Diseases of crayfish: A review

Journal of Invertebrate Pathology 106 (2011) 54–70

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Journal of Invertebrate Pathology journal homepage: www.elsevier.com/locate/jip

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Diseases of crayfish: A review Matt Longshaw ⇑ Cefas Weymouth Laboratory, Barrack Road, The Nothe, Weymouth, Dorset DT4 8UB, UK

a r t i c l e

i n f o

Keywords: Crayfish Virus Fungi Bacteria Protista Metazoa

a b s t r a c t A systematic review of parasites, pathogens and commensals of freshwater crayfish has been conducted. All major groups of disease causing agents have been covered including viruses, bacteria, fungi, protistans and metazoans. Most agents tend to cause limited problems for crayfish. Exceptions to this include fungi, bacteria and viruses. However, in many cases, these tend to be isolated reports in either a specific geographical location or in individual animals. The apparent absence of pathology associated with these agents in crayfish should not be taken to suggest that movements of crayfish to new geographical areas is necessarily acceptable. Several examples are given where seemingly healthy animals have been moved to new areas leading to mortality of other crayfish within the same area as a direct result of transmission of pathogens to naïve hosts. Some future research needs are proposed, including the need for pathogen characterisation and production of disease-free crayfish for aquaculture. Crown Copyright Ó 2010 Published by Elsevier Inc. All rights reserved.

Contents 1.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1. Viruses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1.1. Intranuclear bacilliform viruses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1.2. Birnaviridae. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1.3. Nimaviridae. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1.4. Parvoviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1.5. Picornaviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1.6. Reoviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1.7. Totiviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1.8. Unidentified viral infections . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2. Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.1. Coxiella cheraxi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.2. Nocardia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.3. Spiroplasma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.4. Vibrio. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.5. Aeromonas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3. Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.1. Class Oomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.2. Class Sordariomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3.3. Microsporidia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4. Mesomycetozoea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.5. Protista. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.5.1. Ciliata . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.5.2. Phylum Apicomplexa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6. Digenea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.1. Family Allocreadiidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.2. Family Choanocotylidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.3. Family Cladorchidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

⇑ Fax: +44 (0)1305 206601. E-mail address: [email protected] 0022-2011/$ - see front matter Crown Copyright Ó 2010 Published by Elsevier Inc. All rights reserved. doi:10.1016/j.jip.2010.09.013

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1.6.4. Family Gorgoderidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.5. Family Haematoloechidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.6. Family Macroderoididae. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.7. Family Microphallidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.8. Family Opecoelidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.9. Family Orchipedidae. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.10. Family Paragonimidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.11. Superfamily Plagiorchioidea. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.12. Family Psilostomatidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.13. Family Reniferidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6.14. Family Troglotrematidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.7. Cestoda. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.8. Acanthocephala . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.9. Nematoda. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.10. Branchiobdellida . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.11. Temnocephalida . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.12. Other fouling organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.13. Idiopathic conditions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions and future directions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction Freshwater crayfish are widespread crustaceans occurring on all continents except Antarctica, either as native species or following anthropogenic movements. Several species have been used for aquaculture purposes and more recently, there has been an increase in the sale of crayfish for aquaria. Movements of several species have been responsible for the transfer of the devastating crayfish plague (Aphanomyces astaci) which has led to the complete or near extinction of several populations of native crayfish in Europe. There has therefore been a large focus of research on crayfish plague including characterisation, development of diagnostic methods, description of geographical distribution and evaluation of impact at individual and population levels. However, because of the recognition that A. astaci is a major mortality driver, some cases of unexplained mortalities may have been attributed to crayfish plague despite absence of clear diagnostic evidence (Alderman and Polglase, 1988). Perhaps reflecting the locality of researchers and of major aquaculture production areas, much of the research focus has been in Australia, the United Kingdom, Germany and the United States of America with other countries contributing to a lesser extent. Many papers published in the field of crayfish disease are of a descriptive nature, describe pathogen life cycles or provide a case review of a mortality event. Furthermore, due to problems associated with white spot virus syndrome (WSSV), there has been a focus on the distribution, detection and transmission of the virus to crayfish and other Crustacea (Stentiford et al., 2009). In addition, with the recent advent of molecular diagnostic tests, there have been several publications describing new methods for identifying pathogens in crayfish, such as A. astaci. With increasing evidence of other pathogens of importance, in particular viruses, it is timely to provide a review of the current knowledge of crayfish pathogens. This review is not intended to be exhaustive; rather it will cover the major groups of pathogens including viruses, bacteria, fungi, protistans, metazoan parasites and idiopathic conditions. It will consider the impact of these infections on individuals and populations, provide information on available diagnostic methods and provide gaps in our knowledge and suggest future research needs. 1.1. Viruses Several viruses have been reported from crayfish, although as indicated by Edgerton et al. (2002a) they remain ‘‘a conspicuously

