MABV strains are associated with fish bait (Table 7). Therefore, restriction to non-susceptible

species should substantially reduce the risk associated with MABV strains and be a viable risk

management option.

MABV strains have a wide geographical range (Table 7). Their occurrence in wild stocks in

other regions is likely to be low or negligible (Munro & Midtlyng, 2011). Restriction through the

BNZ Country Approval Procedures, to regions where these strains have not been reported should

substantially reduce the MABV risk and be a viable risk management option.

The presence of MABV strains in a shipment of fish bait may be verified by batch testing using

standard sampling procedures (Corsin et al., 2009; OIE, 2019a). Standard diagnostic procedures

for detection of these strains are provided in McColl et al. (2004b, 2009). Verification of a

negative test from a shipment should substantially reduce the risks associated with MABV

strains and be viable risk management options.

MABV strains are unaffected by freezing, so frozen storage (at -18°C for either 168 hours, or for

4 months), is not a risk management option.

MABV strains are denatured by ionising radiation at 50 kGy (DAFF, 2013). This irradiation

treatment should substantially reduce the MABV risk and be a viable risk management option.

7.3.1 Risk management options

In addition to the general risk management measures that are proposed for all imported fish bait,

one or more of the following risk management options would reduce the MABV specific risk to

an acceptable level:

Option 1

• Competent Authority attestation that the commodity does not include: Japanese jack

mackerel (Trachurus japonicus) (Carangidae); Atlantic herring (Clupea harengus), Bali

sardinella (Sardinella lemuru), Indian oil sardine (Sardinella longiceps), Pacific herring

(Clupea pallasii pallasii), round sardinella (Sardinella aurita), South American pilchard

(Sardinops sagax), spotted sardinella (Ambylygaster sirm) (Clupeidae); and Atlantic

mackerel (Scomber scombrus) (Scombridae) should substantially reduce the risks

associated with MABV strains, and the commodity could be imported without any further

restrictions.

Option 2

• Acceptance of region/country freedom through the BNZ Country Approval Procedures

should substantially reduce the MABV risk so the commodity could be imported without

any further restrictions.

Option 3

• A negative test result from a shipment of the commodity, following approved sampling and

diagnostic procedures through the BNZ Country Approvals Procedures should substantially

reduce the MABV risk, so the commodity could be imported without further restrictions.

Option 4

• Treatment with ionising radiation (at 50 kGy) should substantially reduce the MABV risk, so

the commodity could be imported without further restrictions.


Iridoviridae: Megalocytivirus (red sea bream iridovirus - RSIV) and associated viruses



The Genus Megalocytivirus is one of five genera classified within the family Iridoviridae (OIE,

2019a). Four main species of Megalocytivirus are recognised:

• Infectious spleen and kidney necrosis virus (ISKNV) (two genotypes);

• Red sea bream iridovirus (RSIV) (two genotypes);

• Turbot reddish body iridovirus (TRBIV);

• Three-spine stickleback iridovirus (TSIV).

These represent a species complex consisting of at least 3 genotypes, with closely related strains

that infect a wide range of marine and freshwater fish species (Nolan et al., 2015; Blackwell,

2019). While other uncharacterised strains, including scale drop disease virus of Asian seabass

(Lates calcarifer) (SDDV) are reported (Halaly et al., 2019), natural viral recombinations within

this species complex also occur (Kurita & Nakajima, 2012; Sriwanayos et al., 2013; Nolan et al.,

2015; Blackwell, 2019). For example, the RSIV-Ku strain isolated from red sea bream (Pagrus

major) from aquaculture in Chinese Taipei is a natural recombination of ISKNV and RSIV

genotypes (Shui et al., 2018).

8.1.2 OIE status

Infection with red sea bream iridovirus (RSIV) and infectious spleen and kidney iridovirus

(ISKNV) are recognised as agents of the OIE-listed red sea bream iridovirus disease (RSIVD)

(OIE, 2019a). Other strains within the species complex are not agents of OIE-listed diseases.


RSIV and ISKNV have not been reported from New Zealand from active monitoring

programmes for RSIVD. New Zealand is considered free from RSIVD (OIE 2019a) and this is a

notifiable disease in New Zealand (Anon, 2016). RSIV and ISKNV are considered exotic to New

Zealand.


RSIV and ISKNV strains are not considered zoonotic (OIE, 2019a).

8.1.5 Epidemiology


RSIV and ISKNV strains have a very wide host range from 20 families of marine and freshwater

fish (Gibson-Kueh et al., 2004; Song et al., 2008; Xu et al., 2010; Blackwell, 2019; OIE 2019a,

2019b).

The species associated with RSIV and ISKNV in fish bait and their geographic/pathogen range

are given in Table 8. Prevalence in wild fish stocks is poorly known. (Gibson-Kueh et al., 2004;

Song et al., 2008; OIE, 2019b).


Table 8. Fish bait species susceptible to red sea bream iridovirus (RSIV), infectious spleen and kidney necrosis virus (ISKNV) and associated iridoviruses

Family Host Species Host geographical range1 Pathogen range2


Northwest Pacific, Japan, Korea, Southeast Asia

Northwest, Central Pacific

Clupeidae Pacific herring (Clupea pallasi pallasii)

Arctic, Northwest, Northeast Central Pacific

Northeast, Central Pacific


Cosmopolitan in tropical, subtropical and temperate waters


Scombridae Chub mackerel (Scomber japonicus)

Indo-Pacific, Eastern, Western, Central Pacific Southeast Atlantic (South Africa)


Notes 1 Host geographical range (from FishBase, 2019) 2 Pathogen geographical range (from literature). The Northwest and Central Pacific includes Australia, China, Chinese Taipei,

Hong Kong, South Korea, Malaysia, the Philippines, Singapore and Thailand.


A wide range of marine and freshwater fish are considered susceptible to RSIV and ISKNV

strains. These include snapper (Pagrus auratus) (Sparidae), silver trevally (Pseudocaranx

dentex), trevally (Caranx georgianus) and jack mackerel (Trachurus spp.) (Carangidae). ISKNV

also occurs in freshwaters, where susceptible species include galaxiids (Galaxias spp.) and

cyprinids such as grass carp (Ctenopharyngodon idella) (Diggles, 2003; Blackwell, 2019).

Pathogenicity

All host life stages are susceptible, but juveniles are more susceptible than adults (OIE, 2019a).

Pathogenicity varies among and between viral clades and strains. While pathogenicity ranging

from 50% to 100% occurs in farmed red sea bream (S. auratus) (Sparidae) (Sano et al., 2011),

strains affecting wild fish species are of lower pathogenicity (OIE, 2019a). Pathogenicity is

temperature related, with higher mortalities generally associated with temperatures over 25 °C

(Jun et al., 2009).

The infectious dose (determined experimentally by intra-peritoneal (i.p.) injection of S. auratus

as 103 TCID50 -1) resulted in mortalities from 20% to 60% (Shinmoto et al., 2009; Kurita &

Nakajima, 2012). The lethal dose (105 TCID50 -1) caused 100% mortality, 7–10 days post

infection. Infection was associated with liver, kidney, spleen, heart and muscle tissues (Shinmoto

et al., 2009). Experimental infection (by bath inoculation) indicated subclinical infection was

widespread in surviving carrier fish that remained carriers of disease (Ito et al., 2013). RSIV and

ISKNV strains are not zoonotic.

Clinical signs

Clinical signs of infection vary widely, and infection may progress with no external signs (Noga,

2010; Kurita & Nakajima, 2012; OIE, 2019a).

Pathogen survival

Experimental data indicate megalocytivirus strains survive in water for at least 7 days at 15 °C

(Ito et al., 2013) and for longer periods at 4 °C. They are unaffected by frozen storage in fish

tissues (to -80 °C) and remain viable after repeated freeze-thaw cycles (OIE, 2019a, 2019b).


While viability in natural waters is unknown (see section 6.1.2), RSIV strains remained viable in

frozen baitfish used as feed in Asian aquaculture (Laijimin et al., 2015).

Transmission

Disease transmission is horizontal, through the water column (OIE, 2019a).


Infection occurs by direct contact with viral particles, or through consumption of infected fish

(Sano et al., 2011; OIE, 2019a). Disease progression is initially focused on the viscera (including

the heart). Infection then typically spreads to the gills, skeletal muscle, connective tissue and

spleen (Whittington et al., 2010; OIE, 2019a, Ito et al., 2013).

Infection with ISKNV strains may also be subclinical (Hine & Diggles, 2005), where overt

disease may occur when the host is stressed due to adverse environmental conditions such as

temperature or salinity (Mohr et al., 2015)

Inactivation

Megalocytiviruses are unaffected by short (16 hours) or medium-term frozen storage (to at least

4 months at -20 °C) (OIE, 2019a). They are likely to be inactivated by ionising radiation of

frozen fish bait (at 50 kGy) (DAFF, 2013).

8.2 Risk assessment


RSIV and ISKNV strains are associated with fish bait species (Table 8), including Pacific

herring (Clupea pallasii pallasii) (Clupeidae) imported from the Asia-Pacific region in high

volumes during 2018 (Table 3). Prevalence in wild fish is unknown, but assumed to be low-

moderate, as these viruses remain viable in frozen fish bait (Laijimin et al., 2015). Surviving fish

may be carriers with no external signs of infection (Ito et al., 2013).

As the volume of susceptible fish bait derived from the Asia-Pacific region is high, the risk of

entry of RSIV and ISKNV strains is assessed as low-medium.


RSIV and ISKNV strains are unaffected by short or medium-term frozen storage and multiple

freeze-thaw cycles. They remain viable in seawater for up to 7 days at 15 °C (Ito et al. 2013) and

for longer periods at 4 °C (OIE 2019a). All life stages of the host are susceptible (OIE 2019a).

The infectious dose (determined experimentally) in red sea bream (S. auratus) is low (103

TCID50 -1), with mortalities varying from 20% to 60% (Kurita & Nakajima, 2012; Shinmoto et

al., 2009).

Pathogenicity in wild stocks is unknown, but these pathogens may be spread through fish bait,

where used as aquaculture feed (Sano et al., 2011; Laijimin et al., 2015; OIE, 2019a). A wide

variety of potential host species are present in New Zealand waters.

RSIV and ISKNV strains are environmentally persistent. The infectious dose is assumed to be

low and infection can be initiated through fish bait. The likelihood of exposure and establishment

is therefore assessed to be medium.




Infection with RSIV, ISKNV are OIE-listed diseases (OIE, 2019a), so their establishment would

result in loss of trade access for New Zealand. They have moderate-high morbidity and mortality

(Nakajima et al., 1998; Van Beurden et al., 2012). Their establishment could result in lower

productivity and increased mortality in over 20 families of marine and fresh water fish

(Blackwell, 2019), including snapper (Pagrus auratus) (exports valued at $33 million) and jack

mackerel (Trachurus spp.) (exports valued at $68 million) in 2018 (Aquaculture New Zealand,

2019).

Productivity losses and increased mortality would also negatively impact the developing

aquaculture of yellowtail kingfish (Seriola lalandi) and grouper (Polyprion oxygeneios)

(Diggles, 2003; Symonds et al., 2014).

Establishment of freshwater strains may also adversely affect grass carp (Ctenopharyngodon

idella) and silver carp (Hypophthalmichthys molitrix) aquaculture, used for weed control

(Clayton & Wells, 1999; Blackwell, 2019).

Social consequences

RSIV and ISKNV strains are not considered zoonotic. They have a wide host range (Hine &

Diggles, 2005; Blackwell, 2019), so establishment may negatively affect target species of

importance for customary and recreational fishing (Wynn-Jones et al., 2014, 2019). Recreational

fishing is the fifth most popular recreational activity for adults, involving 700,000 marine fishing

trips annually, with a contribution of NZ$ 638 million to the GDP (Gross domestic product) in

2015 (Southwick et al., 2018).

The snapper recreational target fishery contributed NZ$ 492 million to the GDP (Gross domestic

product) in 2015 (Southwick et al., 2018) and is a taonga for Maori culture (MPI, 2018c). In

fresh water, ISKNV infects galaxiids (Galaxias spp.), which forms a major part of customary

and recreational fisheries for whitebait (NIWA, 2019).

Many of these marine and freshwater fish stocks are considered stressed (MPI, 2018a, 2018b,

2018c). A reduction in productivity and increase in mortality due to the establishment of RSIV

and ISKNV strains would have significant social impacts for New Zealand.


RSIV and ISKNV strains are reported from over 20 fish families and 60 species of fish in marine

and fresh waters (Xu et al., 2010; OIE, 2019a, 2019b; Blackwell,2019). Inshore species

including clupeid, mugilid and scombroid fish are considered likely to encounter and consume

discarded fish bait. Clupeids feed on phytoplankton, zooplankton and detritus and represent a

major link between trophic levels in food webs (Varpe et al., 2005; DFO, 2009; Dunn et al.,

2012). Clupeid and mugilid species provide a significant food source for other fish, seabirds and

marine mammals (Dunn et al., 2012).

Reductions in Pacific herring (C. p. pallasii) abundance (from over-fishing) caused a reported

60% decrease in the relative abundance of wild salmon (Oncorhynchus tshawytscha) in coastal

British Columbia, with major biological, economic and social consequences for humans and wildlife (Pagowski et al., 2019). The wide host range of these pathogens may potentially cause

population level epizootics in wild fisheries with significant environmental effects on the New


Zealand coastal environment (Whittington et al., 2008; Paul et al., 2001; Fish & Game, 2014;

Lafferty et al., 2015).

ISKNV strains may be dispersed into fresh waters by infected euryhaline fish species (Go &

Whittington, 2006; Go, 2015), or through the use and discarding of bait in fresh water

(Blackwell, 2013). If ISKNV strains established in wild cyprinids (such as goldfish, Carassius

auratus), these could act as reservoir hosts for infection of native galaxiids (Galaxias spp.)

(Galaxiidae) (Clayton & Wells, 1999; Blackwell, 2019), as well as threatened aquatic

amphibians (Hine & Diggles, 2005).

Infection may also become endemic, with overt infection occurring when the host is

environmentally stressed (Mohr et al., 2015). Hine & Diggles (2005) assessed the environmental

consequences of establishment of RSIV and ISKNV strains (through introduced ornamental fish)

as high to catastrophic.

The introduction of RSIV and ISKNV strains would have significant negative economic, social

and environmental consequences. The consequences of establishment are assessed to be high.


The entry, exposure, establishment, and consequence assessments for RSIV and ISKNV strains

are medium to high, so the risk estimation is assessed to be medium-high. Under the procedures

followed in this risk analysis, risk management measures may be justified.

8.3 Risk management

RSIV and ISKNV strains have been assessed to be a risk in fish bait. Infection with these strains

is OIE-listed (OIE, 2019a), so the OIE Code provides some guidance on control measures

necessary for inactivation. However, the OIE-listed options require heat treatment which would

render the fish bait unfit for purpose. These are not considered further.

RSIV and ISKNV strains are associated with some fish bait species (Table 8), while other fish

bait species are not associated with RSIV and ISKNV. Therefore, restriction of the commodity to

species not associated with these strains should substantially reduce the risk and be a viable risk

management option.

RSIV and ISKNV strains have a wide geographical range (Table 8). Their occurrence in wild

stocks in other regions is likely to be low or negligible (OIE, 2019a). Restriction, through the

BNZ Country Approval Procedures, to regions where these strains have not been reported should

substantially reduce the risk and be a viable risk management option.

The presence of RSIV and ISKNV strains in a shipment of fish bait may be verified by batch

testing using standard sampling procedures (Corsin et al., 2009; OIE, 2019a). Standard

diagnostic procedures for detection of these strains are provided in McColl et al. (2004b, 2009).

Verification of a negative test for RSIV and ISKNV from a shipment should substantially reduce

the risk and be a viable risk management option.

RSIV and ISKNV are unaffected by frozen storage (OIE, 2019b). Therefore, frozen storage

(at -18°C) for either 168 hours or 4 months is not a risk management option.

RSIV and ISKNV strains are denatured by exposure to ionising radiation (to 50 kGy) (DAFF,

2013). This irradiation treatment should substantially reduce the risk and be a viable risk

management option.



In addition to the general risk management measures that are proposed for all imported fish bait

(section 6.2.1), one, or more of the following risk management options would reduce the RSIV

and ISKNV specific risk to an acceptable level:

Option 1

• Competent Authority attestation that the commodity does not include: Japanese mackerel

(Trachurus japonicus) (Carangidae); Pacific herring (Clupea pallasii pallasii) (Clupeidae);

flathead grey mullet (Mugil cephalus) (Muglidae); and chub mackerel (Scomber japonicus)

(Scombridae) should substantially reduce the RSIV and ISKNV risk, so the commodity

could be imported without any further restrictions.

Option 2

• Acceptance of region/country freedom through the BNZ Country Approval Procedures,

should substantially reduce the RSIV and ISKNV risk, so the commodity could be

imported without any further restrictions.

Option 3

• A negative test result from a shipment of the commodity, following sampling and diagnostic

procedures approved through the BNZ Country Approvals Procedures should substantially

reduce the RSIV and ISKNV risk, so the commodity could be imported without further

restrictions.

Option 4

• Treatment with ionising radiation (to 50 kGy) should substantially reduce the RSIV and

ISKNV risk, so the commodity could be imported without any further restrictions.

Iridoviridae: Erythrocytic necrosis virus (ENV)


9.1.1 Aetiology

Erythrocytic necrosis virus (ENV) is the agent of viral erythrocytic necrosis disease (VEN). It is

a double-stranded DNA virus, classified within family Iridoviridae, originally isolated from

salmonids and herring (Clupea spp.) (Walker & Sherburne, 1977; Winton & Hershberger, 2014).

It may represent a new species within the Iridoviridae (Emmenegger et al., 2014; Hick et al.,

2016; Purcell et al., 2016).

Several ENV strains exist among host species and within geographical ranges, but their genetic

relationships are unknown (Winton & Hershberger, 2014; Hick et al., 2016). Viral isolates from

Chinook salmon (O. tshawytscha) and Pacific herring (Clupea pallasii pallasii) were over 99%

similar, suggesting that cross-species infection occurs (Hershberger et al., 2006, 2009; Winton &

Hershberger, 2014; Pagowski et al., 2019).

9.1.2 OIE status

Infection with VEN is not an OIE-listed disease (OIE, 2019a).



ENV has not been reported from New Zealand (Anderson, 1990; Diggles, 2011). Infection with

ENV is not a notifiable disease in New Zealand (Anon., 2016).


ENV is not zoonotic (Blackwell, 2019).

9.1.5 Epidemiology


ENV is reported from over 10 families and 20 species of elasmobranch and teleost marine fish

(MacMillan et al., 1980; Dannevig & Thorud, 1999; Davies et al., 2009; Emmenegger et al.,

2014; Winton & Hershberger, 2014; Hick et al., 2016; Blackwell, 2019). It is reported from wild

and farmed fish (Hick et al., 2016). The species associated with ENV in fish bait and their

geographic/pathogen range are given in Table 9 (Nicholson & Reno, 1981; Hershberger et al.,

2011; Emmenegger et al., 2014).

ENV occurs in wild Pacific herring (C. p. pallasii) at a prevalence of 27%, reaching 67% in

epizootics (Reno et al., 1985). Prevalence in Californian anchovy (E. mordax) is 38%, reaching

50% in epizootics (Hershberger et al., 2009, 2011; Pagowski et al., 2019).

Table 9. Fish bait species susceptible to erythrocytic necrosis virus (ENV)



Baltic Sea, Northeast, Northwest Atlantic

Baltic Sea. Northeast Atlantic, (Europe, Spain, United Kingdom)

Clupeidae Pacific herring (C. pallasii pallasii)

Arctic, Northwest, Northeast Pacific (Alaska to Mexico)

Northwest, Northeast Pacific (Japan)

Engraulidae Californian anchovy (Engraulis mordax)

Northeast Pacific (Canada to Mexico)

Northeast Pacific (USA, Canada)

Notes 1 Host geographical range (from FishBase, 2019) 2 Pathogen geographical range (from literature). ENV is reported from the North Atlantic and North Pacific Oceans, as well as

from Spain, Japan and the United Kingdom. It is not reported from the Southern Hemisphere (Hick et al., 2016).


ENV may potentially affect a range of inshore fish in New Zealand, including dogfish

(Triakidae) (Tubbs et al., 2007), as well as farmed and wild salmonids.

Pathogenicity

ENV is unculturable, so relationships between virus presence and clinical disease are difficult to

determine experimentally (Emmenegger et al., 2014). It may remain dormant where water

temperatures are lower than 20 °C (Pagowski et al., 2019). Pathogenicity varies among ENV

strains, host species, route of infection and intensity of exposure (Reno et al., 1985; Winton &

Hershberger, 2014). All life stages of the host may be infected, and prevalence may be inversely

correlated to host age (Smail, 1982).

Transmission and infection details are uncertain (Winton & Hershberger, 2014; Pagowski et al.,

2019). ENV inclusion bodies are present in erythrocytes collected from kidney and spleen tissues


(Glenn et al., 2012) and infection can be experimentally induced by filtered preparations from

diseased fish (Evelyn & Traxler, 1978).

Clinical signs

Infection usually occurs without external clinical signs, but fish with acute infection (VEN) may

show signs of anaemia (Reno et al., 1985; Winton & Hershberger, 2014).

Pathogen survival

Iridoviruses are unaffected by frozen storage (OIE, 2019b). From experimental data, ENV

remains viable in erythrocytes in solution up to 2 weeks (Nicholson & Reno, 1981). However,

viability may be less in natural waters (see section 6.1.2).

Transmission

Disease transmission is mainly horizontal, but vertical transmission is also reported (Rohovec &

Amandi, 1981).


ENV affects red blood cells, causing long-term systemic anaemia. This may eventually affect up

to 90% of erythrocytes in the host (Reno et al., 1985). Acute disease (VEN) is often associated

with co-infection by opportunistic pathogens (Hershberger et al., 2006; Glen et al., 2012).

Infection occurs through the water column, where the initial prevalence is likely to be low in

wild fish stocks (Hershberger et al., 2006). From a multi-year field study on wild Pacific herring

(C. pallasii pallasii), population-level VEN epizootics were an important component of natural

mortality (Hershberger et al., 2009; Pagowski et al., 2019).

Subclinical infection of iridoviruses rarely causes high morbidity or mortality in wild fish (Reno

et al. 1985). However, co-infection may occur with other viruses (Hershberger et al., 2006) and

overt disease may then develop where the host is stressed by environmental stressors, such as

low dissolved oxygen and low salinity (Meyers et al., 1986; Winton & Hershberger, 2014; Mohr

et al., 2015).

Inactivation

Iridoviruses including ENV are unaffected by either short (168 hours) or medium-term (4

months) frozen storage (at -20 °C) (OIE, 2019b). ENV is likely to be denatured by exposure to

ionising radiation (to 50 kGy) (DAFF, 2013).

9.2 Risk assessment


ENV is reported from listed fish bait species from a wide geographical area (Table 9). Viral

prevalence ranges from 27% to 38% in wild bait fish, reaching 67% during epizootics

(Hershberger et al., 2009; Pagowski et al., 2019). As viral shedding rates increase when herrings

are stressed, viral particles may also be concentrated in the water used to freeze baitfish (Hick et

al., 2016). ENV is likely to remain viable in frozen fish bait (OIE 2019b).

ENV is associated with fish bait across a wide geographical area, at a moderate-high prevalence

and may also be present in meltwater. The likelihood of entry is therefore assessed to be high.



The transmission mechanisms of ENV are poorly known. Experimental data indicate iridoviruses

remain viable in seawater for up to 7 days at 15 °C (Ito et al., 2013) and for longer periods at

4 °C (Anon., 2018). However, the length of time that ENV may remain infective under natural

conditions is unknown (see section 6.1.2).

All life stages of the host may be infected (Smail, 1982). Infection of farmed salmonids,

including Chinook salmon (Oncorhynchus tshawytscha), through wild fish bait species is

reported, suggesting the infectious dose of ENV is likely to be low (Hershberger et al., 2006,

2009; Sano et al., 2011; Winton & Hershberger, 2014; OIE, 2019a; Pagowski et al., 2019).

A wide variety of potential hosts are present in New Zealand waters. ENV remains viable in

seawater and is environmentally resistant. Infection can be initiated in aquaculture species when

fish bait is used as feed. Therefore, the likelihood of exposure and establishment is assessed as

medium.



Infection with VEN is not an OIE-listed disease, so establishment would not affect New

Zealand’s trade status. Subclinical infection commonly occurs in wild fish stocks, but rarely

causes high morbidity or mortality (Reno et al., 1985). However, ENV is the agent of viral

erythrocytic necrosis (VEN) in salmonids farmed in marine waters. ENV may be associated with

high mortality and morbidity (Hershberger et al., 2009; Winton & Hershberger, 2014; Pagowski

et al., 2019). It would likely have a significant effect on New Zealand marine aquaculture, which

is focussed on Chinook salmon (O. tshawytscha), valued at $77 million in export earnings in

2018 (Hick et al., 2016; Seafood New Zealand, 2019).

ENV has a wide host range, affecting over 10 families of elasmobranch and teleost marine fish

(Blackwell, 2019). Establishment would also have a major economic effect on the developing

aquaculture of yellowtail kingfish (Seriola lalandi) (Carangidae) (Diggles, 2003; Symonds et

al.,2014).

The effects of introduction of viral pathogens on wild fish stocks are unknown (Lafferty et al.,

2015), but are likely to be low (Reno et al., 1985; Kahn et al., 1999). ENV may affect

commercial inshore fisheries including jack mackerel (Trachurus spp.), valued at $68 million in

2018 (Seafood New Zealand, 2019).

Social consequences

ENV is not considered zoonotic. Establishment would have significant social implications for the

recreational hatchery-based fisheries for rainbow trout (O. mykiss) and brown trout (Salmo

trutta) (Marsh & Mkwra, 2013; Fish & Game, 2014), due to lost productivity and increased

mortality. The Lake Taupo trout fishery alone was valued in 2012 at NZ$ 70–80 million (Marsh

& Mkwra, 2013).

Recreational marine fishing is the fifth most popular recreational activity for adults, involving

700,000 marine fishing trips in 2015, with a contribution of NZ$ 638 million to the GDP (Gross

domestic product). Recreational yellowtail kingfish fishing contributed NZ$ 79.3 million to the

New Zealand GDP in 2015 (Southwick et al., 2018).



ENV is reported from over 20 species of marine and anadromous fish (Hine & Diggles, 2005;

Blackwell, 2019). Little is known about the potential effects of virus infection of wild fish stocks

(Lafferty et al., 2015), but subclinical infection is rarely associated with high morbidity and

mortality (Reno et al., 1985; Kahn et al., 1999). However, inshore fish species including clupeid

and engraulid fish are considered likely to encounter and consume discarded fish bait. Clinical

infection in North American clupeid and engraulid fisheries has been associated with population

level epizootics (Hick et al., 2016). Reductions in herring abundance (from over-fishing) caused

a reported 60% decrease in the relative abundance of wild salmon (O. tshawytscha) in coastal

British Columbia. This resulted in major biological, economic and social consequences for

humans and wildlife (Pagowski et al., 2019). Pacific herring (C. p. pallasi) is a keystone species

in marine food webs (Varpe et al., 2005; DFO, 2009), while clupeid and engraulid species

represent significant feed for other marine organisms, including fish, seabirds and marine

mammals (Dunn et al., 2012). The introduction of agents capable of causing population level

epizootics would have significant environmental effects on New Zealand coastal fisheries

(Whittington et al., 2008; Paul et al., 2001).

The introduction of ENV would have significant negative economic, social and environmental

consequences. The consequences of establishment are assessed to be high.


The entry, exposure, establishment, and consequence assessments are medium-high, so the risk

estimation for ENV is medium-high. Under the procedures followed in this risk analysis, risk

management measures may be justified.

9.3 Risk management

ENV has been assessed to be a risk in fish bait. Infection with VEN is a not an OIE-notifiable

disease, so the OIE Aquatic Code (OIE, 2019a) provides no specific guidance or processing

requirements that would ensure the destruction of the virus.

ENV is associated with fish bait (Table 9). Therefore, restriction to non-susceptible species in

the commodity should substantially reduce the ENV risk and be a viable risk management

option. ENV has a wide geographic range (Table 9). The occurrence is likely to be low in other

geographic regions. Restriction through the BNZ Country Approval Procedures to regions where

ENV has not been reported should substantially reduce the ENV risk and be a viable risk

management option.

The presence of ENV strains in a shipment of fish bait may be verified by batch testing using

standard sampling procedures (Corsin et al., 2009; OIE, 2019a). Standard diagnostic procedures

for detection of the virus are provided in Emmenegger et al. (2014) and Pagowski et al. (2019).

Verification of a negative test from a shipment should substantially reduce the ENV risk and be a

viable risk management option.

ENV is unaffected by freezing (OIE, 2019b) so frozen storage (at -18°C for either 168 hours, or

for 4 months) is not a viable risk management option. ENV is denatured by ionising radiation at

50 kGy (DAFF 2013). This irradiation treatment should substantially reduce the ENV risk and be

a viable risk management option.




(section 6.2.1), one or more of the following risk management options would reduce the ENV

specific risk to an acceptable level:

Option 1

• Competent Authority attestation that the commodity does not include European herring

(Clupea harengus), Pacific herring (Clupea pallasii pallasii) (Clupeidae) and European

anchovy (Engraulis mordax) (Engraulidae), should substantially reduce the ENV risk, so

the commodity could be imported without any further restrictions.

Option 2

• Acceptance of country/zone freedom through the BNZ Country Approval Procedures

should substantially reduce the ENV risk so the commodity could be imported without any

further restrictions.

Option 3

• A negative test result from a shipment of the commodity, following sampling and diagnostic

procedures through the BNZ Country Approvals Procedures should substantially reduce the

ENV risk, so the commodity could be imported without further restrictions.

Option 4

• Treatment with ionising radiation (to 50 kGy) should substantially reduce the ENV risk, so


Nodaviridae: Nervous necrosis virus (NNV)


10.1.1 Aetiology

Nodaviruses are non-enveloped positive-stranded RNA viruses, classified as a Betanodavirus.

The type specimen is nervous necrosis virus (NNV). Four main genotypes are recognised by the

OIE (OIE, 2019b): SJNNV-type (striped jack nervous necrosis virus), TPNNV-type (tiger puffer

nervous necrosis virus), BFNNV-type (barfin flounder nervous necrosis virus) and RGNNV-type

(red-spotted grouper nervous necrosis virus) (OIE 2019b). These genotypes broadly correlate

with serotype A (SJNNV), serotype B (TPNNV) and serotype C (BFNNV and RGNNV) (Mori

et al., 2003; OIE, 2019b).

Several other unclassified serotypes have been proposed (Castric, 1997; Johansen et al., 2004;

Kim et al., 2011; Doan et al., 2017; Prado-Alvarez & Garcia-Fernandex, 2019; Bandin & Souto,

2020), but these are not currently recognised by the OIE (OIE, 2019a).

Serotypes are grouped by their optimum growth temperature in vitro (Iwamoto et al., 2000).

Serotype A is infective over a temperature range from 20 °C to 30 °C. Serotype B only affects

tiger puffer (Takifugu rubripes) and is optimal at 20 °C. Serotype C affects fish over a wide

temperature range, including cold water (from 15 °C to 20 °C) and warm water fish (from 25 °C

to 30 °C) (OIE, 2019b). Within each serotype, host specificity is low, as serotypes of one


genotype cause disease in host species that are predominantly infected by other genotypes

(Munday et al., 2002; Tanaka et al., 2003; Sano et al., 2011).

10.1.2 OIE status

Infection with NNV or other nodaviruses is described by the OIE as viral encephalopathy and

retinopathy (VER) (OIE, 2019b). It is not an OIE-listed disease (OIE, 2019a).


No genotype of NNV have been reported from New Zealand (Hine & Diggles, 2005; Tubbs et

al., 2007; Moody & Horwood, 2008). Infection with NNV is not a notifiable disease in New

Zealand (Anon. 2016) but it is considered exotic (Tubbs et al., 2007).


Betanodaviruses including NNV are not considered zoonotic (Adachi et al., 2008; OIE, 2019a).

10.1.5 Epidemiology

NNV causes viral nervous necrosis disease (VNN) or viral encephalopathy and retinopathy

(VER) (OIE, 2019b).


NNV strains are widely distributed (Table 10). They occur in 24 families of fish and coleoid

molluscs in marine and fresh waters. They are also reported from bivalve molluscs (blue mussel,

Mytilus galloprovincialis), crustaceans (blue swimming crab, Portunus pelagicus) and annelid

worms (Nereis spp.) adjacent to infected fish farms (Fichi et al., 2015; Fiorito et al., 2015; Costa

& Thompson, 2016; Doan et al., 2017; Volpe et al., 2018; OIE, 2019b; Blackwell, 2019). The

international trade in nereid worms for fish food may represent an infection pathway (OIE

2019b). NNV strains may also be dispersed by piscivorous birds (OIE 2019b).

NNV occurs in wild Mediterranean fish species at a prevalence of 21%, where the absence of

infected clupeids may reflect their higher susceptibility and mortality (Berzak et al., 2019). NNV

occurs at a prevalence up to 100% in wild mackerel (Scomber spp.) in the Philippines (de la Pena

et al., 2011). NNV strains are a major pathogen of larval and juvenile fish in Chinese, Asian,

Australian and European aquaculture (Castric, 1997; Chi et al., 2016).

The species associated with NNV strains in fish bait and their geographic/pathogen range are

given in Table 10 (Mori et al., 1991, 1992; Nguyen et al., 1996; Ucko et al., 2004; Gomez et al.,

2006; Sakamoto et al., 2008; Tubbs et al., 2007; Gomez et al., 2010; Crane & Hyatt, 2011;

Forrest et al., 2011; Panzarin et al., 2012; Fiorito et al., 2015; Zorriehzahra et al., 2016; OIE,

2019b; Bandin & Sousa, 2020).


Potential species at risk in New Zealand include eels (Anguilla spp.), flatfish (Pleuronectidae),

jack mackerel (Trachurus spp.), snapper (Pagrus auratus), striped trumpeter (Latris lineata),

trevally (Caranx georgianus) and yellowtail kingfish (Seriola lalandi) (Blackwell, 2019).

Pathogenicity

NNV causes economically significant neurological disease (VNN) in a wide range of wild and

farmed fish species (Munday et al., 2002; Sano et al., 2011). Mortalities may reach 100% in


aquaculture, particularly for larval and juvenile fish (OIE, 2019b). Infection in older, larger fish

is often subclinical, where surviving fish may represent reservoirs of disease (Sano et al., 2011;

Chi et al., 2016).

Mortality of farmed sea bream (S. auratus) infected with NNV may reach 50% in marine

aquaculture, with no clinical signs (Cherif et al., 2008).

Experimental infection (by injection) of sevenband grouper (Epinephelus septemfasciatus) at a

rate of 105.5 TCID50 fish-1 resulted in cumulative mortalities up to 44%. Viral titres between 109.5

and 1010.0 TCID50 g-1 were recovered from brain tissues. Viral titre in the tank seawater reached

105 TCID50 L-1. All surviving fish had viral titre in brain tissues of 104 to 107 TCID50 g

-1 (Nishi et

al. 2016). While these data indicate high mortality and high viral shedding rates in surviving

fish, little is known about infectivity in wild fish populations (Lafferty et al., 2015).


Table 10. Fish and coleoid cephalopod mollusc species susceptible to nervous necrosis virus (NNV)


Fish families

Carangidae Atlantic horse mackerel (Trachurus trachurus), Chilean jack mackerel (T. murphi)

Mediterranean, Eastern, Southwest Atlantic (Argentina), Pacific

Mediterranean, Indo-Pacific, Pacific, Northeast, Southwest Atlantic

Japanese jack mackerel (T. japonicus), Pacific jack mackerel (T. symmetricus)

Eastern, Central, South Pacific, Northwest Pacific, Japan, Southeast Asia

Indo-Pacific, North Pacific


Northeast, Northwest Atlantic, North Sea, Baltic Sea, Black Sea

Northeast, Northwest Atlantic, Mediterranean


Cosmopolitan in coastal tropical, subtropical and temperate waters (Eastern and Western Atlantic and Pacific, Indo-Pacific, Black Sea, Mediterranean)

Indo-Pacific, North Pacific, Northeast, Northwest Atlantic Mediterranean

Thinlip grey mullet (C. ramada)

Indo-West Pacific, East, Southwest Pacific

Indo-Pacific

Golden grey mullet (Chelon (Mugil) auratus)

Eastern Atlantic, Black Sea, Mediterranean


Thicklip grey mullet (Chelon (Mugil) labrosus)

Eastern Atlantic, Black Sea, Mediterranean



Eastern Atlantic, Mediterranean

Northeast Atlantic (Norway), Mediterranean (Greece)

Blue mackerel (S. australiasicus)

Indian Ocean, Indo-Pacific, Pacific

Northwest Pacific

Chub mackerel (Scomber japonicus)

Indo-Pacific, Eastern, Western Pacific

North Pacific (Japan)

Coleoid cephalopod mollusc families

Ommastrephidae Japanese flying squid (Todarodes pacificus (=Ommastrephes pacificus)

Indo-Pacific, North Pacific North Pacific

Octopodidae Common Octopus (Octopus vulgaris)

Europe, Mediterranean Mediterranean

Notes 1 Host geographical range (from FishBase, 2019) 2 Pathogen geographical range (from literature). NNV strains are reported from the Indo-Pacific (Australia, China, India, and Southeast Asia), Northeast and Northwest Atlantic, Europe and North America. They have not been reported from South Africa, South America or New Zealand (Hine & Diggles, 2005; Moody & Horwood, 2008; Sano et al., 2011).

Clinical signs

Infection with NNV causes a neurological disease which usually progresses with no external

clinical signs (Sano et al., 2011; Blackwell, 2019).

Pathogen survival

Serotype A NNV can be transmitted in dead and decomposing fish tissues (Chi et al., 2016) and

through fomites in aquaculture (OIE, 2019b). From laboratory experiments using artificial and

sterilised seawater, NNV remains viable in seawater (below 25 °C) for at least 12 months


(Frerichs et al., 2000). Survival is assumed to be at least 6 months and possibly longer (OIE,

2019b), but may be influenced in natural waters by dilution and predation factors (see section

6.1.2).

Transmission

Disease transmission may be vertical and horizontal, but life cycle details are poorly known

(Sano et al., 2011; OIE, 2019b). Wild fish may be reservoirs for infection. Farmed gilthead sea

bream (Sparus aurata) in Israeli aquaculture were infected at a prevalence of 20%, where the

infection was derived from adjacent wild fish stocks in the Mediterranean Sea (Berzak et al.,

2019).


Infection usually occurs through the water column. Disease progression may initially follow

several routes (Costa & Thompson, 2016). In clinical infection, NNV is neuro-invasive, affecting

the spinal cord, swim bladder, then progressing to the brain and retinal tissues. Viral particles are

released into the water column by carrier fish through gonad and digestive excretions (Sano et

al., 2011).

NNV infection is reported in aquaculture through use of infected fish bait of Family Clupeidae

and Japanese flying squid (Todarodes pacificus) (Family Ommastrephidae) as feed (Mori et al.,

2005; Gomez et al., 2010; OIE, 2019b).

Inactivation

No simple control methods are available (Doan et al., 2017). Nodaviruses are unaffected by

either short (168 hours), or medium-term (4 months) freezing (at -20 °C) (OIE, 2019b). NNV

strains are likely to be denatured by ionising radiation (at 50 kGy) (DAFF, 2013).

10.2 Risk assessment


NNV has a wide geographical distribution and a wide host range (Table 10) (Moody &

Horwood, 2008; Sano et al., 2011). Host specificity is low (Munday et al. 2002) and each NNV

genotype commonly affects several species within a geographical region (Shetty et al., 2012).

Prevalence is low-moderate (up to 21%) in wild fish and mollusc stocks (Castric, 1997; Chi et

al., 2016; Berzak et al., 2019), including fish bait species imported in high volumes (Tables 1 &

2).

NNV strains remain viable in frozen fish. Experimental data suggest survival in marine or fresh

waters may be up to 6 months (Frerichs et al., 2000; OIE, 2019b). Survival in natural waters is

unknown (see section 6.1.2). Therefore, the likelihood of entry is assessed as medium.


NNV strains remain viable during frozen storage, transportation and processing (Sano et al.,

2011). They remain viable in water and in decaying tissues for extended periods (OIE, 2019b).

Infection in farmed fish has been traced to the use of frozen wild-sourced fish and coleoid

cephalopods as feed in aquaculture (Castric, 1997; Mori et al., 2005; Gomez-Casado et al., 2011;

Chi et al., 2016; OIE, 2019b).


NNV strains may be dispersed by scavenging piscivorous birds, fomites, or through

contaminated fish processing equipment (Sano et al., 2011). The host range is wide (OIE, 2019b)

and the infectious dose is assumed to be low-moderate (105.5 TCID50 fish-1) (Nishi et al., 2016).

Viral titre is high in decaying fish (between 109.5 and 1010.0 TCID50 g-1) and in seawater (up to

105 TCID50 L-1) due to the high shedding rates of surviving fish (Nishi et al., 2016).

The likelihood of exposure and establishment is assessed as medium.



Infection with NNV is not an OIE-listed disease (OIE, 2019a) so establishment would not result

in restrictions on trade access for New Zealand. NNV strains cause disease with moderate-high

morbidity and mortality in over 70 species of marine and freshwater fish (Castric, 1997; Munday

et al., 2002; Gomez-Casado et al., 2011; Sano et al., 2011; Doan et al., 2017; Blackwell, 2019).

Establishment would likely have economic consequences for the developing marine aquaculture

of yellowtail kingfish (Seriola lalandi), hapuku/grouper (Polyprion oxygeneios) and snapper

(Pagrus auratus). (NIWA, 2017a, 2017b; Plant & Food, 2017).

Little is known about the effect of introduced viruses on wild fish populations (Lafferty et al.,

2015). Clinical disease is uncommon in wild stocks, but the prevalence of subclinical infection

varies widely (from 20% to 100%) (de la Pena et al., 2011; Berzak et al., 2019). Establishment

may result in decreased productivity and increased mortality in inshore commercial fisheries

including jack mackerel (Trachurus spp.) (export value NZ $68 million in 2018) and snapper

(Pagrus auratus) (export value $33 million in 2018) (Plant & Food, 2017; MPI, 2018a, 2018b,

2018c; Seafood New Zealand, 2019). Infected wild species may also act as a reservoir for

infection of farmed fish (Berzak et al., 2019).

NNV strains may also be dispersed by piscivorous birds (OIE, 2019b) into the freshwater

environment. Establishment may result in lowered productivity and increased mortality in grass

carp (Ctenopharyngodon idella) and silver carp (Hypophthalmichthys molitrix) farmed for use in

weed control (Clayton & Wells, 1999; Blackwell, 2019), as well as developing aquaculture of

eels (Anguilla spp.) (NIWA, 2017c). Other introduced cyprinids including goldfish (Carassius

auratus) may act as reservoir hosts (Doan et al., 2017).

Social consequences

NNV strains are not considered to be zoonotic. Their establishment may cause reduced

productivity and increased mortality in marine and freshwater fish and shellfish species of

importance for customary and recreational fishing (Wynne-Jones et al., 2019, 2019; Blackwell,

2019).



domestic product) in 2015 (Southwick et al., 2018).

The snapper, hapuku and trevally recreational fisheries contributed NZ$ 402 million, NZ$ 12.5

million and NZ$ 71 million respectively to the GDP in 2015 (Southwick et al., 2018). Several

species including snapper and eels (Anguilla spp.) are taonga for Maori culture and important for

customary fishing (MPI, 2019c). These fish stocks are considered stressed (MPI, 2018a, 2018b,

2018c) and further reduction in their abundance would have significant social effects for New

Zealand.



NNV strains affect a wide range of marine and freshwater fish (Blackwell, 2019). However, little

is known about the effects of viral infection in wild fish stocks (Lafferty et al., 2015). Species

such as clupeids perform an important role linking trophic levels in food webs and represent a

significant food source for other fish, seabirds and marine mammals (Varpe et al., 2005; Dunn et

al., 2012; Whitfield et al., 2012). Reduction in herring (C. p. pallasi) abundance (from over-

fishing) caused a reported 60% decrease in the relative abundance of wild salmon

(Oncorhynchus tshawytscha) in coastal British Columbia, with major biological, economic and

social consequences for humans and wildlife (Pagowski et al., 2019).

NNV strains affect invertebrates including bivalve and coleoid molluscs, crustaceans and annelid

worms (OIE, 2019b; Volpe et al., 2018; Blackwell, 2019), but the environmental significance is

unknown.

NNV strains may also be dispersed by piscivorous birds (OIE, 2019b) and affect freshwater

species including eels (NIWA, 2017c), where introduced cyprinids including goldfish (Carrasius

auratus) may act as reservoir hosts (Doan et al., 2017). The environmental significance of

establishment in freshwater species is unknown.

The establishment of NNV strains would have negative economic, social and environmental

consequences. The consequences of establishment are assessed to be medium.


The entry, exposure, establishment and consequence assessments for NNV strains are medium,

so the risk estimate is medium. Under the procedures followed in this risk analysis, risk

management measures may be justified.

10.3 Risk management

NNV strains have been assessed to be a risk in the commodity. Infection with NNV is a not an

OIE-notifiable disease, so the Aquatic Code (OIE, 2019a) provides no specific guidance or

processing requirements that would ensure the destruction of the virus.

NNV strains are associated with several finfish and cephalopod species (Table 10). Therefore,

restriction to non-susceptible species in the commodity should substantially reduce the risk

associated with NNV strains and be a viable risk management option.