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understudied group of pathogens of crayfish”. Most crayfish viruses reported are found in certain commercially important species such as members of the genus Cherax, and the majority are intranuclear bacilliform viruses (IBVs). Furthermore, studies on crayfish viruses reflect research interests of selected individuals and their associated institutions rather than necessarily the apparent geographical bias in distribution. Further examples of viral infections will likely be identified as more hosts are examined specifically for viruses. The taxonomy of crayfish viruses is in a state of flux; the higher taxonomic status for some is currently unresolved. 1.1.1. Intranuclear bacilliform viruses Intranuclear bacilliform viruses (IBVs) are a group of nonoccluded double stranded DNA viruses; due to a lack of molecular, immunological and biochemical data on these viruses, their taxonomic position is currently unknown. The IBV’s were previously assigned to the family Baculoviridae (Stentiford et al., 2004). They are restricted to the hepatopancreas and the gut. 1.1.1.1. Astacus astacus bacilliform virus (AaBV). To date AaBV has only been reported from A. astacus from Finland at prevalences up to 100% and with variable intensity and no apparent mortality or morbidity (Edgerton et al., 1996b). Infections occur in the hepatopancreas with infected cells being only slightly hypertrophied and with emarginated chromatin. In some, infected nuclei were compartmentalised by remaining chromatin. Sloughing of infected cells and necrosis and encapsulation of affected tubules were also noted. The virions have a rod-shaped nucleocapsid with a trilamellar envelope and a subapical unilateral expansion. Virions measure 70  340 nm with nucleocapsids measuring 50  260 nm. 1.1.1.2. Austropotamobius pallipes bacilliform virus (ApBV). Edgerton et al. (2002b) reported mortalities of the white clawed crayfish A. pallipes in France which appeared to be associated with the presence of ApBV; however no consistent clinical signs were apparent. In addition, the virus was present at relatively low intensity, suggesting that the virus was unlikely to be the primary cause of the mortality. Subsequently, Edgerton (2003) showed that the virus was present in other populations of A. pallipes; it was recently found in several populations in England and Wales with limited effects (Longshaw, Stebbing and Hockley, in preparation). Infected hepatocytes and cells of the midgut are hypertrophied with emarginated chromatin. Although not reported by Edgerton et al. (2002b) or Edgerton (2003), infected nuclei can contain septae giving rise to

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nuclear partitioning (Longshaw, personal observations). Similar to other IBVs, the virions consist of a nucleocapsid with a nucleoprotein core and a less electron dense capsid. A trilamellar envelope with a unilateral expansion surrounds the nucleocapsid. Virions measure 60  260 nm and the nucleocapsids measure 50  220 nm.

Infected cells are hypertrophied with emarginated chromatin and may be sloughed into tubule lumina. Virions are rod-shaped, measuring 70  240 nm and the nucleocapsids measure 190  40 nm. However, the virus remains to be properly described.

1.1.1.3. Cherax destructor bacilliform virus (CdBV). CdBV was reported from C. destructor from two farms in south Australia at an overall prevalence of 21% with low intensity (Edgerton, 1996). Infected nuclei in the hepatopancreas are hypertrophied and contain emarginated chromatin with amorphous eosinophilic inclusions; intranuclear septae may be present which compartmentalise the nuclear contents. In the single study of CdBV, most capsids were developing; in those virions that were somewhat more mature, the envelope was loosely applied with an apparent kink midway without a true lateral envelope expansion or nucleocapsid tail (Edgerton, 1996). Virions measured 70  300 nm (atypically 450 nm long); nucleocapsids measured 50  260 nm.

1.1.2. Birnaviridae Birnaviridae are double stranded RNA viruses. An important pathogen of fish, infectious pancreatic necrosis virus (IPNV) has been associated with mortalities in salmonid fish. Halder and Ahne (1988) exposed A. astacus to IPNV through various routes. They were able to transmit the virus back to naïve rainbow trout fry and eggs. IPNV could be detected in haemocytes of the antennal gland, gills and hepatopancreas of exposed crayfish although no pathological changes were noted in infected crayfish. The virus persisted in crayfish for up to a year after exposure, suggesting replication is possible and raising concerns over the role of crayfish in transmission of the virus to fish.

1.1.1.4. Cherax quadricarinatus bacilliform virus (CqBV). Originally reported as Cherax bacilliform virus from crayfish in northern Australia by Anderson and Prior (1992), CqBV has since been reported in C. quadricarinatus farmed in other parts of Australia, the USA, Chile and Ecuador (Bateman et al., 2005; Edgerton, 1996; Edgerton and Owens, 1999; Groff et al., 1993; Hauck et al., 2001; Romero and Jiménez, 2002). Although Groff et al. (1993) postulated that the infection was introduced into the USA via importation of broodstock from Australia, Hauck et al. (2001) could not determine how the virus was spread to Utah, USA. Furthermore, they suggested that crayfish from infected sites in Utah had been translocated to several other US states and several countries, including Bermuda, Haiti, Jamaica, Virgin Islands, Colombia, India and Saudi Arabia. Conflicting information on impact of the virus exists, with Anderson and Prior (1992) and Romero and Jiménez (2002) suggesting that CqBV infections were not associated with mortality or disease. However, CqBV causes stunted growth and low grade mortality (Edgerton et al., 2002a; Groff et al., 1993). All ages of crayfish are susceptible to the virus (Edgerton and Owens, 1997). No pathognomonic external signs of infection have been reported, although heavily infected individuals are weakened with reduced or absent tail-flick response (Edgerton, 1996). At the light microscope level infected cells have hypertrophied nuclei with emarginated chromatin and nucleoli and in some, intranuclear septae as nuclear partitions (Edgerton, 1996; Groff et al., 1993). Tubular degeneration and loss of infected cells occurred in infected crayfish (Groff et al., 1993; Romero and Jiménez, 2002). Virions consist of a rod-shaped nucleocapsid surrounded by a loose trilaminar envelope and are distributed throughout infected nuclei (Edgerton, 1996). The unilaterally expanded envelope contains a flexed tail-like structure. Measurements are variable among studies with Anderson and Prior (1992) suggesting virions measure 70  200 nm, close to the measurements observed by Hauck et al. (2001) of 70  220 nm. Measurements provided by Edgerton (1996) and Groff et al. (1993) are slightly larger at 100  260 nm and 100  290 nm respectively. Similarly, nucleocapsid measurements are variable with measurements ranging from 30  150 nm by Anderson and Prior (1992), to 40  180 nm by Hauck et al. (2001) and 50  215 nm by Groff et al. (1993). Differences in measurements may correspond to different strains of either host or virus, or may equally be due to differences in fixation and processing methods (Edgerton et al., 2002b; Hauck et al., 2001).