NNV strains have a wide geographic range (Table 10). Their occurrence in wild stocks in other

regions is likely to be low. Restriction through the BNZ Country Approval Procedures to regions

where NNV strains have not been reported should substantially reduce the NNV risk and be a


The presence of NNV strains in a shipment of fish bait may be determined by batch testing,

using standard sampling procedures (Corsin et al., 2009; OIE, 2019a). Standard diagnostic

procedures for detection of the virus are available (Arimoto et al., 1982; Panzarin et al., 2010;

OIE, 2019b). Verification of a negative test from a shipment should substantially reduce the

NNV risk and be a viable risk management option.

NNV strains are unaffected by freezing (OIE, 2019b). Therefore, frozen storage (at -18 °C to -20

°C for either 168 hours, or for 4 months) is not a viable risk management option.


NNV strains are denatured by ionising radiation (at 50 kGy) (DAFF, 2013). This treatment

should substantially reduce the NNV risk and be a viable risk management option.



(section 6.2.1), one or more of the following risk management options would reduce the NNV


Option 1

• Competent Authority attestation that the commodity does not include: Atlantic horse

mackerel (Trachurus trachurus), Chilean jack mackerel (T. murphi), Japanese jack mackerel

(T. japonicus), Pacific jack mackerel (T. symmetricus) (Carangidae); Atlantic herring

(Clupea harengus) (Clupeidae); flathead grey mullet (Mugil cephalus), golden grey mullet

(Chelon auratus), thicklip grey mullet (Chelon labrosus), thinlip grey mullet (Chelon

ramada) (Mugilidae); blue mackerel (Scomber australasicus), chub mackerel (Scomber

japonicus), Atlantic mackerel (Scomber scombrus) (Scombridae), Japanese flying squid

(Todarodes pacificus) (Ommastrephidae); and common octopus (Octopus vulgaris)

(Octopodidae) should substantially reduce the risk associated with NNV strains, so the

commodity could be imported without any further restrictions.

Option 2


should substantially reduce the risk associated with NNV strains, so the commodity could

be imported without any further restrictions.

Option 3


diagnostic procedures, through the BNZ Country Approval Procedures should substantially

reduce the risk associated with NNV strains, so the commodity could be imported without

further restrictions.

Option 4.

• Treatment with ionising radiation (at 50 kGy) should substantially reduce the risk

associated with NNV strains, so the commodity could be imported without any further

restrictions.

Orthomyxoviridae: Infectious salmon anaemia virus (ISAV)



Infectious salmon anaemia virus (ISAV) is classified in the Genus Isavirus, within the Family

Orthomyxoviridae (OIE, 2019a). Two forms of ISAV are recognised: highly pathogenic HPR

(highly polymorphic region)-deleted ISAV, and low-virulent HPR0 ISAV (the non-deleted HPR

form of ISAV). Both forms may be present in fish populations (Cárdenas et al., 2014). Outbreaks

of ISAV may have resulted from the emergence of HPR-deleted ISAV from low-virulent HPR0

ISAV (Cunningham et al., 2002; Mjaaland et al., 2002; Christiansen et al., 2017; Gagné &

LeBlanc, 2018; OIE, 2019a).


Several distinct strains of each form have been isolated, based on geographical area (Miller &

Cipriano, 2003; OIE, 2019b).

11.1.2 OIE status

Infection with either HPR0 or HPR-deleted ISAV strain is an OIE-notifiable disease (OIE,

2019a).


New Zealand is considered free from both HPR0 and HPR-deleted forms of ISAV (Stone et al.,

1997; Norman et al., 2013). Infection with either form of ISAV is a notifiable disease in New

Zealand (Anon., 2016).


ISAV strains are not zoonotic (CABI, 2019b).

11.1.5 Epidemiology


The HPR0 and HPR-deleted strains of ISAV are widely distributed (Table 11). Prevalence varies

widely (from 5% to 90%), depending on pathogen strain and host species (Le Blanc et al. 2018).

The species associated with ISAV strains in fish bait and their geographic/pathogen range are

given in Table 11. Infection of Atlantic herring (C. harengus) with HPR0 strains mainly occurs in

close proximity to infected farmed salmonids (OIE, 2019b).

HPR-deleted strains are not reported from wild fish bait species (OIE, 2019a). However, Atlantic

herring (C. harengus) and Pacific herring (C. p. pallasii)) can be experimentally infected (by

bath immersion) with isolates from infected salmonids (Table 11) (Biacchesi et al., 2007; OIE,

2019a; CABI, 2019b). Other, as yet unknown marine host species may also function as

reservoirs of infection (OIE, 2019b)


Table 11. Fish bait species susceptible to strains of infectious salmon anaemia virus (ISAV)

HPR0 strains (low virulence)



Atlantic, cosmopolitan Northwest, Northeast Atlantic (Canada, Europe, Norway, UK, USA)

HPR-deleted strains (high virulence)



Atlantic, cosmopolitan Not reported from wild fish3


Indo-Pacific, Pacific, cosmopolitan

Not reported from wild fish3

Notes 1 Host geographical range (from FishBase, 2019) 2 Pathogen geographical range (from literature). The HPR0 strains are widely reported from wild salmonids in marine and

freshwater salmon growing regions including Norway, Scotland, Ireland, New Brunswick, Nova Scotia, Maine and Chile (OIE, 2019a). The HPR-deleted strains are widely reported from Atlantic salmon (Salmo salar) in the Northeastern Atlantic Ocean (Europe, the Faroe Islands), United Kingdom (Scotland), Middle East (Iraq), Northwest Atlantic Ocean (Canada in the Bay of Fundy), the United States (Maine), and South America (Chile) (OIE 2019a; Blackwell 2019).

3 Reported experimental infection of HPR-deleted strains of ISAV


Species at risk in New Zealand include brown trout (Salmo trutta) and rainbow trout

(Oncorhynchus mykiss) in marine and fresh waters, while Pacific herring C. p. pallasii) may

represent a reservoir for infection. Other Pacific salmon species including Chinook salmon (O.

tshawytscha) are considered resistant to HPR0 and HPR-deleted strains of ISAV (Rimstad et al.,

2011; Norman et al., 2013; Kibenge & Godoy, 2016).

Pathogenicity

HPR0 strains cannot be maintained in cell culture, so infectivity and pathogenicity processes are

poorly known (OIE, 2019b). HPR0 strains are of low virulence. They cause transient subclinical

infections in apparently healthy salmonids (Salmo spp.) in marine and fresh waters. All ages of

fish may be infected (OIE, 2019b). While these fish are not considered to be carriers (EFSA,

2012; OIE, 2019a), the role of HPR0 strains in the re-emergence of pathogenic HPR-deleted

strains remains unclear (Cunningham et al., 2002; EFSA, 2012; OIE, 2019a).

HPR-deleted strains cause long-term chronic disease in farmed Atlantic salmon (S. salar) with

associated cumulative mortalities of up to 90% (OIE, 2019a; Blackwell, 2019). Infection in non-

salmonids usually progresses without clinical signs (Cunningham et al., 2002). HPR-deleted

strains isolated from experimentally infected Pacific herring (C. p. pallasii) are infective to

salmonids (OIE, 2019b).

Clinical signs

Infection with HPR0 strains occurs with no clinical signs in non-salmonids (OIE, 2019b).

Infection with HPR-deleted strains in salmonids is characterised by exophthalmia and darkening

of the skin usually within 2-4 weeks post-infection. Infection in non-salmonids typically occurs

with no clinical signs of infection (OIE, 2019a).


The presence of ISAV may be detected by PCR in a range of species, including Chinook salmon

(O. tshawytscha), in the absence of clinical signs (Kibenge & Godoy, 2016). The significance of

these benign infections is uncertain, but they may be the source of clinical disease in farmed

Atlantic salmon (Salmo salar) (Murray et al., 2002; Kibenge & Godoy. 2016).

Pathogen survival

From experimental data, ISAV strains remain viable in marine or fresh water, for up to 4 months

in seawater at 4 °C (Rimstad & Mjaaland, 2002) and up to 14 days at temperatures up to 15 °C

(Falk et al., 1997; MacLeod et al., 2003; Rimstad et al., 2011). These strains survive up to 6 days

in fish tissues held on-ice (Tubbs et al., 2007; OIE, 2019a). They are sensitive to UV radiation

(Oye & Rimstad, 2001), but their survival in natural waters may be influenced by dilution and

predation factors (see section 6.1.2).

ISAV strains may remain viable for up to 4 months in moist decaying fish tissues dumped in

landfill (Rimstad & Mjaaland, 2002; Rimstad et al., 2011).

Transmission

Transmission may be horizontal or vertical, but the epidemiology of ISAV is currently unclear

(OIE, 2019a, 2018b). Dispersal may occur by fomites including fishing boats and gear, transport

and factory processing equipment, as well as through fish wastes, blood, melt and wash water

(OIE, 2019b). ISAV strains survive passage through the avian digestive system (OIE, 2019a) and

may be dispersed between waterways through piscivorous and scavenging birds (OIE, 2019a).


HPR0 strains targets the epithelial cells of the gills (Aamelfot et al. 2012), as well as kidney and

heart tissue (Christiansen et al., 2011; Lyngstad et al.,2011).

HPR-deleted strains cause chronic disease initially infecting the gills, skin surface and intestinal

tract. Disease then becomes systemic, affecting all major organs (OIE, 2019a).

Inactivation

ISAV strains are unaffected by short (168 hours), or medium-term (4 months) frozen storage (at

-18°C) and survive 5 freeze-thaw cycles with no loss in viability (Rimstad et al., 2011; OIE,

2019b). ISAV strains are assumed to be denatured by ionising radiation at 50 kGy (DAFF,

2013).



In fish bait, HPR0 strains are only associated with subclinically infected wild Atlantic herring

(C. harengus) (Table 11), where they are caught in close proximity to infected salmon farms

(OIE, 2019a). While no data are available on disease prevalence, it is reasonable to assume that

fishing for herrings is unlikely to take place close to marine salmon farms. The prevalence is

therefore likely to be low in wild stocks. The likelihood of entry of HPR0 strains is assessed to

be low.

HPR-deleted strains are not reported from fish bait (Table 11). The likelihood of entry of HPR-

deleted strains is assessed to be so low as to be negligible.



HPR0 strains are unaffected by frozen storage. They may remain viable in thawed fish tissues for

up to 6 days and in the water column for a further 7 days, within a temperature range from 4 °C

to 15 °C (MacLeod et al., 2003; Tubbs et al., 2007; OIE, 2019a).

Infection may also occur through fomites including fishing boats and gear, transport and factory

processing equipment, as well as through discarded bait. HPR0 strains may also be dispersed

through offal and discards into waterways by piscivorous birds (OIE, 2019b).

The risk of exposure and establishment of HPR0 strains is considered to be medium.



HPR0 ISAV is an OIE-listed disease, so establishment could result in restrictions on trade access

for New Zealand exports, as well as indirect economic consequences for the fishing, fish

processing and tourist industries. ISAV strains are associated with fish of families Clupeidae,

Gadidae and Salmonidae (Blackwell, 2019). New Zealand finfish aquaculture exports are

focused on Chinook salmon (O. tshawytscha) which is considered resistant to infection (OIE,

2019a), but ISAV presence may be detected by PCR analysis (Kibenge & Godoy, 2016).

While establishment of an OIE-listed disease may have serious economic consequences for New

Zealand trade, the HPR0 strains are of low pathogenicity (OIE, 2019a). The economic

consequences of their establishment on other commercial fisheries are assumed to be negligible.

Social consequences

ISAV strains are not zoonotic (CABI, 2019b). The HPR0 strains are unlikely to have a direct

effect on fish stocks considered significant for customary or recreational fishing (Wynne-Jones

et al., 2014, 2019). The social consequences of establishment are considered negligible.


The HPR0 strains of ISAV are associated with transient, subclinical infection which usually

occurs with no clinical signs (OIE, 2019a, OIE, 2019b). Prevalence in fish bait is likely to be

very low, so establishment would be unlikely to cause significant effects on wild fish (EFSA,

2012; OIE, 2019a, OIE, 3019b). The environmental consequences of establishment of the HPR0

strains are negligible.

The HPR0 strains of ISAV are of low pathogenicity and cause transient subclinical infection.

The economic, social and environmental consequences of establishment through fish bait are

assessed as negligible.


As the entry and consequence assessments are assessed to be negligible, ISAV and other

orthomyxoviruses are not considered to be risks in the commodity. No further assessment is

necessary.


Reoviridae: Piscine aquareovirus (PRV) and associated aquareoviruses



Piscine aquareovirus (PRV) is a group of seven closely associated species (Aquareovirus A -

Aquareovirus G), each comprised of several strains (Lupiani et al., 1993; Bustos et al., 2011; Ke

et al., 2011; King et al., 2011; Blindheim et al., 2015). The relationships between these species

and other associated but uncharacterised strains is unclear (Fauquet et al., 2005; Palacios et al.,

2010; Biering & Garseth, 2012; Kristoffersen et al., 2013). PRV strains are agents of heart and

skeletal muscle inflammation (HSMI), cardiomyopathy syndrome (CMS) and erythrocytic body

inclusion syndrome (EBIS). Co-infection with multiple strains may also occur (Wiik-Nielsen et

al., 2011).

12.1.2 OIE status

Disease caused by PRV strains is not notifiable to the OIE (OIE, 2019a).


PRV strains have not been reported from New Zealand (Cobb, 2008; B. Jones, pers. comm.,

2015). Infection with PRV is not a notifiable disease in New Zealand (Anon., 2016).


PRV strains are not zoonotic (Anon., 2020a).

12.1.5 Epidemiology


PRV strains are widely distributed (Table 12). They are reported from 12 families of fish

(Blackwell, 2019). Infection involves multiple carrier species and virus reservoirs (Roberts,

2012; Wiik-Nielsen et al., 2012; Zhang & Gui, 2015).

The species associated with PRV strains in fish bait and their geographic/pathogen range are

given in Table 12. The prevalence of PRV strains in wild Atlantic horse mackerel (T. trachurus)

and Atlantic herring (C. harengus) is low (Wiik-Nielsen et al., 2012; Carlile et al., 2014).


Table 12. Fish bait species susceptible to piscine aquareovirus and related strains


Carangidae Atlantic horse mackerel (Trachurus trachurus)

Mediterranean, East Atlantic Northeast, Northwest Atlantic (Canada, Denmark, Norway, United Kingdom, Spain)


Atlantic Northeast Atlantic (Denmark, Norway, United Kingdom, Spain)

Notes 1 Host geographical range (from FishBase, 2019) 2 Pathogen geographical range (from literature). PRV is reported from the Northeast Atlantic Ocean (Denmark, Norway,

Scotland, Spain), Northwest Atlantic (Canada); the Northeast and Central Pacific Ocean (Australia, China, Japan), and South America (Chile) (Chen & Jiang, 1984; Fauquet et al., 2005; King et al., 2011; Garseth et al., 2012; Roberts, 2012; Wiik-Nielsen et al., 2012; Carlile et al., 2014; Zainathan et al., 2014).


Susceptible species in New Zealand include salmonids, silverside (Argentina elongata),

mackerel (Trachurus spp.) and Pacific herring (C. p. pallasii) (Paul, 2000; Tubbs et al., 2007).

The New Zealand brill (Colistium guntheri) and turbot (C. nudipinnis) (Pleuronectidae) are

related to the Northern Hemisphere brill (Scophthalmus rhombus) and turbot (S. maximus)

(Pleuronectidae) (Chanet, 2003). Thus, it is reasonable to conclude that pathogens affecting

turbot and brill (Scophthalmus spp.) may infect endemic brill and turbot species (C. Johnston,

pers. comm., 2015).

Pathogenicity

PRV strains reported from salmonids are of high morbidity, low-medium pathogenicity, with a

mortality rate of 5–20% (Kongtorp et al., 2004a, 2004b; Wiik-Nielsen et al., 2012). Subclinical

infection also occurs in wild salmonids at low prevalence rates, with no clinical signs (Palacios

et al., 2010; Wiik-Nielsen et al., 2012; Anon., 2020a).

Pathogenicity is complex, varying among strains and between host species (Wiik-Nielsen et al.,

2012). It is also influenced by water temperature and the presence of co-infecting viral pathogens

(Crane & Carlisle, 2010; Wiik-Nielsen et al., 2012; Zainathan et al., 2014; Johansen et al., 2015;

Garver et al., 2016; Wessel et al., 2017).

PRV strains associated with Atlantic horse mackerel (T. trachurus) and Atlantic herring (C.

harengus) are of low pathogenicity (Carlile et al., 2004). Disease occurrence in these species is

not geographically related to disease occurrence in salmonids and their role as disease reservoirs

for adjacent farmed species is poorly understood (Wiik-Nielsen et al., 2012; Johansen et al.,

2015; Garver et al., 2016).

Clinical signs

Infection in Atlantic horse mackerel (T. trachurus) and Atlantic herring (C. harengus) occurs

with few or no external clinical signs (Garseth et al., 2012; Wiik-Nielsen et al., 2012; Carlile et

al., 2014).

Pathogen survival

From experimental data, PRV strains remain viable for almost 6 months in distilled water

(McDaniels et al., 1983), up to 13 days in seawater (Wiik-Nielsen et al., 2012) and up to 12

months in sewage sludge at 8 °C (Wellings et al., 1976). They remain viable in low moisture


processed foods for 2 months at 5 °C (Pirtle & Beran, 1991). Survival in natural waters may be

influenced by dilution and predation factors (see section 6.1.2), but PRV strains are likely to

remain viable in fish bait discarded into marine waters (Kongtorp et al., 2004a, 2004b; Wiik-

Nielsen et al., 2012).

Transmission

Transmission is horizontal and direct, through the water column (Watanabe et al., 2006; Garver

et al., 2016),


Infection in salmonids initially focuses on skeletal and heart muscle, causing ventricular lesions,

necrosis and myositis. This spreads to the gills, causing haemorrhages and necrosis. Infection

then becomes systemic, affecting the liver, spleen, kidney and intestine. (Wessel et al., 2017),

with haemorrhages of the jaws, operculum and fin bases (Rivas et al., 1996a, 1996b; Aldrin et

al., 2010; Kibenge & Godoy, 2016).

Inactivation

PRV strains remain viable after long-term frozen storage (at -80 °C) and survive 3 freeze-thaw

cycles (Hauge et al., 2016).) They are therefore assumed to be unaffected by either short (168

hours), or medium-term (4 months) frozen storage (at -20 °C). PRV strains are assumed to be

denatured by ionising radiation (at 50 kGy) (DAFF, 2013).



PRV strains occur at low prevalence in fish bait but have a wide geographical distribution (Wiik-

Nielsen et al., 2012). They are associated with Atlantic herring (C. harengus) and Atlantic horse

mackerel (T. trachurus) (Table 12). Imports of T. trachurus as fish bait in 2018 (Table 3)

represented over 10% by volume, while C. harengus was not imported.

PRV strains are unaffected by frozen storage. Infected fish may be present in fish bait at a low

rate of prevalence (Wiik-Nielsen et al., 2012).

The likelihood of entry of PRV strains is assessed as medium.


PRV strains are assumed to be unaffected by frozen storage for at least 4 months (at -20 °C) and

survive multiple freeze-thaw cycles (Hauge et al., 2016). From experimental data, PRV strains

remain viable for up to 13 days in seawater and for up to 12 months in marine sediments

(Wellings et al., 1976; Wiik-Nielsen et al., 2012). Survival in natural waters is unknown (section

6.1.2). Potential alternative hosts exist in New Zealand (Paul, 2000; Tubbs et al., 2007).

Prevalence is low in wild stocks and the strains associated with fish bait are of low

pathogenicity. Disease establishment is complex and likely to require co-infection with other

viral pathogens (Johansen et al., 2011; Wiik-Nielsen et al., 2012; Carlile et al., 2014; Johansen et

al., 2015; Garver et al., 2016; Wessel et al., 2017).

The likelihood of exposure and establishment is therefore assessed to be so low as to be

negligible.



As the likelihood of exposure and establishment of PRV strains through the commodity is

assessed to be negligible, no further risk assessment is necessary.

Rhabdoviridae: Infectious haematopoietic necrosis virus (IHNV)



Infectious haematopoietic necrosis virus (IHNV) is a single-stranded RNA virus classified in the

genus Novirhabdovirus, within the Family Rhabdoviridae (Bootland & Leong, 2011; OIE,

2019a). It was first identified from sockeye salmon (Oncorhynchus nerka) on the West Coast of

the United States (Rucker et al.,1953; Wingfield et al.,1969). One serotype is recognised

(Bootland & Leong, 2011), with several genogroups. These vary widely in pathogenicity and are

geographically based. They have a wide host range within each region (Bootland & Leong,

2011).

13.1.2 OIE status

Infection with IHNV is listed by the OIE as a notifiable disease (OIE, 2019a).


IHNV has not been reported from New Zealand (Boustead et al., 1993; Stone et al., 1997). It is

considered an exotic notifiable disease (Anon., 2016).


IHNV is not zoonotic (OIE, 2019a).

13.1.5 Epidemiology


IHNV occurs in 11 families of marine and freshwater fish (Blackwell, 2019), as well as from

marine and freshwater invertebrates (Bootland & Leong, 2011; OIE, 2019b).

IHNV is widely distributed (Table 13). Infection in non-salmonids commonly occurs without

clinical signs, so the host range is poorly known. Reports are based on targeted experimental

challenge and from incidental detection during surveillance programmes or surveys (Dixon et

al., 2016).

The species associated with IHNV in fish bait and their geographic/pathogen range are given in

Table 13 (Hart et al., 2011; OIE, 2019a).


Susceptible fish hosts in New Zealand include all salmonids and a wide range of other marine,

estuarine and freshwater fish (Tubbs et al., 2007).


Table 13. Fish bait species susceptible to infectious haematopoietic necrosis virus (IHNV)



Pacific Northeast, Northwest Pacific

Notes 1 Host geographical range (from FishBase, 2019) 2 Pathogen geographical range (from literature). IHNV is widely distributed in marine and fresh waters of North America

(Pacific and Northwest USA), South America (Bolivia, Chile), Asia (China, Japan, South Korea and Chinese Taipei (Taiwan), Europe (Austria, Croatia, Czech Republic, France, Germany, Italy, Netherlands, Poland, Slovenia and Spain), Iran, Kuwait, Russia and Pakistan (Tubbs et al., 2007; Dixon et al., 2016; OIE, 2019b).

Pathogenicity

Mortality in farmed fish varies widely (from 50% to 90%), depending on viral strain, host

species and environmental factors (Munro & Midtlyng, 2011; Dixon et al., 2016). Epizootics are

associated with water temperatures between 8 °C and 14 °C (Dixon et al., 2012).

Pathogenicity is much lower in wild stocks, where chronically infected surviving fish may act as

carriers (Bootland & Leong, 2011).

Clinical signs

Infection may occur with no clinical signs, but external lesions, skin darkening and petechial

haemorrhages at the base of the fins, vent, gills and mouth may be present (Bootland & Leong,

2011).

Transmission

Transmission can be vertical or horizontal (OIE, 2019b). Piscivorous birds and invertebrates

including mayflies (Callibaetis spp.), salmon lice (Lepeophtheirus salmonis), salmon leeches

(Piscicola spp.) and parasitic copepods (Salmonicola spp.) may function as vectors in marine and

fresh waters (Bootland & Leong, 2011; Kent et al., 1998; OIE, 2019a, OIE, 2019b).


The infective dose is relatively low (103 pfu ml-1) (Traxler et al., 1993; Bootland & Leong,

2011). Entry occurs through the epidermal cells of the gills, skin, fin bases, oral region, or

through the digestive system (Stone et al., 1997; Bootland & Leong, 2011).

Infection in salmonids is associated with the haematopoietic tissues (liver, spleen and kidney),

(Munro & Midtlyng, 2011). Infection may spread to brain tissue (Samuelsen et al., 2006).

The infectivity of IHNV in wild non-salmonid fish is poorly understood. Infection may be

transient and progress with no clinical signs (Garver & Wade, 2017; OIE, 2019b). The role of

non-salmonids as reservoir hosts for salmonids is uncertain (Garver & Wade, 2017).

From experimental data, strains isolated from eels (Anguilla spp.) are infective to salmonids

(OIE, 2019a). Experimental infection of salmonids (by bath immersion at a low dose rate (102 to

103 pfu ml-1 (plaque forming units) for 10 minutes) with strains from Pacific herring (C. p.

pallasii) did not cause clinical infection. However, an asymptomatic carrier state developed, with

low (<10%) mortality, at a prevalence of up to 70% (Bergmann et al., 2002; Bootland & Leong,

2011; Hart et al., 2011). These experimental data may not fully reflect disease progression in the

natural environment (Free et al., 2006; Dixon et al., 2016).


Pathogen survival

IHNV remains infective over a wide temperature range (from 3°C to 18°C) (Bootland & Leong,

2011; OIE, 2019a). From experimental data, it remains viable for up to 30 days in fresh waters,

14 days in marine or estuarine waters and up to 9 weeks in sediment (Stone et al., 1997;

Yoshimizu et al., 2005; OIE, 2019b). However, survival in natural waters is unknown, and may

be influenced by dilution and predation factors (see section 6.1.2).

Inactivation

IHNV remains viable in frozen fish brain tissue (to -20 °C for 5 months) and is resistant to

multiple freeze-thaw cycles (Stone et al., 1997; Tubbs et al., 2007; Dixon et al., 2016). It is

therefore assumed to be unaffected by either short (168 hours), or medium-term (4 months)

frozen storage (at -20 °C).

IHNV is assumed to be denatured by ionising radiation (at 50 kGy) in frozen fish tissue (DAFF,

2013).



IHNV remains viable in the frozen commodity (Stone et al., 1997; Bootland & Leong, 2011;

Dixon et al., 2016) and infection may occur with no external signs. The prevalence in fish bait is

unknown, but likely to be low (Lafferty et al., 2015).

The likelihood of entry is therefore assessed as low.


IHNV remains viable for several months in marine or fresh waters and sediments (Kahn et al.,

1999; Yoshimizu et al., 2005). IHNV strains are geographically related, with low apparent host

specificity among fish and invertebrate hosts (Bootland & Leong, 2011; OIE, 2019b). The

infectious dose is low (103 pfu ml-1) (Traxler et al., 1993; Bootland & Leong, 2011), but strains

vary widely in pathogenicity, morbidity and mortality (OIE, 2019a).

The epidemiology of IHNV in Pacific herring (C. p. pallasii) is poorly known and disease may

be transient (Garver & Wade, 2017). Surviving fish may remain carriers of disease (Bootland &

Leong, 2011; Mussely & Goodwin, 2012; Sim-Smith et al., 2014; Symonds et al., 2014).

Experimental data indicate IHNV isolates from Pacific herring can infect salmonids (Foot et al.,

2006; Hart et al., 2011; Dixon et al., 2012), but their potential to act as a reservoir host is

uncertain (Garver & Wade, 2017).

The likelihood of exposure and establishment is therefore assessed as low.



Infection with IHNV is an OIE-listed disease, so establishment would result in loss of market

access, with far-reaching consequences for the New Zealand economy. Establishment in

salmonid aquaculture would also result in lowered productivity and increased mortality, with

serious economic consequences through lost production. Salmonid aquaculture in New Zealand


was valued at $77 million in 2018 (Aquaculture New Zealand, 2019). Other significant indirect

costs would result from destocking, disinfection and ongoing monitoring to regain disease-free

status (OIE, 2019a).

IHNV has a wide host range affecting 11 families of fish (Blackwell, 2019; OIE, 2019a). Little is

known about the economic consequences on non-salmonid fish (Lafferty et al., 2015). These

may include losses in productivity and increased mortality in the developing aquaculture and

major inshore fishery for snapper (Sparus aurata) (Sparidae). Snapper exports from capture

fishing were valued at $33 million in 2018 (Seafood New Zealand, 2019).

Social consequences

IHNV is not zoonotic. Establishment in the major customary and recreational fisheries (Wynne-

Jones et al., 2014, 2019) including silver sea bream (snapper) (Pagrus auratus) in coastal waters,

and eels (Anguilla spp.) in fresh water (MPI, 2018b), would be likely to incur significant social

consequences. These species are considered taonga for Maori (MPI, 2013a, 2013b, 2013c).

Recreational fishing is the 5th most popular recreational activity for adults, involving 700,000


domestic product) in 2015 (Southwick et al., 2018). The snapper recreational fishery contributed

NZ$ 402 million to the GDP (Southwick et al., 2018).


Inshore fish species including clupeids are considered likely to encounter and consume discarded

fish bait. IHNV has a wide host range, but pathogenicity varies widely among strains and

between hosts (Bootland & Leong, 2011; Garver & Wade, 2017). The environmental

consequences for wild fish stocks are essentially unknown (Lafferty et al., 2015; Garver &

Wade, 2017).

Clupeids provide a significant food source for other fish, seabirds and marine mammals and a

major link between trophic levels in the aquatic food web (Dunn et al., 2012), so reduction in

biomass due to disease could have significant environmental consequences.

IHNV is an OIE-listed disease affecting a wide host range, with major economic, social and

environmental consequences for New Zealand. The consequences of establishment are assessed

as medium-high.


The entry, exposure, establishment and consequence assessments are low to medium-high. While

uncertainty remains about the pathogenicity, occurrence and epidemiology of the strains

associated with clupeid fish, the risk of entry and establishment of IHNV is estimated to be low-

medium. Under the procedures followed in this risk analysis, risk management measures may be

justified.


IHNV has been assessed to be a risk in the commodity. As an OIE-listed disease, the OIE

Aquatic Code (OIE, 2019a) provides some guidance on processing requirements that would

ensure destruction of the virus. However, the proposed heat treatments would make the

commodity unfit for purpose. These options are not considered further.


IHNV is unaffected by frozen storage (at -20 °C for at least 5 months) (Dixon et al., 2016).

Therefore, frozen storage at -20 °C, for either 168 hours, or for 4 months, is not a viable risk

management option.

Treatment of the commodity using ionising radiation (at 50 kGy) (DAFF, 2013) should

substantially reduce the IHNV risk and be a viable risk management option.

IHNV is associated with the commodity (Table 13). Therefore, restriction to non-susceptible

species in the commodity should substantially reduce the IHNV risk and be a viable risk

management option.

IHNV has a wide geographic range (Table 13). The occurrence is likely to be low or negligible

in other geographic regions. Restriction through the BNZ Country Approval Procedures to

regions where IHNV has not been reported should substantially reduce the IHNV risk and be a


The presence of IHNV in a shipment of the commodity may be determined by batch testing

using standard sampling procedures (Corsin et al., 2009; OIE, 2019a). Standard diagnostic

procedures for detection of the virus are provided in Emmenegger et al. (2000) and OIE (2019b).

Verification of a negative test from a shipment should substantially reduce the IHNV risk and be

a viable risk management option.



(section 6.2.1), one, or more of the following risk management options would reduce the ENV


Option 1

• Competent Authority attestation that the commodity does not include Pacific herring (Clupea

pallasii pallasii) (Clupeidae) should substantially reduce the IHNV risk, so the commodity

could be imported without any further restrictions.

Option 2


should substantially reduce the IHNV risk so the commodity could be imported without


Option 3


diagnostic procedures, through the BNZ Country Approvals Procedures should substantially

reduce the IHNV risk, so the commodity could be imported without further restrictions.

Option 4

• Treatment with ionising radiation (at 50 kGy) should substantially reduce the IHNV risk, so



Rhabdoviridae: Viral haemorrhagic septicaemia virus (VHSV)



Viral haemorrhagic septicaemia virus (VHSV) is a single-stranded RNA virus, classified in the

Genus Vesiculovirus, within the Family Rhabdoviridae (OIE, 2019a). It is the agent of viral

haemorrhagic septicaemia (VHS), an acute and economically significant haemorrhagic viral

disease of marine and freshwater fish (Einer-Jensen et al., 2004; Skall et al., 2005a, 2005b; OIE,

2019b). Four genotypes of VHSV are recognised, with several sub-lineages of varying

pathogenicity (OIE, 2019a).

14.1.2 OIE status

Infection with VHSV is listed by the OIE as a notifiable disease (OIE, 2019a).


Active surveillance is undertaken for VHSV in New Zealand, but VHS has not been reported

from the Southern Hemisphere (Stone et al., 1997; DAFF, 2005, 2014; OIE, 2019a). It has not

been identified in New Zealand (OIE, 2019a), and it is exotic. Infection with VHSV is a

notifiable disease in New Zealand (Anon., 2016).


VHSV is not zoonotic (OIE, 2019a).

14.1.5 Epidemiology

VHS is one of the most economically important diseases affecting marine and freshwater fish

(OIE, 2019a). VHSV strains in European freshwater aquaculture appear to have been transferred

from northern Pacific and Atlantic oceanic waters through marine fish used as feedstock for

aquaculture (Hedrick et al., 2003; Phelps et al., 2014). Management of salmonid fisheries in the

USA now includes movement restrictions on bait fish species as a disease control measure (Skall

et al., 2005a, 2005b; Bain et al., 2010; Phelps et al., 2013, 2014).


VHSV is associated with over 44 families of marine and freshwater fish (Blackwell, 2019), but

may potentially affect all temperate marine species (EFSA, 2008; OIE, 2019b; Anon., 2017a). It

also remains viable in filter-feeding oysters (Crassostrea gigas) and mussels (Mytilus edulis) in

Korea (Choi et al., 2015) without clinical signs for up to 7 days. Extracts from the mollusc

digestive gland failed to initiate infection (by intraperitoneal injection) in olive flounder

(Paralichthys olivaceus), suggesting the likelihood of infection of fish through this pathway is

low (Kim et al., 2017).

VHSV is essentially a cold-water pathogen, occurring where temperatures do not exceed 15 °C,

but it has also been introduced into Iranian and Kuwaiti aquaculture (Hedrick et al., 2003; Tubbs

et al., 2007; Amos et al., 2010; Spikler, 2010; OIE, 2019b). New Zealand oceanic water


temperatures vary between 17 °C in the north, to 12 °C in southern waters (Pinkerton et al.,

2018).

The species associated with VHSV in fish bait and their geographic/pathogen range are given in

Table 14 (Meyers et al., 1994, 1995; Traxler & Kieser, 1994; Meyers & Winton, 1995; Kocan et

al., 1997; Dixon et al., 1997; Marty et al., 1998; Hershberger et al., 1999; Mortensen et al.,

1999; Meyers et al., 1999; Traxler et al., 1999; Kocan et al., 2001; Biosecurity Australia, 2002;

Hedrick et al., 2003; Goodwin et al., 2004; Skall et al., 2005a, 2005b; Lee et al., 2007; Herve-

Claude et al., 2008; Kim & Faisal, 2010; Crane & Hyatt, 2011; Diggles et al., 2011; Purcell et

al., 2012; Kim et al., 2013a; Phelps et al., 2013; Ogut & Altuntas, 2014; Sandlund et al., 2014;

CIFA, 2019; OIE, 2019a).

Table 14. Fish bait species susceptible to viral haemorrhagic septicaemia virus (VHSV)



East, West Atlantic, Black Sea, Europe

Northeast, Northwest Atlantic (Canada, Europe, USA)

Bali sardinella (Sardinella lemuru), Indian oil sardine (Sardinella. longiceps)

North, East, West Indian Ocean, Indo-Northeast Indian Ocean

Indo-Pacific, Northwest, Central Pacific, Northeast Indian Ocean

Pacific herring (Clupea pallasii pallasii), Spotted sardinella (Amblygaster sirm),

North, West, Central Pacific Northeast Pacific (USA)

South American pilchard (Sardinops sagax)

Pacific Northeast Pacific (Canada)

Round sardinella (Sardinella aurita)

East, West Atlantic Northeast, Northwest Atlantic


Northwest Pacific (Korea), Northeast Pacific (Southern California)

Northwest, Northeast Pacific (Canada, Japan, South Korea, USA)



Northeast Pacific (Korea), Northwest Pacific (Canada, USA)

Notes 1 Host geographical range (from FishBase, 2019) 2 Pathogen geographical range (from literature). VHSV is reported from the North Atlantic Ocean (English Channel), Baltic

Sea, Northwest Pacific Ocean (Japan, South Korea), and Eastern Pacific Ocean (from Alaska to California). It has also been introduced into Iranian and Kuwaiti aquaculture (Hedrick et al., 2003; Tubbs et al., 2007; Amos et al., 2010; Spikler, 2010; OIE, 2019b).

The prevalence in wild Pacific herring (C. p. pallasii) varies from 0 to 4-8% (Hedrick et al.,

2003; Arkush et al., 2006), where viral titre in apparently healthy fish is 1,000 to 10,000 times

lower than from fish with active disease (Meyers et al., 1999). Epizootics in wild Pacific herring

are reported in Alaska (Meyers et al., 1999) and in Pacific sardine (S. sagax) in British

Columbia, and Canada (Hedrick et al., 2003).


A wide range of potential marine and freshwater hosts are present in New Zealand (Tubbs et al.,

2007).


Pathogenicity

VHSV causes mortalities ranging from 20% to 100% in marine and freshwater aquaculture.

Pathogenicity in wild fish is variable, but subclinically infected Pacific herring (C. p. pallasii)

are known to shed substantial quantities of virus. Wild fish species may act as reservoirs of

disease (Hershberger et al., 1999).

Mortality is temperature related (OIE 2019a). Experimental infection of salmonids, by

cohabitation with infected Pacific herring (C. p. pallasii) in conditions approximating natural

waters (at 10 °C) resulted in mortality of 81.5% (Gross et al., 2019). Similar experiments

undertaken at higher temperatures (20 °C and above) failed to initiate clinical disease (Arkush et

al., 2008).

Clinical signs

External clinical signs in salmonids include exophthalmia and widespread haemorrhages of the

skin, fin bases and eyes. However, clinically infected fish may show no external signs where

temperatures exceed 5 °C (Munday, 2002). Subclinical infection may progress with no external

signs (Arkush et al., 2006).

Pathogen survival

From experimental data, VHSV remains viable for up to 10 days in sediment at 4 °C (Afonso et

al., 2012; DAFF, 2005). Viability in marine waters varies from up to 13 days at 4 °C, to 1.5 days

at 20 °C). Viability in fresh water varies (from up to 40 days at 4 °C, to less than 1 day at 30 °C).

Infection in wild stocks is therefore considered unlikely where water temperatures exceed 18 °C

(Arkush et al., 2006; Hawley & Garver, 2008). VHSV is sensitive to UV radiation (Oye &

Rimstad, 2001). Viability in natural waters may also be influenced by dilution and predation

factors (see section 6.1.2).

Disease transmission

Transmission is horizontal and direct, through the water column (Arkush et al., 2005).

VHSV can be transferred through frozen fish bait (Phelps et al., 2013), although prevalence in

wild stocks is likely to be low (from 0 to 4-8%) (Arkush et al., 2006). VHSV can be dispersed by

fomites, through meltwater or discharges from commercial processing, ballast water discharges,

or by mechanical transfer through piscivorous birds (DAFF, 2005; Bain et al., 2010; Bowser et

al., 2010).

Frozen sardines (Clupea p. pallasii) from California have been imported as bait into South

Australia without the introduction of VHS. This has been attributed to: the low prevalence of

VHSV in wild fish stocks; the effect of medium-term frozen storage; and the temperature of the

receiving waters being higher than the threshold (18 °C) necessary for VHSV establishment

(DAFF, 2002; Arkush et al., 2006; Herve-Claude et al., 2008; Anon., 2010; Pinkerton et al.,

2018).


Pathogen entry occurs through the epithelial cells of the gills, viscera or skin. Fish bait species

(such as Pacific herring) swim in dense shoals and the infectious dose is low (101.5 pfu mL-1)

(DAFF, 2005). From experimental data, infection may be initiated by cohabitation in healthy fish

after only 1 hour of water borne exposure (Kocan et al., 1997; Hershberger et al., 2007; Hawley

& Garver, 2008). The viral titre in infected South American pilchard (Sardinops sagax) ranged

from 1.26 × 105 to 1.56 × 108 pfu g–1 of tissue (Arkush et al., 2006).


Disease may follow several alternative pathways, depending upon the VHSV strain and the host

species (DAFF, 2005; OIE, 2019a). Infection may be immediate and focussed at the point of

entry or be delayed until viral particles reach the endothelial cells of the vascular tissues, brain or

spleen (DAFF, 2005).

In acute infection, disease initially infects the haemopoietic tissues. It then spreads to the major

organs and musculature, causing extensive necrosis. Infection may also spread to the brain and

neural tissues (OIE, 2019a).

Chronic infection progresses slowly, affecting all major organs, usually with no external signs

(DAFF, 2005). Surviving fish are carriers and continue to shed viral particles through urine,

faeces, mucous and sexual fluids, for months or years (DAFF, 2005).

Homogenates derived from infected fish are infectious to a wide range of species (Oidtmann et

al., 2011). Strains isolated from European marine fish are of low virulence for Atlantic salmon

(Salmo salar) (Dale et al., 2009), but strains from Pacific herring are virulent to salmonids.

Caution is advised before assuming a species is resistant to infection (Skall et al., 2005a, 2005b).

Inactivation

Short-term frozen storage (at -20 °C for 16 hours) is only partially effective against VHSV

(Wolf, 1988; Arkush al., 2006; Phelps et al., 2014; OIE, 2019a). Viral titre is reduced (by up to

90%) following repeated freeze-thaw cycles (Meyers et al., 1994). Studies approximating

commercial frozen storage of sardine Sardinops sagax used as bait indicated VHSV was

completely inactivated after 50 days (7 weeks) frozen storage (at -20 °C) (Arkush et al. 2003).

However, other studies indicate sufficient VHSV may remain viable in thawed fish tissues to

initiate infection where viral titre is high (Meyers at al., 1994; Arkush et al., 2006; Herve-Claude

et al., 2008; Phelps et al., 2013). VHSV is also inactivated where the receiving water

temperature exceeds 18 °C (Herve-Claude et al., 2008; Anon., 2010) and both frozen storage and

the receiving water temperature are recognised as significant risk factors (Anon., 2010). VHSV

is therefore assumed to be substantially, but not completely inactivated by medium-term frozen

storage (for 4 months at -20 °C).

VHSV is assumed to be denatured by ionising radiation (at 50 kGy) of frozen fish tissue (DAFF,

2013).



VHSV may be present in the commodity (Table 14). While Pacific herring was not imported

during 2018, clupeids represented over 50% (by volume) of the finfish imported as bait in 2018

(Table 3).

Prevalence in wild stocks is low (from 4-8%) in wild Pacific herring (C. p. pallasii) except

during epizootics (Hedrick et al., 2003; Arkush et al., 2006), but VHSV occurs in both Pacific

herring (C. p. pallasii) and South American pilchard (S. sagax) (Meyers et al., 1999).

VHSV may be present in carrier fish (Hershberger et al., 1999; Hedrick et al., 2003; Arkush et

al., 2006; Oidtmann et al., 2011; Phelps et al., 2013). VHSV may also be introduced through

meltwater and water used in processing fish bait (OIE, 2019a). Viability is substantially reduced

following frozen storage (Arkush et al., 2006).


VHSV may remain viable in the commodity. However, prevalence is generally low in wild fish

bait stocks and viability is substantially reduced following frozen storage (Arkush et al. 2006;

Herve-Claude et al., 2008), The likelihood of entry is assessed to be low.


VHSV is recognised as a major risk organism in the international fish bait trade (Biosecurity

Australia, 2002; Dalton, 2004; Goodwin et al., 2004; Arkush et al., 2006; Anon., 2010;

Oidtmann et al., 2011; Phelps et al., 2013, 2014). VHSV remains viable in marine and fresh

water, or in sediments for extended periods, where water temperatures do not exceed 18 °C

(DAFF, 2005; Arkush et al., 2006; Herve-Claude et al., 2008; Hawley & Garver, 2008; Afonso

et al., 2012).

The infectious dose is low (101.5 pfu mL-1) (DAFF 2005) and VHSV has a wide host range (OIE,

2019a). Wild fish species may act as reservoirs of disease (Hershberger et al.,1999) and infection

may progress without clinical signs of disease (Arkush et al., 2006).

VHSV can be transferred through frozen fish bait (Herve-Claude et al., 2008; Anon., 2010;

Phelps et al., 2013). It is substantially inactivated by long-term storage (at -20 °C for at least 50

days) (Arkush et al., 2006; Herve-Claude et al., 2008; Anon., 2010), but sufficient VHSV may

remain viable where viral titre is high to establish infection where the temperature of the

receiving water is less than 18 °C (Arkush et al., 2006; Herve-Claude et al., 2008; Hawley &

Garver, 2008), such as Chile or New Zealand (Stone et al., 1997; Herve-Claude et al., 2007;

Pinkerton et al, 2018).

The use of frozen menhaden (Brevoortia tyrannus) as feed in Australian aquaculture is restricted

to marine waters where temperatures exceed 14 °C, due to concerns about VHSV infection

(Diggles, 2007). Movement controls (restricting bait to the water body in which it was caught)

have been placed on bait in the continental USA, while additional processing is also required for

imported fish bait in some US States (Phelps et al., 2013, 2014).

The likelihood of exposure and establishment through the commodity is assessed as low.



Infection with VHSV is an OIE-listed disease primarily affecting salmonids (OIE, 2019a). Its

establishment would result in loss of market access for New Zealand, with significant direct

economic consequences. It is highly pathogenic, with mortalities up to 100% in salmonid

aquaculture (OIE, 2019a).