1.1.3. Nimaviridae White spot syndrome virus is a double stranded DNA virus in the family Nimaviridae which affects a wide range of crustacean hosts, including crayfish (Stentiford et al., 2009). Virions are elliptical to rod-shaped, bound by a trilamellar envelope and measure 80–120  250–380 nm. Because of its importance, White Spot Disease (WSD) is listed under EU legislation, which is reflected in the plethora of research papers written. Experimental transmission of the virus to naïve crayfish has been demonstrated using haemolymph from infected Penaeus chinensis to C. quadricarinatus by Shi et al. (2000); from rotifers to Procambarus clarkii possibly via mechanical transmission rather than true biological transmission (Yan et al., 2007); to A. astacus by Jiravanichpaisal et al. (2004); from Penaeus monodon to Cherax destructor albidus by Edgerton (2004) and from P. monodon gill homogenate to P. leniusculus (Jiravanichpaisal et al., 2001). Du et al. (2007) suggested that virus replication in P. clarkii is so successful that it can be used as a biological method to amplify virus particles for subsequent studies. Decreased temperatures, although slowing infection progression, does not entirely stop transmission and development of the virus in crayfish (Du et al., 2008). Recently, Baumgartner et al. (2009) demonstrated that both farmed and wild P. clarkii and Procambarus zonangulus in Louisiana were natural hosts for WSSV. Several methods for WSSV identification have been developed, including molecular methods (Huang et al., 2001; Jiravanichpaisal et al., 2001; Yan et al., 2007). The official OIE method recommends a nested polymerase chain reaction (PCR) approach using primer sets developed by Lo et al. (1997). However, Claydon et al. (2004) expressed concerns with false WSSV positive results associated with the primer sets of Lo et al. (1997) when used with C. quadricarinatus. Although crayfish and other invertebrates do not have a true adaptive immune system, efforts have been made to induce protection to WSSV through exposure of naïve crayfish to viral envelope proteins or to binary-ethylenimine inactivated WSSV. The WSSV envelope proteins VP28 and VP19 have been produced in several expression vectors, and injected intramuscularly into crayfish. Groups of crayfish subsequently exposed to WSSV had a lower overall mortality (Du et al., 2006; Xu et al., 2006). The slightly different approach of Zhu et al. (2009) involved use of binaryethylenimine inactivated WSSV, which also led to mortality reductions in crayfish exposed to WSSV.

1.1.1.5. Pacifastacus leniusculus bacilliform virus (PlBV). This virus was originally reported as a personal communication in Hauck et al. (2001) without details of virus or host. Subsequently, data were presented in conference proceedings by Hedrick et al. (1995). The virus apparently infects tubule cell nuclei of the hepatopancreas.

1.1.4. Parvoviridae Parvoviruses are single stranded DNA viruses; several parvoviruses and presumptive parvoviruses occur in Australian Cherax spp.

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1.1.4.1. Cherax quadricarinatus parvo-like virus (CqPlV) of Edgerton et al. (2000). C. quadricarinatus farmed in Australia were reported to be affected by a parvo-like virus by Edgerton et al. (2000). Nuclei of affected gills showed typical emarginated chromatin and peripheral nucleoli. Ultrastructurally, icosahedral virus particles measuring 20 nm in diameter were observed in affected nuclei. Limited mortality and morbidity were associated with this infection (Edgerton et al., 2000). Molecular details are lacking for this virus. 1.1.4.2. Cherax quadricarinatus parvovirus of Bowater et al. (2002). A second parvovirus causing mass mortality was described from farmed C. quadricarinatus in northern Queensland, Australia by Bowater et al. (2002). Affected crayfish were moribund, had soft shells and were anorexic and weak. Intranuclear inclusion bodies with emarginated chromatin were present in gills, cuticular epithelium, gut epithelium and connective tissue. To a lesser extent the antennal gland, haematopoeitic tissue, epithelial cells of the seminiferous tubules and interstitial tissue of the ovary were affected. No infections were detected in hepatopancreatocytes, neurones or the heart by histology. Ultrastructurally, viral particles were hexagonal with an average diameter of 19.5 nm. Despite the very similar size and site of infection reported by Edgerton et al. (2000), Bowater et al. (2002) considered the form reported by them to be distinct since they did not find gill lesions associated with infection. Molecular data are required to determine the relationship for this and the parvovirus of Edgerton et al. (2000) which were found in the same host from the same geographical area. 1.1.4.3. Cherax destructor systemic parvo-like virus (CdSPV). A systemic infection of most tissues, with focus in gills, was reported from a single C. destructor farmed in south Australia. The infected animal was moribund with an opaque abdomen (Edgerton et al., 1997, 2000). Intranuclear Cowdry type A inclusions were noted in all affected tissues with nuclei having emarginated chromatin. Tissues affected included hepatopancreas, midgut, epicardium, abdominal muscle and connective tissues. Necrosis and haemocyte infiltration was associated with infection. Virus particles were icosahedral and measured 21 nm (Edgerton et al., 1997). 1.1.4.4. Spawner-isolated mortality virus (SMV). Spawner-isolated mortality virus (SMV) has been tentatively placed in the Parvoviridae and was originally described from P. monodon (Fraser and Owens, 1996). The virus is associated with high levels of mortality in shrimp which show haemocyte infiltration and necrosis of affected tissues. No pathognomonic signs are apparent; there are no intranuclear inclusion bodies. Ultrastructurally, virions are icosahedral and measure 20 nm in diameter. In naturally infected C. quadricarinatus, positive in situ hybridisation reactions using an SMV probe were noted in hepatopancreas, midgut, epithelium of gonadal tissue, and occasionally in heart, haemocytes and connective tissues (Fraser and Owens, 1996; Owens and McElnea, 2000). However, similar to P. monodon infections, no external signs were noted in affected C. quadricarinatus. 1.1.4.5. Penaeus merguiensis densovirus (PmergDNV). Penaeus merguiensis densovirus is a hepatopancreatic parvovirus which causes reduction in growth of P. monodon and infects at least ten species of penaeid prawns worldwide (La Fauce and Owens, 2007). C. quadricarinatus experimentally exposed to PmergDNV were lethargic with limited tail-flick response within a week of being exposed to the virus via oral or injection routes of infection. Mortalities of virally challenged animals occurred within 24 h. However, no histopathological changes, such as intranuclear inclusion bodies attributable to PmergDNV, were noted in challenged animals and there was no evidence of viral replication in C. quadricarinatus.