Other consequences of establishment include reduced productivity and increased mortality, as

Chinook salmon (O. tshawytscha) is particularly susceptible to VHSV (OIE, 2019a). New

Zealand salmonid aquaculture of Chinook salmon (O. tshawytscha) was worth $77 million in

export earnings in 2018 (Seafood New Zealand, 2019). Further indirect economic costs would

include fallowing and destocking, as well as disinfection of farms and processing equipment in

the supply chain.

VHSV has a wide host range in marine and fresh waters (OIE, 2019a, 2018b; Blackwell, 2019).

Wild stocks may represent reservoirs of infection for other farmed species (Hershberger et al.

1999). Establishment may affect the developing aquaculture of brill (Colistium guntheri) and


turbot (C. nudipinnis) (Pleuronectidae), hapuku/grouper (Polyprion oxygeneios) (Polyprionidae),

snapper (Sparus aurata) (Sparidae) and yellowtail kingfish (Seriola lalandi) (Carangidae).

Several wild fisheries may also be affected in marine and fresh waters (Diggles, 2003. 2004a,

2004b; Hickman & Tait, 2008; Symonds et al., 2014; NIWA, 2017b; Plant & Food, 2017; MPI,

2018a, 2018b, 2018c). For example, the snapper export fishery was valued at $33 million in

2018 (Seafood New Zealand, 2019).

Social consequences

VHSV is not zoonotic (OIE, 2019a). It has a wide host range, including many coastal marine fish

species (Tubbs et al., 2007). While effects on wild fish species are uncertain (Lafferty et al.,

2015), establishment is likely to result in lowered productivity and increased mortality for

susceptible species of significance for customary and recreational fishing (Wynn-Jones et al.,

2014, 2019; MPI, 2018a, 2018b, 2018c).



domestic product) in 2015 (Southwick et al., 2018). The snapper, hapuku and trevally

recreational fisheries contributed NZ$ 402 million, NZ$ 12.5 million and NZ$ 71 million

respectively to the GDP in 2015 (Southwick et al., 2018).


VHSV is highly pathogenic and potentially affects all temperate fish species (OIE, 2019b).

Establishment may result in lowered productivity and increased mortality in a wide range of

marine and freshwater fish species, many of which are considered stressed (MPI, 2018a, 2018b,

2018c). While effects of introduced viruses on wild fish stocks are uncertain (Lafferty et al.,

2015), VHSV has caused epizootics in Pacific herring (C. p. pallasii) and in South American

pilchard (S. sagax) (Meyers et al., 1999; Hedrick et al., 2003).

Inshore fish species including clupeids and mugilids provide a significant food source for other

fish, seabirds and marine mammals and are a major link between trophic levels in the aquatic

food web (Dunn et al., 2012). Wild fish may also act as reservoirs of disease (Hershberger et al.,

1999).

The establishment of VHSV is likely to have significant direct and indirect economic, social and

environmental effects for New Zealand. Therefore, the consequences of establishment are

assessed to be high.


The entry and exposure assessments for VHSV are low, but the consequences are assessed to be

high. The risk estimate is assessed to be medium. Under the procedures followed in this risk

analysis, risk management measures may be justified.


VHSV has been assessed as a risk in the commodity. VHS is an OIE-listed disease, so the OIE

Aquatic Code and Aquatic Manual provide some guidance to avoid the spread of VHSV (OIE,

2019a, 2019b). As the OIE-recommended heat treatment (OIE 2019a) would render the

commodity unfit for purpose as fish bait, this option is not considered further.


VHSV is associated with fish bait (Table 14). Short-term frozen storage (for 168 hours) is

unlikely to be effective in denaturing VHSV (Arkush et al., 2006; Phelps et al., 2013). Medium-

term frozen storage (-18 °C to -20 °C) for at least 4 months (16 weeks) should denature most

VHSV, but sufficient virus may remain present to initiate infection where viral titre is high

(Arkush et al., 2006; Herve-Claude et al., 2008; Anon., 2010).

The VHSV Working Group recommended batch testing of frozen fish bait (Anon., 2010). When

combined with medium-term frozen storage (see section 6.2.1), batch testing should substantially

reduce the risk associated with VHSV and be a viable risk management option.

Treatment of the commodity using ionising radiation (at 50 kGy) (DAFF, 2013) should

substantially reduce the VHSV risk and be a viable risk management option.

14.3.1 Risk management options.


(section 6.2.1), one or more of the following risk management options would reduce the VHSV


Option 1

• Competent Authority attestation that the commodity does not include Atlantic herring

(Clupea harengus), Bali sardinella (Sardinella lemuru), Indian oil sardine (Sardinella.

longiceps), Pacific herring (Clupea pallasii pallasii), spotted sardinella (Amblygaster sirm),

South American pilchard (Sardinops sagax), round sardinella (Sardinella aurita)

(Clupeidae); flathead grey mullet (Mugil cephalus) (Mugilidae); and Chub mackerel

(Scomber japonicus) (Scombridae) should substantially reduce the IHNV risk, so the

commodity could be imported without any further restrictions.

Option 2


should substantially reduce the VHSV risk so the commodity could be imported without


Option 3


diagnostic procedures through the BNZ Country Approvals Procedures, should substantially

reduce the VHSV risk, so the commodity could be imported without further restrictions.

Option 4

• Treatment with ionising radiation (at 50 kGy) should substantially reduce the VHSV risk, so


Edwardsiella spp.



Edwardsiella spp. are a species complex of Gram-negative rod-shaped filamentous bacteria

classified in the Family Enterobacteriaceae (Farmer et al., 1984, 1985). Five genetically distinct


but phenotypically ambiguous species are recognised (E. ictaluri, E. tarda, E. piscida, E.

hoshinae and E. anguillarum) with high intergenomic heterogeneity (Reichley et al., 2017;

Katharios et al., 2019). They vary widely in their responses to environmental parameters

(salinity, temperature, and pH), host specificity, geographical range, specific target organs, drug

resistance and pathogenicity (Park et al., 1983; Reichley et al., 2017).

High resolution phylogenetic methods, such as multilocus sequence analysis (MLSA),

multilocus sequence typing (MLST) and whole genome sequencing (Shao et al., 2015) are

necessary to identify species within the species complex (Katharios et al., 2019). For the

purposes of this risk analysis, these species are considered together as Edwardsiella spp.

15.1.2 OIE status

Infection with Edwardsiella spp. is not notifiable to the OIE (OIE, 2019a).


Edwardsiella spp. are reported from migratory seabirds and cetaceans that inhabit Antarctic and

Sub-Antarctic waters and may occasionally enter New Zealand waters (Sakazaki & Tamura,

1975; Shao et al., 2015; EPA, 2015; Tindall, 2016).

Edwardsiella spp. have not been reported from fish in Antarctic and Sub-Antarctic waters (S.

Fenwick, Massey University, reported in Stone et al., 1997; Leotta et al., 2009; Cools et al.,

2013). They have not been reported from New Zealand marine or freshwater fish (Johnston,

2008; M. Bestbier, BNZ, pers. comm., 2020). Therefore, for the purposes of this risk analysis,

Edwardsiella spp. are assumed to be exotic. Infection with Edwardsiella spp. is not a notifiable

disease in New Zealand (Anon., 2016).


Edwardsiella spp. are zoonotic. Bacterial gastroenteritis may result from eating undercooked

aquatic products. Wound infections associated with fishing and fish processing can cause

cellulitis and gas gangrene (Jordan & Hadley, 1969; Nagel et al., 1982). Infection is rarely

serious but may progress to systemic conditions including meningitis, septicaemia and

osteomyelitis (Janda et al., 1993; Hirai et al., 2015).

15.1.5 Epidemiology


Edwardsiella spp. are reported from marine and freshwater invertebrates, fish, amphibians,

reptiles and marine mammals. They also occur in oceanic birds, as well as terrestrial animals

including cattle and swine (Blackwell, 2019). Edwardsiella spp. have a wide geographical

distribution (Table 15).

The species associated with Edwardsiella spp. in fish bait and their geographic /pathogen range

are given in Table 15.

https://www.frontiersin.org/articles/10.3389/fmicb.2019.00141/full#B46


Table 15. Fish bait species susceptible to Edwardsiella spp.



Indo-Pacific Indo-Pacific (India)


Cosmopolitan in coastal waters of the tropical, subtropical and temperate waters of all seas.


Notes 1 Host geographical range (from FishBase, 2019) 2 Pathogen geographical range (from literature). Edwardsiella spp. are widely distributed, in over 30 countries, as well as from

the Antarctic, Atlantic, Indian and Pacific Oceans (Leotta et al., 2009).

Potential host fish species in New Zealand

A wide range of potential host fish species are present in New Zealand marine and freshwaters,

including members of Carangidae (yellowtail kingfish, Seriola lalandi), Pleuronectidae (turbot,

Colistium nudipinnis), Salmonidae (Oncorhynchus spp., Salmo spp., Salvelinus spp.), Sparidae

(snapper, Pagrus (Chrysophrys) auratus) and Polyprionidae (hapuku, Polyprion spp.) (Tubbs et

al., 2007; Blackwell, 2019).

Pathogenicity

Pathogenicity in farmed Chinook salmon (O. tshawytscha) varies from 5% to 15%. The lethal

dose (LD50) is low, estimated at 4.1 × 106 cells for chinook salmon (Park et al., 2012) and 1 ×

105 cells for gilthead seabream (Sparus aurata) (Baya et al., 1997).

Edwardsiella spp. are generally regarded as opportunistic pathogens of lower pathogenicity in

wild fish stocks (Amandi et al., 1982; Park et al., 2012). Epizootics occur in grey mullet (Mugil

cephalus) in Japanese coastal waters (Kusuda et al., 1976).

Clinical signs

External signs may include white skin patches and cutaneous lesions, but disease expression

varies widely between pathogen strains and among host species. Infection may also progress

with no external clinical signs (Evans et al., 2011).

Pathogen survival

Edwardsiella spp. are environmentally resistant. From experimental data, they remain viable in

marine or fresh waters up to 76 days, over a wide temperature range (from 45 °C to Antarctic

waters). They remain viable on dead and moribund fish tissues and survive in sediments in a

viable but not culturable (VBNC) state for extended periods (Evans et al., 2011). However, the

length of survival under natural conditions is unknown (see section 6.1.2).


Disease transmission is horizontal and direct, through the water column (Amandi et al., 1982).

Edwardsiella spp. may be dispersed by fomites, through contaminated water, aerosol

transmission, or by piscivorous birds (Johnston, 2008; Evans et al., 2011).Edwardsiella spp. are

opportunistic pathogens with a wide host range and may become ubiquitous in endemic areas

(Buller, 2014; Wiedenmayer et al., 2006; Harai et al., 2015).



Edwardsiella spp. are significant bacterial pathogens affecting farmed marine and freshwater

fish of all ages (Plumb & Hansen, 2011). From experimental infection per os, the infectious dose

is very low (from 1 × 101 to 9 × 107 cells) for channel catfish (Ictalurus punctatus) (Park et al.,

2012). Edwardsiella spp. infection in aquaculture is usually associated with poor rearing

conditions including high water temperature, poor water quality and high organic content (Meyer

& Bullock, 1973; Rashid et al., 1994; Evans et al., 2011).

Disease progression may follow a renal or hepatic pathway. In renal infection, micro-abscesses

form in kidney tissues. In hepatic infection, large abscesses develop in the liver, with extensive

tissue liquefaction (Miyazaki & Egusa, 1976; Evans et al., 2011). In both pathways, infection

becomes systemic, infecting the spleen, liver, heart and digestive tissue mucosa. Necrotic lesions

may also form in the lateral musculature (Miyazaki & Kaige, 1985).

Inactivation

Edwardsiella spp. are moderately cold-resistant. They survive frozen storage (up to 50 days

at -20 °C) in frozen fish tissues (Castro et al., 2006; Johnston, 2008), but are rendered non-viable

by longer-term frozen storage (at least 50 days) (Brady & Vinitnantharat, 1990). It is assumed

therefore that Edwardsiella spp. would be inactivated by medium-term frozen storage (at -20 °C

for 4 months).

Ionising radiation (at 50 kGy) is also likely to denature Edwardsiella spp. in frozen fish tissues

(DAFF, 2013).



Edwardsiella spp. are associated with the commodity (Table 15). Prevalence may reach 14% in

wild Indian oil sardine (Sardinella longiceps) (Clupeidae) in the Bay of Bengal during epizootics

(Kumar et al. 2014).

The prevalence in flathead grey mullet (Mugil cephalus) (Mugilidae) in Japanese waters is

unknown but assumed to be low (Kusuda et al., 1976; Evans et al., 2011). M. cephalus

represented 13% and S. longiceps represented less than 1%, respectively, by volume of finfish

bait imports from all countries in 2018 (see Table 3). Edwardsiella spp. strains remain viable in

dead and moribund fish tissues (Evans et al., 2011) and in fish bait frozen for up to 50 days

(Castro et al., 2006; Johnston, 2008). Infected fish may show no external signs of disease (Brady

& Vinitnantharat, 1990).

Edwardsiella spp. have a limited distribution where associated with fish bait and the prevalence

in healthy wild stocks is assumed to be low.

The likelihood of entry is assessed as low.


Edwardsiella spp. are non-obligate opportunistic pathogens. From experimental data, they

remain viable in a resistant VBNC state in the aquatic environment for extended periods, over a

wide temperature range (from 0 °C to 45 °C) (Evans et al., 2011; Park et al., 2012). Survival in

natural waters is unknown (see section 6.1.2).


Infection occurs through the water column, by cross-contamination of working surfaces (where

fish bait is used on fishing vessels), through the discharge of contaminated meltwater (Johnston,

2008; Evans et al., 2011). Edwardsiella spp. may be dispersed from landfill or between

waterways by piscivorous birds (Evans et al., 2011).

Edwardsiella spp. are zoonotic pathogens and human exposure may occur through cross-

contamination or handling of infected fish bait (Janda & Sharon, 1993; Hirai et al., 2015).

The host range in fish bait species is small and the prevalence is low in healthy wild fish stocks.

The likelihood of exposure and establishment is assessed as low-medium.



Edwardsiella septicaemia is not listed by the OIE (OIE, 2019a), so establishment would not

result in trade restrictions for New Zealand. However, Edwardsiella spp. have a wide host range

affecting farmed marine and freshwater fish worldwide. Infection results in lowered productivity

and increased mortality. Chronic infection causes tissue spoilage that makes the product

commercially unmarketable (Evans et al., 2011; CABI, 2015).

Establishment may result in serious economic consequences for farmed and fished species in

New Zealand, including salmonids. These include Chinook salmon (Oncorhynchus tshawytscha)

(Salmonidae) which was valued at $77 million in exports in 2018 (Seafood New Zealand, 2019).

However, the prevalence of clinical disease in farmed fish overseas is commonly related to over-

stocking and poor management practices (Evans et al., 2011).

Other potentially affected farmed species include yellowtail kingfish (Seriola lalandi)

(Carangidae), brill (Colistium guntheri) and turbot (C. nudipinnis) (Pleuronectidae) and gilthead

sea bream (snapper) (Sparus aurata) (Sparidae) (Diggles, 2003; Hickman & Tait, 2008; Mussely

& Goodwin, 2012; Sim-Smith et al., 2014; Symonds et al., 2014; NIWA, 2017a, 2017b; Plant &

Food, 2016, 2017). Establishment could also affect grass carp (Ctenopharyngodon idella) and

silver carp (Hypophthalmichthys molitrix) aquaculture for weed control in fresh waters (Clayton

& Wells, 1999; NIWA, 2014).

While prevalence of Edwardsiella spp. is generally low in wild fish stocks, epizootics occur in

wild flathead grey mullet (Mugil cephalus) (Evans et al., 2011). The establishment of

Edwardsiella spp. may result in reduced productivity, increased mortality and increased product

rejection in New Zealand coastal inshore fisheries (MPI, 2018a, 2018b, 2018c). For example, the

snapper export fishery was valued at $33 million in 2018 (Seafood New Zealand, 2019).

Social consequences

Edwardsiella spp. are zoonotic, generally causing minor bacterial gastroenteritis and cellulitis

through handling infected fish (Jordan & Hadley, 1969; Nagel et al., 1982). Serious infection is

rare, but may include meningitis, septicaemia and osteomyelitis (Janda et al., 1993; Hirai et al.,

2015).

Establishment of Edwardsiella spp. may affect freshwater aquaculture. This includes brown trout

(Salmo trutta) and rainbow trout (Oncorhynchus mykiss) farmed for recreational fishing (Fish &

Game, 2014). The Lake Taupo trout fishery alone is valued at NZ$ 70–80 million annually

(Marsh & Mkwra, 2013).


The establishment of Edwardsiella spp. may affect other species of significance for customary

and recreational fishing in marine and freshwater, including eels (Anguilla spp.). These are

considered stressed by the level of current fishing activity (Wynn-Jones et al., 2014, 2019;

NIWA, 2017c). Eels (Anguilla spp.) are considered a taonga in Maori culture (MPI, 2018a,

2018b, 2018c)

Recreational fishing is the 5th most popular recreational activity for adults, involving 700,000


domestic product) in 2015. Of this, snapper contributed NZ$236 million and trevally NZ$46

million to the GDP in 2015. These fisheries provided an estimated total of NZ$ 402 million and

NZ$ 71 million respectively, to the New Zealand economy in 2015 (Holdsworth et al., 2016;

Southwick et al., 2018).


Edwardsiella spp. are opportunistic pathogens of generally low pathogenicity in wild fish stocks

(Park et al., 2012). The infectious dose is very low (Park et al., 2012) and the host range includes

20 families of marine and freshwater fish (Evans et al., 2011; Blackwell, 2019). Edwardsiella

spp. may become endemic, with reported prevalence of up to 60% in wild freshwater fish

(Vandepitte et al., 1980). Wild fish may also function as reservoir hosts for human and animal

infection, including aquaculture (Vandepitte et al., 1980; Evans et al., 2011; Cools et al., 2013).

Edwardsiella spp. are opportunistic, zoonotic pathogens with a wide host range. While of low

pathogenicity in wild fish stocks, they may become significant pathogens of farmed fish,

particularly when aquaculture occurs under sub-optimum conditions.

The consequences of establishment are assessed as low.


As the entry, exposure and consequence assessments are low, or low-medium, the risk estimate

is for Edwardsiella spp. is assessed to be low. Under the procedures followed in this risk

analysis, risk management measures may be justified.


Edwardsiella spp. have been identified as risk organisms in previous aquatic risk analysis

(Johnston, 2008). Infection with Edwardsiella spp. is not an OIE-notifiable disease, so the OIE

Aquatic Code (OIE, 2019a) provides no specific guidance or processing requirements that would

ensure the destruction of the pathogen.

Edwardsiella spp. are associated with fish bait (Table 15). Edwardsiella spp. are unlikely to be

denatured by short-term frozen storage (at -20 °C for 168 hours). They are likely to be

substantially or completely denatured by medium-term frozen storage (to -20 °C for at least 4

months) (Brady & Vinitnantharat, 1990). An extended period of frozen storage is consistent with

current fish bait industry practice (B. Burney, pers. comm., 2020). Adoption of this general risk

management option should substantially reduce the risk associated with Edwardsiella spp. and

be a viable risk management option.


If the general risk management measures proposed for all imported fish bait (section 6.2.1) are

adopted, including medium-term frozen storage (at -18 to -20 °C for four months), then no


pathogen-specific risk management measures would be necessary to reduce the risk associated

with Edwardsiella spp. to an acceptable level.

Francisella spp.



Francisella spp. represent a species complex of rickettsia-like intracellular bacteria classified in

Family Francisellaceae, within the Proteobacteriaceae (Arkush & Bartholomew, 2011). The host

relationships and distribution within this species complex are unclear (Ottem et al., 2009; Arkush

& Bartholomew, 2011; Colquhoun & Duodu, 2011). Strains of Francisella piscida (= F.

noatunensis) associated with finfish include F. noatunensis orientalis in warm water fish species

and F. noatunensis noatunensis in cold water fish species (Colquhoun & Duodu, 2011).

16.1.2 OIE status

Infection with Francisella spp. is not listed by the OIE (OIE, 2019a).


Francisella spp. are considered exotic (Johnston, 2008) and have not been reported from New

Zealand. Infection with Francisella spp. is not a notifiable disease in New Zealand (Anon.,

2016).


Francisella spp. affecting fish are not considered zoonotic (Colquhoun & Duodu, 2011).

16.1.5 Epidemiology


Francisella spp. are reported from 9 families of marine and freshwater fish and invertebrates

(Johnston, 2008; Colquhoun & Duodu, 2011; Blackwell, 2019). They have a wide geographical

distribution (Table 16).

The species associated with Francisella spp. in fish bait and their geographic /pathogen range are

given in Table 16. Prevalence in wild Atlantic cod (Gadus morhua) reached 20% off the

Swedish coast in 2004 (Alfjorden et al., 2006), while lower levels of prevalence (from 7-11%)

were reported in Atlantic cod from coastal Norwegian waters (Ottem et al., 2008).

Francisella spp. is also reported at a prevalence of 11%, in Atlantic mackerel (S. scombrus) from

areas immediately adjacent to infected Atlantic cod farms. However, these prevalence data were

determined from a very small sample size (n=9 fish) The bacterial load in Atlantic mackerel was

much lower than for infected Atlantic cod (Colquhoun & Duodu, 2011).

Strains of Francisella spp. may be host specific. Given the small sample size available for

Atlantic mackerel (Ottem et al., 2008), the significance of this infection is difficult to estimate

(Colquhoun & Duodu, 2011).


Table 16. Fish bait species susceptible to Francisella spp.



North Atlantic, Mediterranean Sea

Northeast Atlantic (Europe)

Notes 1 Host geographical distribution (from FishBase 2019) 2 Pathogen geographical range (from literature). Francisella spp. are reported from the East, South and West Atlantic (Central

and South America, Norway, United Kingdom, United States), and the North, Central and South Pacific (Asia, China, Hawaii, Central and South America) (Colquhoun & Duodu, 2011).

Potential fish host species in New Zealand

Potential hosts in New Zealand include Atlantic salmon (Salmo salar) and brown trout (S.

trutta), as well as mackerel (Scomber spp.) and flatfishes (Order Pleuronectiformes) (Colquhoun

& Duodu, 2011).

Pathogenicity

Francisella spp. infects farmed Atlantic cod (G. morhua) with mortalities up to 40%.

Subclinically infected wild fish, including Atlantic mackerel (S. scombrus), have been recovered

adjacent to Atlantic cod aquaculture in the Northeast Atlantic, since the 1980s (Ottem et al.,

2008). The role of these fish as reservoir hosts for farmed species is unclear (Colquhoun &

Duodu, 2011).

Clinical signs

Clinically infected fish may have darkened skin due to extensive haemorrhages of the dermal

musculature. Subclinical infection usually progresses with mild or no clinical signs (Nylund &

Ottem, 2006; Colquhoun & Duodu, 2011).

Pathogen survival

Francisella spp. may enter a VBNC state. Experimental data indicate they may survive for up to

30 days at 4 °C, and up to 16 days at 12 °C in marine and fresh waters, with longer survival in

sediment (Ottem et al., 2009; Colquhoun & Duodu, 2011). However, survival time in natural

waters is unknown and may be influenced by other factors (see section 6.1.2).


Transmission is horizontal and direct, through the water column, but the exact mechanisms are

unclear (Colquhoun & Duodu, 2011). Francisella spp. may be dispersed by fomites including

fish bait processing equipment (Arkush & Bartholomew, 2011).


Francisella spp. are considered emerging pathogens causing systemic, chronic, granulomatous

infections in wild and farmed fish (Colquhoun & Duodu, 2011).

Experimental data suggest the infectious dose in intracellular bacteria ranges from 10 cfu fish-1

(Nylund & Ottem, 2006; Colquhoun & Duodu, 2011), to 105 TCID50 ml-1 (Almendras et al.,

1997; Birbeck et al., 2004; Nylund et al., 2006). It is likely that a relatively high dose would be

necessary to induce infection with Francisella spp. (Johnston, 2008).

Pathogen entry occurs through the skin or viscera. Different strains target different organ

systems of the host. While clinical disease is associated with extensive necrotic granulomas of


the skin, mouth, gill and nasal tissue, the pathogenic mechanisms remain unclear (Arkush &

Bartholomew, 2011; Colquhoun & Duodu, 2011).

Inactivation

Francisella spp. remain viable after prolonged frozen storage (at -80 °C) (Colquhoun & Duodu,

2011). Related species survive for up to 75 days in frozen sheep muscle stored at -20 °C

(Johnston, 2008). Francisella spp. are unlikely therefore, to be affected by either short (168

hours) or medium-term (4 months) frozen storage (to -20 °C).

Francisella spp. are assumed to be denatured by ionising radiation (at 50 kGy) (DAFF, 2013).



In fish bait, Francisella spp. occur rarely in wild Atlantic mackerel (Scomber scombrus) with no

clinical signs. The prevalence estimates of 10% are based on a small sample size (n=9 fish) and

infection only occurs in fish taken immediately adjacent to Atlantic cod (G. morhua) fish farms

previously infected with Francisella spp. (Ottem et al., 2009). The significance of this infection

is unknown but is assumed to be minor (Colquhoun & Duodu, 2011).

Commercial fishing is a large-scale operation (FAO, 2014) and Atlantic mackerel represented

less than 3% of imported finfish bait in 2018 (Table 3). The likelihood of entry of Francisella

spp. through fish bait is therefore assessed to be so low as to be negligible.


As the entry assessment is negligible, Francisella spp. is assessed as not representing a risk in

the commodity. No further risk assessment is necessary.

Pseudomonas anguilliseptica



Pseudomonas anguilliseptica is a Gram-negative, rod-shaped motile bacterium, classified in

Family Pseudomonadaceae. Three main serotypes are recognised (I-III), with high phenotypical

homogeneity (Lopez-Romalde et al., 2003). Each serotype includes several isolates reported

from a wide range of marine and freshwater fish hosts and across geographical areas (Daly &

Akoi, 2011).

17.1.2 OIE status

Infection with P. anguilliseptica is not listed by the OIE (OIE, 2019a).


P. anguilliseptica has not been reported from New Zealand and is considered exotic (Duignan et

al., 2003). Infection with P. anguilliseptica is not a notifiable disease in New Zealand (Anon.,

2016).



P. anguilliseptica is not zoonotic (Haenen, 2017).

17.1.5 Epidemiology

P. anguilliseptica is a significant emerging pathogen, causing “Winter disease” in farmed fish

(Daly, 1999; Daly & Akoi, 2011; Wheeler, 2012; Blackwell, 2019).


P. anguilliseptica infection is reported from over 17 families of wild and farmed marine and

freshwater fish (Blackwell, 2019). It has a wide geographical distribution (Table 17).

The species associated with P. anguilliseptica in fish bait and their geographic/pathogen range

are given in Table 17.

Table 17. Fish bait species susceptible to Pseudomonas anguilliseptica



Atlantic Northeast Atlantic (Europe)

Notes 1 Host geographical range (from FishBase 2019) 2 Pathogen geographical range (from literature). P. anguilliseptica is reported from the North Atlantic and Mediterranean Sea

(Canada, Denmark, Egypt, Finland, France, Portugal, Spain and the United Kingdom), the Indo-Pacific and Northern Pacific Ocean (Australia, India, Japan, Malaysia and Chinese Taipei). It is also reported from deep sea microbial and hydroid communities in oceanic waters off Fiji (Berthe et al., 1995; Daly, 1999; Balboa et al., 2003; Romanenko et al., 2008; Wheeler, 2012; Andree et al., 2013; Mastan, 2013).

Prevalence in farmed fish varies from 12% (in salmonids), to 83% (in seabream, Sparus aurata)

(Wiklund & Lonnstrom, 1994; Fadel et al., 2018). Prevalence in wild Atlantic herring (C.

harengus) from coastal Finnish waters may reach 50% in epizootics (Lonnstrom et al., 1994)

(Table 17).

Infected wild fish may function as reservoir hosts (Daly & Akoi, 2011). P. anguilliseptica

infection has been transferred to farmed fish by herrings (Clupea spp.) (Clupeidae) when used as

aquaculture feed (Nash et al., 1987).


Potential hosts in New Zealand include jack mackerel (Trachurus sp.) (Carangidae), herrings

(Clupea spp.) (Clupeidae), salmonids (Oncorhynchus spp., Salmo spp., Salvelinus spp.), and

snapper (Pagrus auratus) (Sparidae) (Blackwell, 2019).

Pathogenicity

P. anguilliseptica is an opportunistic pathogen of wild fish (Lonnstrom et al., 1994). It causes

epizootic infections in a wide range of wild and farmed species (Daly & Akoi, 2011). Mortality

in marine aquaculture ranges from 2% (in Atlantic cod, Gadus morhua), to 50% (in Atlantic

salmon, Salmo salar) (Blackwell, 2019). However, pathogenicity is strongly influenced by water

quality and stocking levels (Nash et al.,1987) and varies between hosts (Muroga et al., 1975).

Experimental infection per os failed to initiate clinical disease in rainbow trout (Oncorhynchus

mykiss) (Lonnstrom et al., 1994), or in wild turbot (Scophthalmus (Psetta) maximus) (Magi et

al., 2009). In both studies, subsequent infection by intra-peritoneal (i.p.) injection resulted in

https://www.sciencedirect.com/topics/immunology-and-microbiology/sea-bream

https://www.sciencedirect.com/topics/immunology-and-microbiology/sparus-aurata


80% mortality in rainbow trout and 100% mortality in turbot, respectively. These data reflect the

opportunistic nature of P. anguilliseptica infection (Magi et al., 2009) and indicate P.

anguilliseptica is a secondary pathogen (requiring another agent to initiate infection) (Daly &

Akoi, 2011).

Strains of P. anguilliseptica associated with wild Atlantic herring (Clupea harengus) are of low

pathogenicity (Lonnstrom et al., 1994) where P. anguilliseptica may also be a secondary

pathogen, (Wiklund, 2016).

Clinical signs

External signs of infection may include exophthalmia, petechial haemorrhages, ulceration of the

ventral surface, mouth, anal region and fin bases, as well as eye lesions and punctured corneas

(Lonnstrom et al., 1994; Wiklund, 2016). The disease may also progress with no external clinical

signs (Mastan, 2013; Wiklund, 2016).

Pathogen survival

P. anguilliseptica survives in dead fish tissues (Sakr & El-Rhman, 2008). Experimental data

indicate P. anguilliseptica remains viable in marine and fresh water for extended periods (Ciric

et al., 2009), but its survival in natural waters is unknown (see section 6.1.2) Growth is limited

above 37 °C (Ferguson et al., 2004).

P. anguilliseptica survives in fish offal dumped in landfill and has been recovered from leachates

in groundwater (Tian et al., 2014). It is also present in deep-sea invertebrate communities

(Romanenko et al., 2008).


Disease transmission is horizontal, through the water column. Surviving fish are carriers (Fadel

et al., 2018). P. anguilliseptica may be dispersed through infected discharge water, biofilm and

fomites (Lonnstrom et al., 1994).


Infection occurs through cohabitation, or by consumption of infected fish, but the details of

infection are poorly understood (Austin, 2005; Austin & Austin, 2007). The infective dose varies

among isolates (Magi et al., 2009).

Disease may follow several pathways. Infection may be systemic, with necrotic vacuolation and

degeneration of liver, kidney and spleen tissues. This spreads to connective tissues, skeletal

musculature, the brain and cartilage, causing myolysis and pyogranulomatous inflammation

(Sakr & El-Rhman, 2008). Infection may also target the brain and nervous system, causing

corneal opacity, cephalic osteochondritis and meningitis. Surviving fish are carriers of disease

(Magi et al., 2009). Infection associated with the nervous system occurred at a prevalence of

33% in wild Atlantic herring (C. harengus) (Lonnstrom et al., 1994) and 40% in sea bream

(Sparus aurata) (Fadel et al., 2018).

Inactivation

P. anguilliseptica is resistant to freezing. It is unaffected by frozen storage (to -20 °C)

(Algammal et al., 2020) and remains viable after long-term storage (to -70 °C) (Hine &

MacDiarmid 1997; Magi et al. 2009). It is therefore assumed to be unaffected by either short

(168 hours), or medium-term (4 months) frozen storage (at -20 °C).


P. anguilliseptica is likely to be denatured by ionising radiation (at 50 kGy) (DAFF 2013).



P. anguilliseptica is associated with Atlantic herring (C. harengus) from Finnish waters at a

prevalence of up to 50%. Infected fish show no external signs of infection (Lonnstrom et al.,

1994). P. anguilliseptica is likely to remain viable in the commodity (Sakr & El Rhman, 2008;

Magi et al., 2009). As the host range and geographical range associated with fish bait are limited,

the likelihood of entry is assessed as low.


P. anguilliseptica survives in marine and fresh waters for extended periods (Ferguson et al.,

2004; Ciric et al., 2009). It remains viable in landfill and can be dispersed between waterways by

mechanical vectors such as piscivorous birds (Tian et al., 2014).

Exposure may occur through consumption of infected fish bait (Nash et al., 1987; Lonnstrom et

al., 1994), or by cohabitation through infected meltwater (Magi et al., 2009). It has a wide range

of potential hosts in New Zealand waters (Tubbs et al., 1997).

The strains associated with Atlantic herring are of low pathogenicity (Lonnstrom et al. 1994).

and P. anguilliseptica may be a secondary pathogen (Wiklund, 2016). It may be introduced into

farmed fish stocks through the use of fish bait as aquaculture feed, but establishment is strongly

influenced by water quality and stocking levels (Lopez-Romalde et al., 2003; Sakr & El Rhman,

2008; Magi et al., 2009; Jansson & Vennerstrom, 2014).

The likelihood of exposure and establishment of P. anguilliseptica through fish bait is assessed

to be so low as to be negligible.


As the likelihood of exposure and establishment is assessed to be negligible, P. anguilliseptica is

not assessed to be a risk in the commodity. No further risk assessment is necessary.

Streptococcus spp. (S. agalactiae serotype III: 283, S. iniae)



Streptococcus agalactiae (serotype III: 283) and S. iniae are gram-positive non-spore forming

bacteria of the Streptococcus species complex, classified within the family Streptococcaceae

(Gao et al., 2014; Mishra et al., 2018). S. agalactiae is classified in Lancefield’s serotype group

B Streptococcus (GBS).

Pathogenicity and host specificity vary widely among the members of this species complex

which include free-living, commensal and zoonotic organisms (Agnew & Barnes, 2007;

Rajendram et al., 2016; Zhou et al., 2011, 2018; Mishra et al., 2018; Barkham et al., 2019).

Many Streptococcus spp. are considered opportunistic pathogens (Bunch & Bejerano, 1997). S.


agalactiae III: 283 and S. iniae are regarded as primary fish pathogens, causing streptococcosis

disease in fish (Salati, 2011; Mishra et al., 2018).

18.1.2 OIE Status

Streptococcosis is not an OIE-listed disease (OIE, 2019a).


Serotypes of S. agalactiae are endemic in New Zealand, causing mastitis in dairy animals and

mild streptococcosis in wild and farmed fish (Hine & Diggles, 2005; Johnston, 2008; DermNet,

2014).

The pathogenic serotypes of S. agalactiae including serotype III: 283 reported from fish

(Barkham et al., 2018) have not been reported from New Zealand and are considered exotic

(Blackwell, 2019).

S. iniae has never been reported from New Zealand (Hine & Diggles, 2005; Johnston, 2008). It is

also considered to be exotic. Infection with Streptococcus spp. is not a notifiable disease in New

Zealand (Anon., 2016).


S. iniae and S. agalactiae III: 283 are zoonotic pathogens. S. iniae infection is mainly associated

with puncture wounds arising from the handling of fish. This generally causes mild bacteremic

cellulitis. In rare cases, however, infection progresses to endocarditis, meningitis, arthritis,

sepsis, pneumonia, osteomyelitis and toxic shock (Shoemaker et al., 2001; Baiano & Barnes,

2009).

S. agalactiae infection occurs through consumption of under-cooked fish. Infection may result in

meningitis and pneumonia (Agnew & Barnes, 2007; Baiano & Barnes, 2009).

Of the 2,824 hospitalised cases in Australia attributed to atypical Streptococcus spp. during

1999–2000 and in 2006–2007, 278 cases in 1999–2000, and 430 cases in 2006–2007, were due

to S. iniae, respectively. As S. iniae is not included in routine biochemical testing or clinical

databases, these values are likely to be under-estimates (Baiano & Barnes, 2009).

18.1.5 Epidemiology


S. agalactiae (serotype III: 283) (= S. difficule) and S. iniae occur in over 20 families of fish

(Blackwell, 2019). They are widely distributed (Table 18 and 19).

The species associated with S. agalactiae III: 283 in fish bait and their geographic/pathogen

range are given in Table 18 (Eldar et al., 1995; Evans et al., 2002). Streptococcus infection also

occurs in South American pilchard (Sardinops sagax) in Kenyan coastal waters (Odhiambo et

al., 2018). This is not considered further as the species of Streptococcus is not defined.


Table 18. Fish bait species susceptible to Streptococcus agalactiae III: 283



East, West Atlantic, Mediterranean East, West Pacific

Northeast Atlantic (Florida) Mediterranean (Europe)

Notes 1 Host geographical range (from FishBase 2019) 2 Pathogen geographical range (from literature). Streptococcus agalactiae III; 283 is reported from Europe (Belgium, Israel,

Iran, Italy, Kuwait, Spain), Asia (Hong Kong, Singapore, South Korea, Japan, Thailand, Viet Nam), North America (USA), South America (Brazil, Columbia, Costa Rica) and South Africa, (Buchanan et al. 2008; Zhou et al. 2011; Delannoy et al., 2013; Chen, 2019).

The species associated with S. iniae in fish bait and their geographic/host range are given in

Table 19 (Minami et al., 1979; Yasunaga et al., 1982; Sako, 1984; Johnston, 2008; Salati, 2011;

Kluzik & Woodford, 2016). The prevalence in flathead grey mullet (M. cephalus) may reach

50% during epizootics (Plumb et al., 1974).

Table 19. Fish bait species susceptible to Streptococcus iniae



Northwest Pacific, Japan, Korea

Northwest Pacific

Clupeidae South American pilchard (Sardinops sagax (= S. melanostictus)

West Pacific West Pacific Ocean


East, West Atlantic, Pacific. Mediterranean Sea

Mediterranean Sea (Israel)



Northwest Pacific

Notes 1 Host geographical range (from FishBase 2019) 2 Pathogen geographical range (from literature) S. iniae is globally distributed (Susanna et al., 2003).


A wide range of marine and freshwater fish host species are present in New Zealand (Tubbs et

al., 2007; Blackwell, 2019).

Pathogenicity

S. agalactiae III: 283 affects wild species of marine and freshwater fish where water

temperatures exceed 17 °C (Salati, 2011) and rarely affects farmed species where temperatures

are lower than 25 °C (Tavares et al., 2018). New Zealand coastal water temperatures vary from

17 °C in the north, to 12 °C in the south (Pinkerton et al., 2018). While S. agalactiae may

survive in New Zealand waters, it is likely to only affect farmed fish species in Northern New

Zealand waters.

S. iniae infection occurs in a wide range of tropical and temperate waters from 10 °C to 45 °C.

Optimal pathogenicity is associated with water temperature range from 35°C to 45 °C, but

infection occurred in farmed Australian barramundi (Lates calcarifer) at 10 °C (Bromage et al.

1999; Bromage & Owens, 2002).

Mortalities range from 10% to 60% in the coastal marine aquaculture of rainbow trout

(Oncorhynchus mykiss) (Salmonidae), turbot (Scophthalmus maximus) (Scophthalmidae),


gilthead sea bream (Sparus aurata) (Sparidae) in Europe and Australia (Carson et al., 1993;

Prieta et al., 1993, cited in Musquiz et al., 1999; Toranzo et al., 2009). Mortality is strongly

influenced by water quality and stocking levels.

Clinical signs

External signs of infection may include exophthalmia, distended abdomen, haemorrhages in the

eyes, fin bases and opercula, darkening of the skin and spine displacement (Amal & Saad, 2011).

Infection may also progress with no external signs (Yanong & Francis-Floyd, 2002; Salati,

2011).

Pathogen survival

S. iniae survives in marine waters for up to 7 days (Bromage et al., 1999). Survival in natural

waters is unknown (see section 6.1.2).


Transmission is horizontal and direct, either through consumption of infected fish tissues, or as

opportunistic infection, following abrasion and skin damage (Yasunaga, 1982; Shoemaker et al.,

2000). Subclinically infected or surviving fish may act as carriers of infection (Baiano & Barnes,

2009).


Infectivity is maintained for long periods in the aquatic environment by sequential subclinical

infection (Kitao et al., 1979; Nguyen et al., 2002). S. iniae remained present in water and

sediment samples taken from a bastard halibut (Paralichthys olivaceus) farm for two years

following initial infection (Bromage et al., 1999).

Infection is internalised by invasion and survival within macrophage cells (Yoshida et al., 1996;

Bromage et al., 1999; Baiano & Barnes, 2009). Experimental infection per os indicate the

infectious dose is low, ranging from 1 × 103 cfu ml-1 (in for barramundi, Lates calcarifer), to 2 ×

107 cfu ml-1 (in tilapia, Oreochromis sp.), after 1-minute exposure (Bromage & Owens, 2002;

Shoemaker et al., 2000).

Acute infection mainly occurs in younger fish, but all ages may be infected. Disease progresses

from the skin to systemic septicaemia of the spleen, liver, heart and kidney tissues, eyes, brain

and nervous system (Amal & Saad, 2011; Salati, 2011). Chronic infection may develop into a

carrier state in older fish (Baiano & Barnes, 2009; Salati, 2011).

Wild fish may be reservoirs of infection for farmed species. Streptococcus sp. infection was

transferred from wild Klunzinger’s mullet (Liza klunzingeri) to gilthead seabream (Sparus

aurata) farmed in Kuwait (Zlotkin et al.,1998; Evans et al., 2002; Salati, 2011).

Levels of S. iniae in infected host tissues may reach 109 cfu g-1 in spleen, kidney and brain tissue

and 109 cfu mL-1 in blood (Nguyen et al., 2002; Shutou et al., 2007). Infection results in lowered

productivity, while the presence of blood-filled lesions in fish tissues renders the product

commercially unmarketable (Muzquiz et al., 1990; Salati, 2011).

Inactivation

Streptococcus spp. remain viable in fish tissues after short term frozen storage (at -70 °C for 180

days) and long-term frozen storage (at -20 °C for up to 9 months) (Evans et al., 2004; Hine &

Digges, 2005; Diggles, 2011). Streptococcus spp. remain viable in frozen sardine (S. sagax)


where used as feed for yellowtail (Seriola quinqueradiata) aquaculture in Japan (Yasunaga,

1982) and are likely to remain infective in frozen fish bait. It is assumed therefore, that

Streptococcus spp. would be unaffected by either short term (168 hours), or medium-term (4

months) frozen storage (at -20 °C).

Streptococcus spp. are likely to be denatured by ionising radiation (at 50 kGy) (DAFF, 2013).



S. agalactiae III: 283 and S. iniae are associated with fish bait (Table 18 and Table 19). Infected

fish may show no external signs of infection (Evans et al., 2011). Prevalence in wild fish bait is

unknown, but this may reach 50% following epizootics (Plumb et al., 1974; Lafferty et al.,

2015). Streptococcus spp. are unaffected by frozen storage (Evans et al., 2004; Hine & Diggles,

2005, Diggles, 2011) and may be present in the meltwater associated with fish bait (Bromage et

al., 1999).

The likelihood of entry is assessed as medium.


Experimental data indicate Streptococcus spp. survive for up to 7 days in the aquatic

environment (Bromage et al., 1999) within the temperature range of New Zealand coastal marine

waters (Pinkerton et al., 2018).

The pathogen load in tissues or in meltwater is high (Nguyen et al., 2001; Shutou et al., 2007),

while the infectious dose is low (Bromage & Owens 2002). Infectivity can be maintained in the

environment by sequential subclinical infection of a range of carrier hosts (Bromage et al., 1999;

Nguyen et al., 2002).

Infection has occurred through use of frozen fish bait (S. sagax) as feed in aquaculture

(Yasunaga, 1982). Infection can also be transferred from wild to farmed fish (Plumb et al., 1974;

Zlotkin et al., 1998; Salati, 2011). S. iniae was introduced into the USA in 1994, demonstrating

its ability to translocate (Perera et al., 1994; Johnston, 2008).

The likelihood of exposure and establishment of S. agalactiae III: 283 and S. iniae is assessed as

medium.



Streptococcosis (including infection with S. iniae or S. agalactiae) is not an OIE-listed disease

(OIE, 2019a), so establishment would not result in loss of trade access for New Zealand.

Establishment may result in lowered productivity and increased mortality, as well as product loss

resulting from tissue contamination (Muzquiz et al., 1990; Salati, 2011). Streptococcosis causes

economic losses estimated at US$100 million annually in the combined aquaculture of Australia,

Israel, Japan and the USA (Shoemaker et al., 2001).

S. agalactiae III; 283 and S. iniae have a wide host range (Salati, 2011). Their establishment may

affect the aquaculture of Chinook salmon (Oncorhynchus tshawytscha) (Salmonidae), with


exports valued at $77 million in 2018 (Aquaculture New Zealand, 2019). It may also affect the

developing aquaculture of yellowtail kingfish (Seriola lalandi) (Carangidae) and New Zealand

snapper (silver seabream) (Pagrus auratus) (Sparidae) (Plant & Food, 2016, 2017; NIWA,

2017a).