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La Fauce and Owens (2007) suggested that C. quadricarinatus could act as a carrier for the virus. 1.1.5. Picornaviridae Picornaviridae are single stranded RNA viruses. Following a survey of yabbies by Edgerton (1999), a new picorna-like virus (Cherax albidus picorna-like virus (CaPV)) associated with mortalities was reported. Subsequently, Jones and Lawrence (2001) reported the same virus associated with mortality in farmed C. albidus in western Australia. Although prevalence was <5%, its distribution throughout the farming area was widespread. Infections were characterised by large basophilic inclusions occupying the entire nuclei or small eosinophilic intranuclear nuclei inclusions in histological section. Nuclear changes occurred in the interstitial tissues of the digestive gland, labyrinth epithelium of the antennal gland and the hindgut. Ultrastructurally, non-enveloped virions in paracrystalline arrays measured 13–19 nm. Jones and Lawrence (2001) were unsure about the exact taxonomic position of the virus and related it to either the Circoviridae or the Picornaviridae. 1.1.6. Reoviridae Reoviruses are double stranded RNA viruses and a single example, Cherax quadricarinatus reolike virus (CqRV) has been described. Edgerton et al. (2000) reported eosinophilic, cytoplasmic inclusion bodies in the hepatopancreas of C. quadricarinatus in northern Queensland Australia. Affected tubules were surrounded by a haemocytic infiltration; ultrastructurally infected nuclei contained regularly sized non-enveloped hexagonal and pentagonal shaped virions measuring 35–40 nm. The single affected animal examined by Edgerton et al. (2000) was moribund; Bowater et al. (2002) and La Fauce and Owens (2007) considered the infection to be relatively benign. 1.1.7. Totiviridae Totiviruses are double stranded RNA viruses. Cherax quadricarinatus Giardia-like virus (CGV) was originally reported as Cherax Giardia-like virus (GCV) from the hepatopancreas of C. quadricarinatus from Australia; affected host cells were mildly hypertrophic (Edgerton et al., 1994). The infections are more prominent in juvenile crayfish and result in low grade mortality in farmed populations (Edgerton and Owens, 1997). Ultrastructurally, non-enveloped virions measured 25 nm in cross section and were hexagonal or pentagonal, suggesting an icosahedral shape. It continues to occur in Queensland, causing limited mortalities (Owens and McElnea, 2000). Poulos et al. (2006) considered that since the virus had not been isolated or characterised it could not be unequivocally placed alongside other Giardia-like viruses and considered the virus should be referred to as a putative toti virus. 1.1.8. Unidentified viral infections A wide range of viruses have been reported in crayfish. However, many have been poorly described or remain unclassified and there thus remains a pressing need to adequately identify and properly describe viruses from crayfish. This includes utilising clinical signs, histology, ultrastructure as well as molecular tools to discriminate crayfish virus. In some cases, presumptive viruses apparent in histological sections have not been further described. For example Romero and Jiménez (2002) report on the presence of Cowdry type A inclusions in the gills and stomach of C. quadricarinatus from Ecuador. These inclusion bodies are typical of viral infections, and although the authors tested affected tissue samples for infectious hypodermal and haematopoietic necrosis virus (IHHNV) and white spot syndrome virus using in situ hybridisation, no further analyses were conducted. Hypertrophied nuclei were noted in the gills of several C. quadricarinatus from Australia; no further characterisation of this putative viral infection was made

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(Edgerton and Owens, 1999). In addition, reports of viral infections in conference abstracts (e.g. P. leniusculus bacilliform virus reported by Hauck et al., 2001) or in unpublished thesis such as the picorna-like virus from the connective tissues of the gill in A. astacus noted by Edgerton et al. (2002a) need to be re-isolated and properly described. 1.2. Bacteria Bacterial infections of crayfish are common and widespread and many consist of opportunistic infections. Typically bacteria isolated from crayfish include representatives of the genera Acinetobacter, Aeromonas, Bacillus, Citrobacter, Corynebacterium, Flavobacterium, Micrococcus, Pseudomonas, Staphylococcus and Vibrio (Alderman and Polglase, 1988; Edgerton et al., 2002a; Quaglio et al., 2006b; Smith and Söderhäll, 1986; Vey, 1986). In certain cases, bacterial infections of crayfish lead to mortalities in both farmed and wild animals, particularly in combination with other underlying poor conditions and in other cases are reported in asymptomatic crayfish (Edgerton et al., 2002a; Quaglio et al., 2006b). Bacteraemia of crayfish presents as perivascular cuffing of haemolymph vessels, haemocyte aggregations and nodule formation (Edgerton and Owens, 1999; Quaglio et al., 2006b; Romero and Jiménez, 2002). 1.2.1. Coxiella cheraxi A systemic intracellular bacterial infection of C. quadricarinatus was characterised using molecular techniques by Tan and Owens (2000) and named as C. cheraxi. The infection was associated with mortalities in the host species. Although it was considered by these authors to be systemic, it appears to primarily infect gill tissue with hepatopancreas becoming infected later during heavy infections (La Fauce and Owens, 2007). The taxonomic position of the bacterium was subsequently confirmed by Cooper et al. (2007) as a member of the previously monotypic genus Coxiella which contains C. burnetii, the causative agent of Q fever in humans and various other mammals. The systemic intracellular bacterial infection described in Ecuadorian C. quadricarinatus by Jiménez and Romero (1997) and Romero et al. (2000) is likely to be closely related to C. cheraxi based on the pathology, morphology and localisation of the infection. A second intracellular bacterial infection was reported as a Rickettsia-like organism (RLO) in the hepatopancreas of C. quadricarinatus by Edgerton and Prior (1999), which was considered distinct from C. cheraxi due to the different tissue tropism. Given that it was only recorded in a single animal, this is unlikely to be of major concern. Although probably closely related to Coxiella spp., the lack of molecular data has meant that this infection must remain as a RLO. An unidentified Rickettsia-like organism was noted in the ovaries, hepatopancreas and heart of the parthenogenetic marbled crayfish with an associated granuloma surrounding the infection (Vogt et al., 2004). 1.2.2. Nocardia Members of the genus Nocardia, containing at least 80 species, are gram-positive, acid-fast, filamentous bacteria. Furthermore, they can form branched substrate hyphae which then fragment into rod-shaped elements which are non-motile (Friedman et al., 1998). Nocardia have been reported in soil and a wide range of animal hosts, including humans, fish and shellfish (Engelsma et al., 2008; Friedman et al., 1998; Kitao et al., 1989). Typically, Nocardia spp. are associated with disease. Alderman et al. (1986) reported on a putative Nocardia in a single A. pallipes in England. The infected animal was lethargic and uncoordinated and moving around in daylight. Externally, the animal had a melanised fracture of the exoskeleton. Several nodules, formed from haemocyte infiltration contained acidfast, branching filaments, led the authors to suggest that the infection was most probably Nocardia. No further characterisation of