Establishment may reduce productivity of important coastal inshore fisheries (MPI, 2018a,

2018b, 2018c). Wild fish are reservoirs of infection for farmed species, including gilthead sea

bream (Sparus auratus) (Sparidae) (Zlotkin et al., 1998). The New Zealand snapper (P. auratus)

export fishery was valued at $33 million in 2018 (Seafood New Zealand, 2019).

Establishment may affect farmed freshwater species, including grass carp (Ctenopharyngodon

idella) and silver carp (Hypophthalmichthys molitrix) used for weed control (Clayton & Wells,

1999). Introduced cyprinids such as the European (red fin) perch (Perca fluviatilis) may function

as reservoir hosts.

Social consequences

Exotic S. iniae and S. agalactiae strains are emerging zoonotic pathogens (Salati 2011). Most

cases are minor, but infection may be serious in older patients (> 50 years) or those with

compromised immune systems (Shoemaker, 2001; Agnew & Barnes, 2007; Baiano & Barnes,

2009; Salati, 2011).

Establishment may affect recreational trout fishing (brown trout, Salmo trutta) and rainbow trout

(Oncorhynchus mykiss) (Fish & Game, 2014). The Lake Taupo trout fishery alone was valued

annually at NZ$ 70–80 million (Marsh & Mkwra, 2013).

The effects of pathogens on wild fisheries are poorly known (Lafferty et al., 2015), but

establishment may affect other species of importance for customary and recreational fishing in

marine and fresh waters (Wynn-Jones et al., 2014, 2019). Recreational fishing is the 5th most

popular recreational activity for adults, involving 700,000 marine fishing trips annually, with a

contribution of NZ$ 638 million to the GDP in 2015 (Gross domestic product) (Southwick et al.

2018). The snapper and trevally recreational fisheries contributed NZ$ 402 million and NZ$ 71

million respectively to the GDP in 2015 (Southwick et al., 2018).


Streptococcus spp. may be introduced through frozen fish bait (Yasunaga, 1982). S. agalactiae

III: 283 and S. iniae have a wide host range. The pathogen load is high, and the infective dose is

low (Bromage & Owens, 2002). Infection has established through sequential subclinical

infection of susceptible wild fish, leading to Streptococcus spp. becoming endemic (Kitao et al.,

1979; Nguyen et al., 2002).

Epizootic infection is reported in flathead grey mullet (Mugil cephalus) (Plumb et al., 1974).

Establishment may cause lowered productivity in a wide range of marine and freshwater fish

species considered stressed at current levels of fishing activity (MPI, 2018a, 2018b, 2018c).

Inshore fish of families Clupeidae, Engraulidae and Mugilidae also represent a significant food

source for other fish, seabirds and marine mammals and a major link between trophic levels

(Dunn et al., 2012).

The consequences of the introduction of S. agalactiae III: 283 and S. iniae are assessed to be

medium.



As the entry, establishment and consequence assessments are medium, the risk estimates for S.

agalactiae III: 283 and S. iniae are assessed to be medium. Under the procedures followed in this

risk analysis, risk management measures may be justified.


S. agalactiae III: 283 and S. iniae have been assessed to be risks in fish bait. Infection with

Streptococcus spp. is not an OIE-listed disease, so the OIE Aquatic Code (OIE, 2019a) provides

no specific guidance or processing requirements that would ensure the destruction of these

pathogens.

S. agalactiae III: 283 and S. iniae are associated with the commodity (Table 18 and Table 19).

Therefore, restriction to non-susceptible species in the commodity should substantially reduce

the risk associated with these pathogens and be a viable risk management option.

S. agalactiae III: 283 and S. iniae have a wide geographic range (Table 18 and Table 19). Their

occurrence is likely to be low in other geographic regions. Restriction through the BNZ Country

Approval Procedures to regions where these pathogens have not been reported should

substantially reduce the risk associated with these pathogens and be a viable risk management

option.

The presence of S. agalactiae III: 283 and S. iniae in a shipment of the commodity may be

verified by batch testing using standard sampling procedures (Corsin et al., 2009; OIE, 2019a).

Standard genomic tests for S. iniae are not available. Recognised diagnostic procedures are

provided in Weinstein et al. (1997), Pereira et al. (2013) and Buller (2014). Verification of a

negative test from a shipment should substantially reduce the risk associated with these

pathogens and be a viable risk management option.

Streptococcus spp. are unaffected by frozen storage (at -20 °C) for up to 9 months (Evans et al.,

2004; Hine & Digges, 2005; Diggles, 2011). Therefore, frozen storage, for either 168 hours or 4

months, is not a viable risk management option.

Streptococcus spp. are assumed to be denatured by ionising radiation (at 50 kGy) (DAFF 2013).

This irradiation treatment should substantially reduce the risk associated with these pathogens

and be a viable risk management option.



(see section 6.2.1), one or more of the following risk management options would reduce the risk

specific to S. agalactiae III: 283 and S. iniae an acceptable level.

Option 1

• Competent Authority attestation that the commodity does not include Japanese jack mackerel

(Trachurus japonicus) (Carangidae); South American pilchard (Sardinops sagax)

(Clupeidae); flathead grey mullet (Mugil cephalus) (Mugilidae); and chub mackerel

(Scomber japonicus) (Scombridae) should substantially reduce the S. agalactiae III: 283 and

S. iniae risk, so the commodity could be imported without further restrictions.


Option 2

• Acceptance of region/country freedom through the BNZ Country Approvals Procedures

should substantially reduce the S. agalactiae III: 283 and S. iniae risk, so the commodity

could be imported without further restrictions.

Option 3

• A negative test from a shipment of the commodity, following approved sampling and

diagnostic procedures through the BNZ Country Approvals Procedures should substantially

reduce the S. agalactiae III: 283 and S. iniae risk, so the commodity could be imported

without further restrictions.

Option 4

• Treatment with ionising radiation (at 50 kGy) of the frozen commodity should substantially

reduce the S. agalactiae III: 283 and S. iniae risk, so the commodity could be imported

without further restrictions.

Myxozoan pathogens


19.1.1 Aetiological agents

Myxozoan (= Myxosporean) fish pathogens are highly specialised multicellular obligate

parasites. They are classified in Class Myxozoa of the Phylum Cnidaria (WoRMS, 2019). The

current classification within the Myxozoa is based on a complex and detailed description of

spore anatomy (Lom & Dykova, 1992). This classification has become controversial. Genomic

studies have shown the Class Myxozoa to be polyphyletic, with several major clades and species

complexes (e.g., Chloromyxum, Kudoa, Myxobolus spp.) (Okamura et al., 2011; Fiala et al.,

2015; Okamura & Gruhl, 2015; Chang et al., 2015; Okamura et al., 2018). Some, including the

Kudoa thyrsites species complex, have a wide host range (Feist & Longshaw, 2006; Fiala et al.,

2015; Vidal et al., 2017).

Myxozoans generally cause little apparent harm to their hosts (Feist & Longshaw, 2006).

However, tissue-inhabiting myxozoans cause extensive post-mortem myoliquefaction,

commonly resulting in post-harvest rejection of the commercial product. Infected wild fish may

become reservoir hosts for farmed species (Langdon et al., 1992; Hine & Jones, 1994; Moran et

al., 1999). Myxozoans have been previously identified as risk organisms in New Zealand fish

(Johnston, 2008; Blackwell, 2019).

This risk analysis is restricted to exotic myxozoans associated with post-mortem

myoliquefaction, where the host range includes fish species of economic, social or

environmental importance to New Zealand. These are reviewed together in this risk analysis, as

the host range is common for several identified myxozoan hazards.

19.1.2 OIE status

Diseases associated with myxozoan pathogens are not listed by the OIE (OIE, 2019a).



Myxozoans are reported from New Zealand fish. These are generally host-specific and are

primarily associated with coelomic spaces (bile duct, gall bladder, urinary bladder) (Hine. 1977;

Hewitt & Little, 1972; Boustead, 1982; McArthur & Sengupta, 1982; Landsber & Lom, 1991;

Hine et al., 2000; Diggles et al., 2002; Bingham, 2009; Lane et al., 2014). They are likely to

have a minimal effect on their host (Lom & Dykova, 1992).

Myxozoan species including Kudoa hexapunctata, K. prunusi, K. shiomitsui and K. yasunagi

(Table 6) are reported from highly migratory pelagic and oceanic tuna species. These tuna

species are widely reported from the Pacific Ocean, including New Zealand (Zhang et al., 2010;

Meng et al., 2011; Kasai et al., 2017). It is considered likely that the myxozoans associated with

these highly migratory species are present, but not reported in New Zealand waters. They are not

considered further.

Kudoa thyrsites has a wide geographical distribution and a wide host range. It has not yet been

reported from New Zealand waters (Hine & Jones, 1994; Stone et al., 1997; Hine et al., 2000;

Blackwell, 2019). It has not been reported from passive testing programmes (M. Bestbier, BNZ,

pers. comm., 2020). It is assumed that K. thyrsites is absent from New Zealand waters.


Myxozoans are minor zoonotic pathogens (Boreham et al., 1998; Feist & Longshaw, 2006;

Iwashita et al., 2013; Kasai et al., 2015). Myxobolus spp. have caused zoonotic infection

following consumption of previously frozen fish fillets (Boreham et al., 1998; Ohnishi et al.,

2016).

19.1.5 Epidemiology


Blackwell (2019) identified several myxozoan species of concern likely to be associated with

eviscerated fish. The present risk analysis considers a wider range of myxozoans likely to be

present in whole (uneviscerated) fish bait species (Table 5). Most myxozoans are host-specific,

at least to the family level, with high tissue or organ-specificity (Lom & Dykova, 1995, 2006;

Molnar & Eszterbauer, 2015). They are of low or low-moderate pathogenicity and are not

regarded as significant fish pathogens in wild fish stocks (Langdon et al., 1992; Lom & Dykova,

1992, 2006; Diamant et al., 1994; Feist & Longshaw, 2006; Lom & Dykova, 2006; Gomez et al.,

2014). These have not been considered further.

Myxozoan species from genera Enteromyxum and Kudoa are associated with post-mortem

myoliquefaction and have a wide host range. E. leei occurs in 46 host species from 6 families. K.

clupeidae occurs in 13 host species from 2 families. K. iwatai occurs in 19 host species from 6

families. K. nova (= K. quadratum) occurs in 20 host species from 2 families, while the K.

thyrsites species complex occurs in 40 host species from 5 families (Lom & Dykova, 2006;

Blackwell, 2019).

The species associated with E. leei, K. clupeidae, K. iwatai, K. nova and K. thyrsites in fish bait

and their geographic/host range are given in Table 20 (Hahn, 1917; Meglitsch, 1947; Heckman

& Jensen, 1978; Kpatcha et al., 1999; Egusa & Shiomitsu, 1983; Lom & Dykova, 1992; Munday

et al., 1998; Reimschuessel et al., 2003; Longshaw et al., 2004; Campbell, 2005; Diamant et al.,

2005; Whipps & Kent, 2006; Langdon, 2007; Montero et al., 2007; Matsukane et al., 2011;


Henning et al., 2013; Mackenzie & Kalavati, 2014; Fall et al., 2015; Kasai et al., 2015; Levsen,

2015; Ovcharenko, 2016; Yurakhno, 2017; Dezfuli et al., 2020).

The reported geographical range of these myxozoans may underestimate their true distribution,

as some regions, such as the North Atlantic, have been more thoroughly investigated than others

(Mackenzie & Kalavati, 2014; Fiala et al., 2015).

Myxozoan prevalence varies widely but is likely to be low in wild fish stocks. Few data are

available, and the sample sizes are often small (Henning et al., 2013; Alama-Bermejo et al.,

2013). Reported prevalence of E. leei in the thinlip grey mullet (C. ramada) was estimated at

83% (from n= 43 fish) (Dezlui et al., 2020). K. thyrsites prevalence varies with season and with

host size in fish bait species. In the South American pilchard (S. sagax), prevalence varies from

17% during spring, to 91% in autumn (Reed et al., 2012); Henning et al., 2019). In Atlantic

mackerel (S. scombrus), overall prevalence was 0.8% (from 1,475 fish), but the prevalence was

higher (8.9%) in older, heavier (> 600 g) fish (Levsen et al., 2008).


Table 20. Fish species susceptible to exotic myxozoan pathogens Enteromyxum leei, Kudoa clupeidae, K. iwatai. K. nova. K. thyrsites

Myxozoa Host family

Host species Host geographical range1

Pathogen range2

Enteromyxum leei

Mugilidae Flathead grey mullet (Mugil cephalus), golden grey mullet (Chelon auratus), thinlip grey mullet (C. ramada)

East Atlantic, West Atlantic, Mediterranean, West Pacific Ocean


Kudoa clupeidae


Northwest Atlantic, Southwest Atlantic

Northeast Pacific (California), Southwest Atlantic

Kudoa iwatai Carangidae Japanese jack mackerel (Trachurus japonicus)

North Pacific Japan, Korea, Red Sea


Widespread, Mediterranean

Mediterranean, Japan

Kudoa nova Carangidae European horse mackerel (Trachurus trachurus)

North Atlantic, Northeast Atlantic, Black Sea, Mediterranean, Pacific

Pacific (Western Australia), Black Sea, North, Northeast Atlantic, West Atlantic, Mediterranean

Scombridae Bigeye tuna (Thunnus obesus), little tunny (Euthynnus alleteratus)

North Sea, Northeast, Northwest Atlantic, Mediterranean

North Sea, Northwest Atlantic, Mediterranean

Kudoa thyrsites

Clupeidae Bali sardinella (Sardinella lemuru), South American pilchard (Sardinops sagax)

Central, Northwest Pacific Northeast, Southeast Atlantic Southeast, Northeast Pacific, Mediterranean

Northwest, Eastern Pacific (Alaska, Australia, Japan, East, Northeast Northwest Atlantic, Mediterranean (Chile, England, South Africa)

Engraulidae Californian anchovy (E. mordax)



Scombridae European horse mackerel (Scomber scombrus), Chub mackerel (S. japonicus)

Northeast, Southeast Atlantic, Pacific (Australia, Canada, Chile, Japan)

Northeast, Southeast Atlantic (South Africa), Pacific (Australia, Japan, Southeast Pacific (Chile), Northeast Pacific (Canada, Alaska)

Notes 1 Host geographical range (from FishBase 2019) 2 Pathogen geographical range (from literature)


A wide range of potential hosts exist in New Zealand (Blackwell, 2019), but the following

commercially farmed fish species are relevant to this chapter: brown trout (Salmo trutta),

Chinook salmon (Oncorhynchus tshawytscha), rainbow trout (O. mykiss) (Salmonidae);

yellowtail kingfish (Seriola lalandi) (Carangidae); snapper (Pagrus auratus) (Sparidae); hapuku

(Polyprion oxygeneios) (Polyprionidae); turbot (Colistium nudipinnis) (Pleuronectidae). The host

range of the myxozoans E. leei, K. clupeidae, K. nova, K. iwatai and K. thyrsites includes finfish

species commercially farmed in New Zealand.


Pathogenicity

Myxozoans have little apparent effect on their fish hosts, even at high levels of infection (Lom &

Dykova, 1992, 2006; Alvarez-Pellitero & Sitja-Bobadilla, 1993; Feist & Longshaw, 2006;

Gomez et al., 2014; Diggles, 2011; Blackwell, 2019). Myxozoan species clades preferentially

infect specific tissue types, but the severity of infection varies among host species. (Egusa &

Nakajima, 1980; Burger et al., 2008).

Clinical signs

Infection in wild fish stocks usually progresses with few or no external signs of infection (Lom

& Dykova, 2006). Chronic infection is usually associated with aquaculture, where stressors may

include high temperatures, overcrowding and poor water quality. Heavily infected fish may

become emaciated, with visible cysts, lesions and necrosis of the skin or fins. These fish may

show poor weight gain and increased mortality (Lom & Dykova, 2006; Gomez et al., 2014;

Stilwell & Yanong, 2018).

Pathogen survival

Little is known about myxozoan survival in marine waters. Experimental data from fresh water

myxozoans such as M. cerebralis suggest infective actinospores may remain viable for up to 7

days in water (Hoffman, 1990; Eszterbauer et al., 2001), or up to 12 months in sediments

(Nehring et al., 2015). However, experimental data may not reflect survival in natural waters

(see section 6.1.2).


The life cycle of marine myxozoans is complex (Chang et al., 2015; Fiala et al., 2015; Okamura

et al., 2015; Naldoni et al., 2019). Life cycle details are only known for 50 freshwater

myxozoans out of the 2,200 species associated with fish. These have an indirect life cycle,

usually involving a bryozoan or annelid intermediate host (Lom & Dykova, 2006).

Little is known about the life cycle details, viability and host range of marine myxozoans (Feist

& Longshaw, 2006; Lom & Dykova, 2006). These are assumed to also have an indirect life

cycle. The intermediate host may be a bryozoan or annelid (Lom & Dykova, 2006), or an

undescribed zooplankton species (Sindermann, 1990). Myxozoans may persist and replicate

through vertical transmission in their intermediate hosts (Feist & Longshaw, 2006; Hallett et al.,

2015; Nehring et al., 2015).

Some marine myxozoans, such as E. leei, have a direct fish-fish life cycle. Infection occurs by

cohabitation, or through consumption of infected tissues, but the details of infection are poorly

understood (Diamant, 1997; Redonto et al., 2002; Yasuda et al., 2002; Diamant et al., 2006;

Lom & Dykova, 2006).

Disease transmission in Kudoa spp. may also be direct, through an extra-sporogonic stage, or

stages in the life cycle. Experimental studies have shown infection with K. thyrsites can be

initiated in naïve Atlantic salmon (Salmo salar) by injection of blood from infected fish

containing these extra-sporogonic stages (Moran et al., 1999; Young & Jones, 2005). While the

significance of these life cycle stages in disease transmission is uncertain (Young & Jones,

2005), K. thyrsites could also be transmitted directly from fish to fish.

Myxozoans survive passage through avian digestive systems. The infective stages may be

dispersed in avian faeces between waterways by piscivorous birds (El-Matbouli & Hoffmann,


1991). Host specificity to the intermediate host is usually low (Gomez et al., 2014) and

alternative invertebrate hosts may be present in New Zealand waters.

Transmission of spores from the fish host may be delayed until the death or predation of the host.

Therefore, the time required for an exotic myxozoan to establish in a new region may be in the

order of decades. The process of establishment is complex and dependent on multiple factors.

These include temperature, salinity, pH, the rate of predation of the host fish relative to the

developmental stage of the pathogen, as well as on the distribution biology of both the

intermediate and final host (Hallett et al., 2015; Jones et al., 2015).


Infective actinospores contact the fish host by invading cells of the skin, fins, buccal cavity,

digestive tract or gills (Lom & Dykova, 2006). Infection may remain localised in the epithelial

tissues, or become systemic, affecting multiple organs (Gomez et al., 2014).

Infection may be coelozoic (inhabiting body spaces) or histozoic infection (inhabiting organs or

muscle tissues). Chronic infection is usually lifelong. Infection generally progresses with little

apparent effect upon the host, where the infection rate is low (Lom & Dykova, 2009).

At higher rates of infection, tissue necrosis, cell detachment and vacuolisation may occur. This

results in chronic enteritis, anorexia, and cachexia (Gomez et al., 2014). Infection of the gill

tissues causes epithelial damage and cyst formation. The resulting hypertrophy and fusion of gill

lamellae may significantly reduce gill function (Lom & Dykova, 2006).

Where infection occurs in the musculature, open skin lesions develop that allow myxospores to

be released to the environment. When these lesions heal, an enclosed pseudocyst is formed. The

processes of pseudocyst formation are poorly understood (Sindermann, 1990; Rothwell et al.,

1997; Gomez et al., 2014). These pseudocysts decompose post-mortem, causing tissue

myoliquefaction, tissue spoilage and economic loss of the commercial fish product (Feist &

Longshaw, 2006). The loss rates from product rejection due to tissue myoliquefaction may be

significant. It is estimated that 4% to 7% of Canadian farmed Atlantic salmon (Salmo salar)

product is rejected due to myoliquefaction (Marshall et al., 2015).

Myxozoans present in deeper tissues of the brain, cartilage, or somatic muscle tissues also cause

little apparent harm to their host. These are released by decomposition, following the death of the

host (Diamant, 1997; Feist & Longshaw, 2006).

Inactivation

Myxospores are relatively resistant to frozen storage in fish tissues. Most inactivation data are

available for freshwater myxozoans. Spore viability of M. cerebralis is reduced following

medium-term (3 months) frozen storage (to -20 °C or below) (Boreham et al., 1998; El Matbouli

& Hoffmann 1991; Yurakhno, 2017). Infectivity is significantly reduced after 5 months storage

(at -20 °C), where only 2 of the 15 samples (13%) remained viable (Hoffman & Putz, 2011). M.

cerebralis spores in salmon tissue stored at -20 °C or -80°C were rendered non-viable after 2

months storage, while spores held at 4 °C and higher remained infective for the Tubifex tubifex

intermediate host (Hedrick et al., 2008).

For marine myxozoans, a 20% reduction in viability of Kudoa nova occurred after freezing

(to -20 °C) (Yurakhno, 2017). Kudoa septempunctata spores in olive flounder (Paralichthys

olivaceus) were rendered non-viable by frozen storage (to -20 °C for 4 hours) (Anon., 2020b).


Marine myxozoans are likely to be unaffected by short-term (168 hours) frozen storage

(at -20 °C), but substantially, or completely denatured by medium-term frozen storage (for at

least 4 months) (El Matbouli & Hoffmann, 1991; Boreham et al., 1998; Lom & Dykova, 2006;

Hedrick et al., 2008; Reed et al., 2012; Ohishi et al., 2016; Henning et al., 2019; Anon., 2020b).

Myxozoan prevalence is generally low in wild fish stocks. Where pathogen load is high,

sufficient spores may remain viable to initiate infection after short-medium term frozen storage

(Hedrick et al., 2008; Levsen et al., 2008; Bartholomew & Kerans, 2015; Henning et al., 2019).

Myxozoans are likely to be denatured by ionising radiation (at 50 kGy) (DAFF, 2013).



The myxozoans E. leei, K. clupeidae, K. iwatai, K. nova and K. thyrsites are associated with fish

bait, including species that are imported in large quantities (Table 3). Infected fish typically

show no external signs of infection (Lom & Dykova, 2006).

The prevalence in fish bait is highly variable depending on host species, fish size and season of

capture (Lom & Dykova, 2006; Levsen et al., 2008; Reed et al., 2012; Henning et al., 2019).

Myxozoans are likely to remain viable in the commodity. As myxozoan prevalence is highly

variable, the likelihood of entry is assessed as low.


The life cycle of marine myxozoans is poorly understood (Lom & Dykova, 2006). Some, such as

E. leei have a direct life cycle, with fish-fish transmission and a wide host range (Diamant et al.,

1997, 2006; Feist & Longshaw, 2006). Others, including Kudoa spp. may also have a direct life

cycle (Moran et al., 1999; Young & Jones, 2005). These may also use an intermediate host such

as an annelid, bryozoan, polychaete or other unknown zooplankton species (Sindermann, 1990;

Bartholomew & Kerans, 2015).

Little data are available on viability of the infective stages, infective dose, or the specificity of

the final, or intermediate hosts of Enteromyxum leei or Kudoa spp. (Feist & Longshaw,2006). It

is possible that alternative host species may be present in New Zealand waters (Blackwell,

2019). Myxozoans can also persist through vertical transmission in their intermediate hosts (Feist

& Longshaw, 2006; Hallett et al., 2015; Nehring et al., 2015). Myxozoans may be dispersed by

piscivorous birds between waterways through faeces or in discarded fish tissues (El-Matbouli &

Hoffmann, 1991).

It is reasonable to assume that infected fish bait used in inshore waters may infect coastal inshore

finfish, through a suitable benthic invertebrate host (if required), in quantities and duration

sufficient to establish an infection. However, the factors influencing establishment success of

myxozoans are complex. As infection is rarely fatal to the finfish host, successful establishment

in wild stocks may be unapparent, even over a time scale of decades (Hallett et al., 2015; Jones

et al., 2015).

Myxozoan establishment may only be apparent in susceptible marine farmed species, where

infection has become established in a susceptible wild fish host, that could act as a reservoir of

infection (Feist & Longshaw, 2006; Montero et al., 2007; Bartholomew & Kerans, 2015).


The likelihood of exposure and establishment of E. leei, K. clupeidae, K. iwatai, K. nova and K.

thyrsites through fish bait is extremely uncertain but is assumed to be low, or very low.


Economic consequences.

Disease associated with E. leei, K. clupeidae, K. iwatai, K. nova and K. thyrsites is not listed by

the OIE (OIE, 2019a), so establishment would not result in loss of general trade access for New

Zealand. However, the presence of K. thyrsites in Chinook salmon exports may result in loss of

access to some North American markets (M. Bestbier, BNZ, pers. comm., 2020).

E. leei, K. clupeidae, K. iwatai, K. nova and K. thyrsites are likely to cause little apparent effect

in wild fish stocks at low levels of prevalence (Lom &Dykova, 2009). However, they may cause

economic losses through reduced productivity and increased mortality in aquaculture (Jones et

al., 2015). This would have significant consequence for New Zealand salmonid aquaculture,

including brown trout (Salmo trutta), Chinook salmon (Oncorhynchus tshawytscha) and rainbow

trout (Oncorhynchus mykiss). Losses in Canadian salmonid aquaculture resulting from Kudoa

thyrsites were estimated at CAD$ 6 million in 2015 (Braden et al., 2017). Chinook salmon

(Oncorhynchus tshawytscha) exports were valued at $77 million in 2018 (Aquaculture New

Zealand, 2019).

E. leei, K. clupeidae, K. iwatai, K. nova and K. thyrsites may also affect aquaculture of non-

salmonid species including yellowtail kingfish (Seriola lalandi) (Carangidae), snapper (Pagrus

auratus) (Sparidae), hapuku (Polyprion oxygeneios) (Polyprionidae) and turbot (Colistium

nudipinnis) (Pleuronectidae) (Moran et al., 1999a; Rigos et al., 1999; Montero et al., 2007;

Rigos & Katharios, 2010; Berger & Adlard, 2011; ; Ovcharenko, 2016; NIWA, 2017a, 2017b,

2017c; Aquaculture New Zealand, 2019). For example, the snapper export fishery was valued at

$33 million in 2018 (Seafood New Zealand, 2019).

Other Kudoa spp. associated with mugilid and scombroid fish species have caused significant

economic losses in Asian, Australian and European aquaculture (Campbell, 2005; Lom &

Dykova, 2006; Montero et al., 2007; Yurakhno & Ovcharenko, 2014; Bartholomew & Kerans,

2015; Jones et al., 2015). These host species are not farmed in New Zealand, so the

consequences of establishment associated with these fish families are assumed to be negligible.

Social consequences

Myxozoans are minor zoonotic pathogens, causing gastroenteritis following consumption of

previously frozen fish fillets (Boreham et al., 1998).

The social consequences for customary and recreational fishing in New Zealand due to the

establishment of E. leei, K. clupeidae, K. iwatai, K. nova and K. thyrsites are unknown, but likely

to be very low, at low rates of prevalence in wild fish stocks.


Evaluation of the consequences of myxozoan establishment in wild fish stocks is complex

(Lafferty et al., 2015). Myxozoans are of low pathogenicity (Egusa & Nakajima, 1980; Diamant

et al., 1994; Burger et al., 2008; Lom & Dykova, 2013; Gomez et al., 2014; Hallett et al., 2015;

Jones et al., 2015). The environmental consequences are unknown, but likely to be very low.

The consequences of establishment of E. leei, K. clupeidae, K. iwatai, K. nova and K. thyrsites

are uncertain, but assessed to be low.



The entry, exposure and consequence assessments for E. leei, K. clupeidae, K. iwatai, K. nova

and K. thyrsites are assessed to be low. The risk estimate is therefore assessed to be low, but with

high levels of uncertainty. Under the procedures followed in this risk analysis, risk management

measures may be justified.


E. leei, K. clupeidae, K. iwatai, K. nova and K. thyrsites have been assessed to be a risk in the

commodity. Infection with myxozoan pathogens is not an OIE-notifiable disease, so the OIE

Aquatic Code (OIE, 2019a) provides no specific guidance or processing requirements that would

ensure the destruction of these pathogens.

Myxozoans have been identified as risk organisms in previous IRAs in New Zealand

(MacDiarmid, 1993; Stone et al., 1997; Diggles, 2011; Blackwell, 2019) and in Australia (Kahn

et al., 1999). They have been identified as risk organisms in risk analyses for emerging

aquaculture species (Nowak et al., 2003; Hutson et al., 2007; Sanchez-Garcia et al., 2014;

Okamura et al., 2015).

Myxozoans E. leei, K. clupeidae, K. iwatai, K. nova and K. thyrsites are associated with fish bait

(Table 21). These species are likely to be substantially or completely denatured by medium-term

frozen storage (to -20 °C for at least 4 months (17 weeks)) (El Matbouli & Hoffmann, 1991;

Boreham et al., 1998; Lom & Dykova, 2006; Hedrick et al., 2008; Reed et al., 2012; Ohishi et

al., 2016; Henning et al., 2019; Anon., 2020b). An extended period of frozen storage is

consistent with current fish bait industry practice. Adoption of this general risk management

option should substantially reduce the risk associated with these myxozoans and be a viable risk

management option.


If the general risk management measures proposed for all imported fish bait (section 6.2.1) are

adopted, then no pathogen-specific risk management measures would be necessary to reduce the

risk associated with E. leei, K. clupeidae, K. iwatai, K. nova and K. thyrsites to an acceptable

level.


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Appendices

Appendix 1. Historical fish bait imports, 2008-2017

Prior to 2008, New Zealand’s commercial fishing industry was largely self-sufficient for fish bait

(Blackwell, 2013). Between 2008 and 2017, increases in demand from recreational and

commercial fishers resulted in the importation of a wide range of finfish (and cephalopod species

as fish bait.

1.1 Historical imports of finfish as bait, 2008-2017

The location of capture varied for each species between years depending on price and

availability (Table 21). Many fish bait species are widely distributed and the fishing fleets range

over a wide geographical area. (B. Burney, pers. comm., 2019; R. Clark, pers. comm., 2019; P.

Lamb, pers. comm., 2019; A. Spencer, pers. comm., 2019). Importantly, the reported country of

export or purchase of imported fish bait species may differ from the actual catch location (Jereb

et al., 2010; Granados-Amores et al., 2014; Norman et al., 2014; FAO, 2018; FishBase, 2019; B.

Burney, pers. comm., 2019; R. Clark, pers. comm., 2019).

Imports of redbait/Cape bonnetmouth (Emmelichthys nitidus nitidus) (Emmelichthyidae), silver

warehou (S. punctata) (Centrolophidae), and coalfish (Pollachius virens) (Gadidae) were minor

(< 1 tonne). These species are no longer imported (B. Burney, pers. comm., 2019). Other species

such as the European pilchard (Sardinia pilchardus) have not been recently imported. These

species were therefore not included in the list of species considered as fish bait (Table 5) and are

not considered further.

Table 21. Taxonomy and geographical distribution of finfish imported as fish bait, 2008-2017

Family Common name Species Geographical distribution1

Carangidae Pacific jack mackerel Trachurus symmetricus Eastern Pacific (Alaska to Mexico, Galapagos Is.)

Chilean jack mackerel Trachurus murphi South and Southwest Pacific, Southwest Atlantic

Japanese jack mackerel Trachurus japonicus Northwest Pacific, Japan, Korea

Centrolophidae Silver warehou Seriolella punctata Eastern Indian Ocean, Southwest and Southeast Pacific Ocean

Clupeidae Atlantic herring Clupea harengus Northwest (USA) and Northeast Atlantic Ocean, Europe

Pacific herring Clupea pallasii pallasii (Clupea pallasii)

Northeast and Northwest Pacific, Pacific Ocean

South American pilchard or sardine

Sardinops sagax Cosmopolitan: S. sagax ocellatus (S. Africa), S. s. neopilchardus (Australia), S. s. caerulus (California), S. s. melanostictus (China, Japan)

Round sardinella Sardinella aurita East and west Atlantic Ocean, Mediterranean Sea

Indian oil sardine Sardinella longiceps South and West Indian Ocean (East Indian coast)

Bali sardinella Sardinella lemuru Pacific Ocean (Asia, SE Asia, Australia)

Spotted sardinella Amblygaster sirm Indo-Pacific (Red Sea to Japan, New Guinea, Northern Australia, Fiji)



European pilchard Sardina pilchardus Northeast Atlantic, North Sea, Mediterranean

Emmelichthyidae Redbait/Cape bonnetmouth

Emmelichthys nitidus nitidus

Indo-Pacific, Eastern Atlantic Ocean

Engraulidae Californian anchovy Engraulis mordax Northeast Pacific Ocean (Vancouver to Mexico)

Peruvian anchovy Engraulis ringens Southeast Pacific Ocean (Peru to Chile)

Exocoetidae Tropical two-wing flying fish

Exocoetus volitans Widespread tropical, subtropical oceans

Sailfin flying fish Parexocoetus brachypterus

Indo-Pacific, West and East Atlantic Oceans

Gadidae Coalfish (saithe) Pollachius virens Northeast, Northwest Atlantic Ocean

Hemiramphidae Balao halfbeak Hemiramphus balao West Atlantic Ocean

Dussumier’s halfbeak Hyporamphus dussumieri Indo-Pacific (India to Okinawa)

Ballyhoo halfbeak Hemiramphus brasiliensis

West Atlantic (New York to Brazil), East Atlantic Oceans (Cape Verde to Angola)

Luke’s halfbeak Hemiramphus lutkei (= H. marginatus)

North Pacific (China, Japan), Southeast Asia, Pacific Islands

Jumping halfbeak Hemiramphus archipelagus (= H. marginatus)

Indo-Pacific, southeast Asia, Polynesia

Southern garfish Hyporhamphus melanochir

East Indian Ocean, Pacific Ocean (Australia)

Japanese halfbeak Hyporhamphus sajorii Northwest Pacific Ocean (Japan to China, Korea)

Mugilidae Flathead grey mullet Mugil cephalus Widespread in tropical, sub-tropical and temperate waters

Sand grey mullet, Myxus elongatus Southwest Pacific, Australia

South African mullet Chelon (Liza) richardsonii Southeast Atlantic, South Africa

Yellow eye mullet Aldrichetta forsteri Southwest Pacific, Australia, New Zealand

Scomberesocidae Sanmar, Pacific saury Cololabis saira Pacific Ocean (Japan, Alaska south to Mexico)

Scombridae Bonito (Bullet tuna) Auxis rochei Atlantic, Indian and Pacific Oceans

Bonito (Frigate tuna) Auxis thazard Atlantic, Indian and Pacific Oceans

Little tunny Euthynnus alletteratus Atlantic Ocean (widespread tropical, subtropical)

Skipjack tuna Katsuwonus pelamis Cosmopolitan (Atlantic, Indian and Pacific Oceans)

Atlantic bonito Sarda sarda Cosmopolitan (East and West Atlantic Ocean, Europe, Mediterranean Sea)

Eastern Pacific bonito Sarda chiliensis East Pacific Ocean (Chile to Peru)

Pacific bonito Sarda lineolata Northeast Pacific Ocean (Alaska to California)

Blue mackerel Scomber australasicus Indo-West Pacific

Chub (Pacific) mackerel Scomber japonicus Indo-Pacific, temperate



Atlantic mackerel Scomber scombrus Atlantic Ocean, Mediterranean Sea

Bigeye tuna Thunnus obesus Cosmopolitan (tropical, subtropical waters)

Source New Zealand fish bait industry: B. Burney, pers. comm., 2019; R. Clark, pers. comm., 2019); D Rutherford, pers. comm., 2019. Notes 1 Reported distribution (FishBase, 2019)

1.2 Historical imports of coleoid cephalopods as bait, 2008-2017

Coleoid cephalopod molluscs (squid, cuttlefish and octopus) support extensive commercial

fisheries worldwide, both for use as bait and for human consumption. Historical imports of

coleoid molluscs into New Zealand during 2008-2017 were mainly squid of the families

Loliginidae and Ommastrephidae (Table 22). As previously noted, the reported country of export

or purchase of imported bait species may differ from the actual catch location.

Coleoid cephalopod molluscs were not considered in the previous IRA for fish food and fish bait

(Cobb 2008) and are not specifically covered in the IHS for fish bait (FISFOOIC.ALL) (MPI

2011). They have historically been imported as a food-grade product under the IHS

(FISMARIC.ALL) (MPI 2008), of which a variable proportion has then been on-sold as bait.

This is because the quality of industrial (non-food grade) product is considered too low for

commercial use as bait (B. Burney, pers. comm. 2018). No trade import data are available

separately for coleoid molluscs imported as fish bait. Of the total imports (by volume) of coleoid

cephalopod molluscs during 2013-2015 (Notes

1 Jereb & Roper (2010).

2 Within the Loglinidae, the genus Loligo is paraphyletic, with species based largely on geographical distribution 3 The classification of species within the Octopus species complex is currently under review, with 350 named species reported

(Norman et al. 2014) and over 150 potentially undescribed species known from tropical and subtropical waters (FishBase 2019).

Table 23), industry sources estimate that 700-800 t, representing approximately 35% of these

annual totals, were redirected annually for use as fish bait (B. Burney, pers. comm. 2019).

Table 22. Classification and distribution of coleoid cephalopod molluscs imported, 2008-2017


Loliginidae (Long-finned squids)2.

Patagonian squid Doryteuthis gahi (= Amerigo (Loligo) gahi)

Southeast Atlantic, Falkland Islands, Southwest Pacific

California market squid, opalescent inshore squid

Doryteuthis opalescens (= Loligo opalescens)

East Pacific (Alaska to Mexico)

Long-finned inshore squid

Doryteuthis pealeii (= Loligo (Amerigo) pealeii)

West Atlantic (Newfoundland to Venezuela)

Loligo squid undefined Loligo vulgaris Europe, Mediterranean, Northeast Atlantic

Spear squid Heterololigo (Loligo) bleekeri

Northwest Pacific (Japan to Korea)

Common squid Loligo forbesii Mediterranean Sea, Northeast Atlantic (Scotland to the Azores)



Chokka squid Loligo reynaudii (= L. vulgaris reynaudii)

Southern Africa, Southeast Atlantic, Southwest Indian Ocean

Swordtip squid Uroteuthis edulis (= Photololigo edulis)

East, West Indian Oceans, Pacific (Southeast Asia, Japan, China)

Spear squid Heterololigo (Loligo) bleekeri

Northwest Pacific (Japan to Korea)

Common squid Loligo forbesii Mediterranean Sea, Northeast Atlantic (Scotland to the Azores)

Chokka squid Loligo reynaudii (= L. vulgaris reynaudii)

Southern Africa, Southeast Atlantic, Southwest Indian Ocean

Swordtip squid Uroteuthis edulis (= Photololigo edulis)

East, West Indian Oceans, Pacific (Southeast Asia, Japan, China)

Juvenile squid Loligo spp. (unclassified) Widespread oceanic waters

Ommastrephidae (flying, short-finned squids)

Subfamily Illicinae Argentine shortfin squid Illex argentinus (= Ommastrephes argentines)

Southwest Atlantic (Brazil to Argentina, Falkland Islands)

Broadtail shortfin squid/ northern shortfin squid

Illex coindetii (= Illex illecebrosus)

East Atlantic (North Sea, to Namibia), West Atlantic (Caribbean to French Guiana), Mediterranean Sea

Sharp-tail shortfin squid Illex oxygonius West North Atlantic (New Jersey to Gulf of Mexico), East Atlantic

Juvenile squid Illex spp. unclassified Widespread oceanic waters

Subfamily Ommastrephinae

Jumbo flying squid Dosidicus gigas East Pacific (Alaska to Peru)

Subfamily Todarodinae

Japanese flying squid Todarodes pacificus (= Ommastrephes pacificus)

Northwest Pacific, Southeast Asia, Australia

European flying squid Todarodes sagittatus (= Loligo saggittata)

Northeast, North, South Atlantic, Mediterranean Sea

Broad squid Nototodarus sloanii, N. gouldi

Southern Ocean, New Zealand

Sepiidae (Cuttlefishes)

Needle cuttlefish Sepia aculeata Indo-West Pacific (India to Japan)

Spineless cuttlefish Sepiella inermis (= Sepia affinis, S. inermis)

Indo-West Pacific (India to Indonesia and South China Sea)

European cuttlefish Sepia officinalis Northeast Atlantic, Mediterranean Sea

Curve spine cuttlefish Sepia recurvirostra Indo-West Pacific (China to the Philippines, Indonesia and Pakistan).

Juvenile cuttlefish Sepia spp., Sepiella spp. Widespread

Octopodidae (Octopus)

Octopus Octopus species complex3

Widespread coastal tropical, subtropical waters

Notes 1 Jereb & Roper (2010). 2 Within the Loglinidae, the genus Loligo is paraphyletic, with species based largely on geographical distribution 3 The classification of species within the Octopus species complex is currently under review, with 350 named species reported

(Norman et al. 2014) and over 150 potentially undescribed species known from tropical and subtropical waters (FishBase 2019).


Table 23. Imports of coleoid cephalopod molluscs, 2013-2015

Total volume1 (t)

Year 2013 2014 2015

Family Common name Scientific name

Loliginidae, Ommastrephidae

Squid Illex spp., Ommastrephes spp., Loligo spp., Nototodarus spp., Sepioteuthis spp.

1,900 2,019 1,949

Sepiidae Cuttlefish Sepia spp. 44 30 41

Octopodidae Octopus Octopus spp. 52 71 47

Total 1,996 2,140 2,037

Source Trade data, Department of Statistics (2017). Notes 1 Estimated volume (t) of imports for human consumption and for use as bait


Appendix 2. Pathogen association by fish bait host species considered in this risk analysis (Table 5)

Table 24. Risk organisms and associated fish bait species where specific risk management measures are proposed

Host MABV RSIV/ISKNV ENV NNV IHNV VHSV Streptococcus agalactiae III: 283

Streptococcus iniae

OIE-listed N Y N N Y Y N N

Family Carangidae

Trachurus japonicus √ √ √ √

Trachurus murphi √

Trachurus symmetricus √

Trachurus trachurus √

Family Clupeidae

Ambylygaster sirm √ √

Clupea harengus √ √ √ √

Clupea pallasii pallasii √ √ √ √ √

Sardinella aurita √ √

Sardinella lemuru √ √

Sardinella longiceps √ √

Sardinops sagax √ √ √

Family Engraulidae

Engraulis mordax √

Engraulis ringens

Family Hemiramphidae

Hemiramphus balao

Hemiramphus brasiliensis

Hemiramphus dussumieri

Hemiramphus lutkei



Streptococcus iniae

Family Mugilidae

Chelon auratus √

Chelon labrosus √

Chelon ramada √

Mugil cephalus √ √ √ √ √

Family Scomberosocidae

Cololabias saira

Family Scombridae

Auxis rochei

Auxis thazard

Euthynnus alletteratus

Katsuwonas pelamis

Sarda chilensis

Sarda lineolata

Sarda sarda

Scomber australasicus √

Scomber japonicus √ √ √ √

Scomber scombrus √ √

Thunnus obesus

Family Loliginae

Doryteuthis opalescens

Loligo spp.

Uroteuthis duvaucelli

Family Ommastrephidae

Illex argentinus

Illex coindetti

Todarodes pacificus √



Streptococcus iniae

Family Sepiidae

Sepia recurvirostrata

Sepia spp.

Family Octopodidae

Octopus vulgaris √

Octopus spp.

Note: General risk management options are likely to provide sufficient reduction in pathogen occurrence for Edwardsiella spp.,Enteromyxum leei, Kudoa clupeidae, K. iwatai, K. nova and K. thyrsites. No specific risk management options are therefore necessary.


Appendix 3. Summary and evaluation of risk management options

Table 25 summarises the qualitative evaluation of the expected reduction in pathogen load

associated with each risk management option in the relevant risk organism chapter.

The estimated levels of reduction in occurrence are:

Eliminate > 95% reduction in pathogen load

Substantial 71-95% reduction in pathogen load

Moderate 51-70% reduction in pathogen load

Slight < 50% reduction in pathogen load

Table 25. Assumed reduction in pathogen load associated with risk management options

Risk organism MABV RSIV/

ISKNV ENV NNV IHNV VHSV

General risk management options

Health certification Moderate Moderate Moderate Moderate Moderate Moderate

Frozen storage (-18 ° to

-20 ° for 4 months (17 weeks)

No effect No effect No effect No effect No effect Slight

Species specific risk management options

Source from species not associated with risk

organism Substantial Substantial Substantial Substantial Substantial Substantial

Source from region/country free of

risk organism Substantial Substantial Substantial Substantial Substantial Substantial


Substantial Substantial Substantial Substantial Substantial Substantial

Ionising radiation Substantial Substantial Substantial Substantial Substantial Substantial

Risk organism Edwardsiella

spp.

Streptococcus agalactiae III:

283

Streptococcus iniae

Enteromyxum leei

Kudoa clupeidae, K. iwatai, K. nova, K.

thyrsites

General risk management options

Health certification Moderate Moderate Moderate Moderate Moderate

Frozen storage ( -18 °

to -20 °C) for 4 months (17 weeks)

Substantial No effect No effect Substantial Substantial

Species specific risk management options

Source from species not associated with risk

organism N/A Substantial Substantial N/A N/A

Source from region/country free of

risk organism N/A Substantial Substantial N/A N/A


N/A Substantial Substantial N/A N/A

Ionising radiation N/Al Substantial Substantial N/A N/A

Notes “No effect” means a risk management option is unlikely to reduce pathogen occurrence N/A” means a risk management option is not applicable.

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