the putative bacteria in A. pallipes was carried out and there have been no further reports of any Nocardia spp. in crayfish. 1.2.3. Spiroplasma Spiroplasmas are small, helical bacteria associated with arthropods, ticks and plants with 40 species described to date. A further several hundred isolates have also been partially described, which may represent new species (Regassa and Gasparich, 2006). Spiroplasma spp., particularly in insects, act as male killing organisms, distorting sex ratios in some populations (Enigl and Schausberger, 2007; Kageyama et al., 2007). To date, no evidence has been provided to show that spiroplasmas have a role in distorting sex ratios of crayfish. A Spiroplasma sp. has been reported in P. clarkii affected by ‘‘crayfish weakness disease” in the same locality as mitten crabs (Eriocheir sinensis) affected by tremor disease (caused by Spiroplasma sp.). Wang et al. (2005) however, were unable to transmit the bacteria from freshwater crabs to crayfish. Subsequently Bi et al. (2008) showed through molecular sequencing, that the forms present in mitten crabs, in P. clarkii, and in Penaeus vannamei, were the same species, closely allied to S. mirum. In controlled experimental transmissions, the bacteria occur systemically in the haemolymph, connective tissue of the gonads, pereiopods, gut hepatopancreas, nerves, heart and gills (Wang et al., 2005). Diagnostic tools including PCR and an enzyme-linked immunosorbent assay (ELISA) have been developed for detection of the pathogen (Bi et al., 2008; Ding et al., 2007; Wang et al., 2009). Experimentally, oxytetracycline is effective in treating Spiroplasma in mitten crabs, with potential for treatment of this bacteria in crayfish (Liang et al., 2009). 1.2.4. Vibrio Several reports of mortalities associated with Vibrio spp. have been made. Although V. cholerae is associated with low level mortalities in P. clarkii, usually the causative agent of Vibrio associated mortality of crayfish is V. mimicus (Thune et al., 1991). Infections are systemic, often within haemolymph, and other than lethargy, no external pathognomonic signs occur (Eaves and Ketterer, 1994; Thune et al., 1991). Several ribotypes of V. mimicus have been isolated, all of which can be pathogenic to crayfish, particularly under aquaculture conditions (Eaves and Ketterer, 1994; Wong et al., 1995). Additional to directly causing mortality in crayfish, Vibrio infections in crayfish can lead to gastroenteritis in humans through ingestion of raw or undercooked crayfish (Bean et al., 1998; Eaves and Ketterer, 1994). 1.2.5. Aeromonas Aeromonas hydrophila, ubiquitous in the freshwater environment, is associated with disease in fish and shellfish and occasionally in humans (Tulsidas et al., 2008). The bacterium is sometimes isolated from apparently healthy crayfish but is considered to have the potential to cause problems under culture conditions (Quaglio et al., 2006a). Jiravanichpaisal et al. (2009) isolated A. hydrophila and several other bacteria from unhealthy Pasifastacus leniusculus collected in Sweden. Following culture of bacteria in broth, bacterial extracellular products were separated from the bacteria and injected into naïve crayfish which were then held at different temperatures. Although several bacteria were considered pathogenic to varying degrees, most mortalities were associated with A. hydrophila. In crayfish injected with A. hydrophila and held at 22 °C, mortalities began 6 h post-injection. Histologically, affected crayfish had necrotic lesions in most tissues and haemocyte aggregations within haemal sinuses. 1.3. Fungi The most studied disease of crayfish is crayfish plague caused by A. astaci. However, many different fungi have been isolated from

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crayfish, some of which have been associated with disease, including Alternaria sp., Hormodendrum sp. (Class Dothideomycetes), Aspergillus sp. (Class Eurotiomycetes) and Uncinula sp. (Class Leotiomycetes) (Edgerton et al., 2002a). 1.3.1. Class Oomycetes The class Oomycetes contains two important genera associated with mortalities in crayfish, namely Saprolegnia and Aphanomyces. Saprolegnia spp. are generally opportunistic, invading dead eggs and moribund larvae of crayfish (Herbert, 1987; Royo et al., 2002; Vey, 1979, 1981). However, they can infect healthy crayfish; infection through damaged cuticle can dramatically increase transmission and mortality (Diéguez-Uribeondo et al., 1994). Declines of the spiny-cheek crayfish Orconectes limosus in Germany were associated with several Saprolegnia spp., found both on external surfaces and more deeply within abdominal tissues (Hirsch et al., 2008). Around 35 species of Aphanomyces have been described in three main lineages; plant parasitic, animal parasitic and saprophytic or opportunistic parasitic (Diéguez-Uribeondo et al., 2009). Additional to A. astaci, crayfish can be infected with A. reptans (saprophytic lineage) and A. frigidophilus (animal parasitic lineage) (Ballesteros et al., 2006; Diéguez-Uribeondo et al., 2009; Royo et al., 2004). A. astaci apparently originated in North America and, through anthropogenic movements of the signal crayfish Pascifastacus leniusculus and others, is now established in many European countries following its first report in Europe in the 1860s (Alderman et al., 1984, 1990; Alderman, 1993; Bohman et al., 2006; Demers and Reynolds, 2002; Diéguez-Uribeondo et al., 1997; Diéguez-Uribeondo, 2005; Edgerton et al., 2004; Harlioglu, 2008; Johnsen et al., 2007; Kozubiková et al., 2006, 2007; Nylund and Westman, 2000; Rahe and Soylu, 1989). Although North American hosts are resistant carriers, native species of crayfish in Europe are highly susceptible with large scale mortalities associated with the fungal infection occurring following non-indigenous crayfish introductions. There are at least two molecular clades of A. astaci based on randomly amplified polymorphic DNA (RAPD) analysis (Huang et al., 1994; Lilley et al., 1997). Similar to most animal parasitic Aphanomyces spp., A. astaci does not produce sexual structures, with transmission occurring via zoospores which are released from infected moribund or dead crayfish into the water. These are motile and can survive for several days in water and several weeks in mud (Alderman and Polglase, 1988; Diéguez-Uribeondo et al., 1995; Edgerton et al., 2002a; Oidtmann et al., 2002). Following chemotactic attraction of the zoospores to crayfish, zoospores penetrate through the cuticle and infect the host; transmission to crayfish can be enhanced through damaged cuticle (Nyhlén and Unestam, 1975; Smith and Söderhäll, 1986; Unestam and Weiss, 1970). There are limited pathognomonic signs of infection, characterised by whitening of muscle; however, this can be indicative of other diseases. At the point of invasion by zoospores there may be small pin prick melanised areas (Alderman and Polglase, 1988; Edgerton et al., 2002a). Furthermore, histological lesions are also limited, in part due to the rapid onset of death following initial infection. The presence of hyphae in the cuticle at the site of infection is an important diagnostic feature of the disease. Recent advances in molecular biology techniques have facilitated development of a PCR approach to diagnosis of crayfish plague (Oidtmann et al., 2004, 2006). However, Ballesteros et al. (2009) showed that these primers lack specificity when used against other species of Aphanomyces isolated from crayfish. Any test used to identify a potential plague outbreak should thus include culture of the pathogen and molecular tools to confirm the presence of A. astaci (Vrålstad et al., 2009). One technique with potential for rapid, reliable identification of A. astaci is through

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amplification of chitinase genes which appear to be conserved in the fungus and may be species specific (Hochwimmer et al., 2009). 1.3.2. Class Sordariomycetes Several fungi within the class Sordariomycetes have been reported in crayfish under the genus name Fusarium, although sometimes these have not been typed to species (Quaglio et al., 2006a,b). Members of the genus are filamentous fungi which are normally distributed in soil and plants, in which they can cause disease. Alderman and Polglase (1985) described an infection of A. pallipes in the United Kingdom associated with Plectosporium (=Fusarium) tabacinum which is a plant pathogen and has been isolated from soil (Palm et al., 1995). Moulting of affected crayfish was delayed and the gills contained numerous melanised encapsulations, which gave rise to the common name of ‘‘black gill disease”. Histologically, melanised encapsulations contained fungal mycelia and were surrounded by hyaline haemocytes. In later stages of infection, increased melanisation of haemocytes developed. Alderman and Polglase (1985) suggested that early reports of fungal shell disease lesions caused by Didymaria cambari (Class Dothideomycetes) may have been due to P. tabacinum (attributed to Ramularia astaci (Class Dothideomycetes) by Alderman and Polglase (1988) and Edgerton et al. (2002a)). Lesions due to F. oxysporum were reported from the gills of Astacus leptodactylus and A. pallipes by Maestracci and Vey (1987), which superficially appear similar to those reported for P. tabacinum by Alderman and Polglase (1985). A further ‘‘Fusarium” infection of crayfish is that caused by Haematonectria haematococca, reported as Fusarium solani causing ‘‘brown abdomen disease” (Chinain and Vey, 1987). Pathology associated with this fungus is similar to that reported for P. tabacinum and includes haemocyte infiltration and subsequent melanisation in areas where fungal hyphae occur. Production of exotoxins by the fungus may be responsible for its lethality to crayfish (Chinain and Vey, 1988). Edgerton et al. (2002a) suggested that the Fusarium sp. in A. leptodactylus reported by Vey (1979) was F. roseum var. culmorum although there appears to be no basis for this decision. 1.3.3. Microsporidia Traditionally classified as protistans, the microsporidians have been placed either as a sister group to or firmly within the fungi (Fischer and Palmer, 2005; Gill and Fast, 2006; Hirt et al., 1999). Vossbrinck and Debrunner-Vossbrinck (2005) attempted to classify the Microsporidia into three new classes, the Aquasporidia, the Marinosporidia and the Terresporidia which reflected their habitat. However, they were unable to resolve the position of the two crayfish Thelohania spp. (Thelohania contenjeani and Thelohania parastaci) included in the analysis. 1.3.3.1. Thelohania spp. Thelohania species in crayfish are generally found within the musculature with infected animals generally appearing opaque or whitish giving rise to the common name of porcelain disease or cotton tail. There is currently debate on the phylogenetic position of the genus; Thelohania spp. occurring in freshwater crayfish occur within a single clade closely related to the Vairimorpha and Nosema clade from insect and crustacean hosts. Being distant from the type species of the genus, it is possible that crustacean Thelohania spp. may be transferred to a different genus (Brown and Adamson, 2006). The first Thelohania to be described from a crayfish was Thelohania contejeani Henneguy, 1892 and has been extensively studied. The parasite caused mass mortalities of crayfish in Europe; it infects A. astacus, A. leptodactylus, A. pallipes, P. leniusculus and Orconectes (Faxonius) limosus (Dunn et al., 2009; Edgerton et al., 2002a). T. contejeani has a dimorphic life cycle in the crayfish host, alternating between a diplokaryotic sporont producing 8 uninucleate spores measuring 4  2 lm with 9–10 turns of the polar tube,

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and single diplokaryotic sporonts producing diplokaryotic spores measuring 4  2 lm with 5–6 turns of the polar tube (Lom et al., 2001). Goodrich (1956) reported mortality of A. pallipes apparently caused by T. contejeani. She described two forms of microsporidians in muscle – one forming octosporous pansporoblasts typical of T. contejeani and a second form consisting of solitary spores which she referred to as Nosema sp. Edgerton et al. (2002a) considered these latter to be Ameson sp. with monosporous pansporoblasts. However, Goodrich (1956) had likely observed solitary T. contejeani spores arising from diplokaryotic sporoblasts as described by Lom et al. (2001). The report of this species in New Zealand Paranephrops zealandicus by Quilter (1976) likely refers to an undescribed species since all other records of this species are from the northern hemisphere. Thelohania spp. reported from Australia are found in crayfish within the superfamily Parastacoidea whereas T. contejeani is a parasite of crayfish from the superfamily Astacoidea (Moodie et al., 2003a). Spore measurements of T. contejeani from Quilter (1976) were smaller than those of Lom et al. (2001). Confirmation of P. zealandicus as a host for T. contejeani therefore needs both molecular and ultrastructural analyses. Additionally, the reports by France and Graham (1985) and by Graham and France (1986) of T. contejeani in Canadian crayfish Orconectes virilis need confirmation through use of molecular and ultrastructural methods. Two Thelohania spp. have been described from muscle of Australian C. destructor. Thelohania montirivulorum produces uninucleate and binucleate spores which measure 6  2.5 lm, with 20–22 turns to the polar filament (Moodie et al., 2003a). T. parastaci described from Cherax destructor albidus, C. d. rotundus and C. d. destructor undergoes a dimorphic life cycle involving either production of diplokaryotic spores, or production of sporophorous vesicles each containing eight uninucleate spores (Moodie et al., 2003b). Binucleate spores measured 4  2 lm and contain a polar filament with 6–8 turns. Uninucleate spores are of a similar size with 12–20 turns to the polar filament. Numerous other Thelohania species have been inadequately described, without molecular or ultrastructural data. These include Thelohania cambari from Cambarus bartoni that measured 4.6  2 lm (Sprague, 1950); Thelohania sp. from Cambarellus shufeldi (Sogandares-Bernal, 1962b, 1965) with spores measuring 3.0–3.5  1.2–1.6 lm (reported erroneously as T. cambari by Edgerton et al., 2002a); Thelohania sp. from the antipodean Paranephrops planifrons which measured 4  2 lm (Jones, 1980); Thelohania sp. in C. albidus from Australia measuring 4  2 lm (Jones and Lawrence, 2001) and Thelohania sp. in C. quadricarinatus measuring 3.5  2 lm. The latter species stopped spawning of females and caused infected animals to appear sluggish with a weak tailflick response (Herbert, 1987, 1988).

1.3.3.2. Pleistophora soganderesi. Originally reported as a Plistophora (sic.) sp. in Cambarellus puer by Sogandares-Bernal (1962b), the pansporoblast of this microsporidian contained 19–21 spores measuring 6–9  4 lm. It was incorrectly referred to as Thelohania soganderseri by Edgerton et al. (2002a), having been named P. soganderesi by Sprague (1966). The infection was apparently rare in crayfish and was absent from a subsequent survey of crayfish in Louisiana by Sogandares-Bernal (1965). This microsporidian and Thelohania sp. of Sogandares-Bernal (1962b) may have caused earlier mortality events in Louisiana. The placing of this microsporidian in the genus Pleistophora may be inappropriate since members of this genus are parasites of vertebrates (Canning and Hazard, 1982; Nilsen et al., 1998; Nilsen, 2000). There is therefore a need to re-evaluate the taxonomic position of this microsporidian through molecular and ultrastructural studies.

1.3.3.3. Vavraia parastacida. Members of this genus typically have a variable number of spores in the pansporoblast ranging from 8, 16, 32 and rarely 64. Vavraia parastacida has been reported from Cherax tenuimanus, C. albidus, Cherax quinquecarinatus and C. quadricarinatus (Langdon, 1991a, 1991b; Langdon and Thorne, 1992). Infected animals apparently have a bluish colouration, particularly lateral and ventral to the tail. Similar to Thelohania sp. reported by Herbert (1987), infected animals are sluggish with limited tail-flick response. Spores measure 5.5  2.6 lm and contain a polar filament with 9–11 turns. 1.3.3.4. Vairimorpha cheracis. This microsporidian infects mainly the muscular tissues of C. destructor and is also found within the intestinal mucosa and the intercellular spaces surrounding the ova in females (Moodie et al., 2003c). Spores are markedly pyriform and measure 3.5  2 lm. Similar to other Microsporidia infections of crayfish muscle, infected animals appear opaque. Moodie et al. (2003c) reported that infected animals do not feed, move abnormally slowly and are likely to die if they moult. However, infected animals held in aquaria lived for up to six months, suggesting that disease progression was slow. 1.3.3.5. Unidentified Microsporidia. A microsporidian infection with no obvious external signs in two C. quadricarinatus was reported by Edgerton and Owens (1999) in which animals were co-infected with CqBV. Microsporidian spores were noted in the connective tissue, typically near the hepatopancreas with an intense haemocytic and melanistic response associated with the spores. The authors could not determine the number of spores in sporophorous vesicles because only sectioned material was available. Spores in section measured 4  2.5 lm and the authors considered that given the tissue tropism and size, it may represent an undescribed species. 1.4. Mesomycetozoea The Mesomycetozoea was erected to incorporate a monophyletic group of organisms at the animal-fungi border that included Dermocystidium, rosette agent, Ichthyophonus and Psorospermium and have been referred to as the DRIPs clade after those four original members (Ragan et al., 1996). Two orders have been described within the Mesomycetozoea, the Ichthyophonida and the Dermocystida. The crayfish parasite Psorospermium spp. was placed in the Ichthyophonida along with Amoebidium parasiticum, Anurofeca richardsi, Ichthyophonus spp., Pseudoperkinsus tapetis, Sphaeroforma arctica, isolate LKM51 and isolate Ikaite un-c53 (Adl et al., 2005; Mendoza et al., 2002). Psorospermium spp. affect a wide range of crayfish in the families Astacidae, Cambaridae and Parastacidae (Vogt and Rug, 1999). Under certain conditions the genus is pathogenic (Vey, 1986). Several Psorospermium morphotypes have been described, based on size, morphology, host and geographical location (Bangyeekhun et al., 2001). Two European morphotypes are differentiated by size. Psorospermium haeckeli is elongate, slightly spindle shaped with roundish or lance-like ends and measures 120–180  50  50 lm, the other type being egg-shaped and measuring 100  60  60 lm (Vogt and Rug, 1995). Both types cooccur in several crayfish species (Henttonen et al., 1997; Vogt et al., 1996; Vogt and Rug, 1995, 1999). The morphotype from P. clarkii and P. zonangulus from the USA measures 120–200  35–70 lm (Bangyeekhun et al., 2001; Henttonen et al., 1992). Despite molecular studies confirming the clear morphological distinction between the European and American forms, Bangyeekhun et al. (2001) were unwilling to erect a new species in the absence of a complete description of all life stages of each American and European morphotype. Jones and Lawrence (2001) reported a Psorospermium sp. in the gills, connective and neural tissues of C. albidus with negligible host

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response, in Australia. Herbert (1987) reported Psorospermium sp. in the same organs and tissues of C. quadricarinatus and occasionally ovary membranes, cardiac and skeletal muscle with no pathology. There was evidence of accumulation of increased numbers with increasing age of the crayfish. In contrast, Edgerton and Owens (1999) found the infection in the connective tissue of the subcutis, hepatopancreas, antennal gland and gill, and rarely in skeletal muscle, and neural and haemopoietic tissues C. quadricarinatus in Australia. However, they found no evidence of increasing intensity of infection with age. Psorospermium sp. in C. quadricarinatus and C. tenuimanus possesses two shell plates, unlike P. haeckeli which possess multiple plates in mature spores (Edgerton and Owens, 1999). Psorospermium spp. may have a diphasic life cycle, and although a putative intermediate host has not been demonstrated, it can be transmitted from P. leniusculus to A. astacus and between P. leniusculus individuals but not from A. astacus to P. leniusculus (Gydemo, 1996). However, Vogt et al. (1996) suggested that Psorospermium was transmissible from A. astacus to Austropotamobius torrentium. A free living stage without the need for an intermediate host may exist (Vogt and Rug, 1999). 1.5. Protista 1.5.1. Ciliata Although ciliates are commonly associated with crayfish, they are normally not considered a problem in the wild. Mortalities generally occur under aquaculture conditions where poorer water quality, elevated temperatures and high host densities increase the risk of problems (Morado, 1995). Life cycles are direct and reproduction is normally through binary fission or budding. Most ciliates are found on the external surfaces of crayfish, including pleopods, periopods, telsons, gills and carapace. Host-parasite checklists of ciliates on crustaceans are provided by Sprague and Couch (1971) and Morado (1995). Tetrahymena pyriformis is a complex of approximately 30 species of holotrich ciliates that occur both as free-living organisms and as opportunistic parasites of both vertebrate and invertebrate hosts, including fish (Sadler and Brunk, 1992; Simon et al., 2008). A systemic infection of three red clawed crayfish from northern Queensland, Australia was caused T. pyriformis, based on morphological characteristics of the parasite. Affected animals exhibited weakened or failed tail-flick response and were unable to right themselves. Histologically, the ciliates occurred usually in the haemocoel and haemal spaces in most tissues. Focal necrosis of the tissues occurred in all affected crayfish (Edgerton et al., 1996a). The other major group of ciliates affecting crayfish are in the order Sessilina, whose defining characteristic are that they are attached permanently to the host (Morado, 1995). Genera occurring on crayfish include Epistylis, Carchesium, Lagenophrys, Paralagenophrys, Zoothamnuim, Opercularia, Vorticella and Cothurnia. Most reports related to the peritrichous ciliates Episytlis spp. suggest they are innocuous, acting as commensals (Brown et al., 1993; Harlioglu, 1999; Hüseyin and Selcuk, 2005; Quaglio et al., 2006b; Vogelbein and Thune, 1988). However, mortalities have been associated with Epistylis sp., usually under culture conditions (Brown et al., 1993) and mortalities associated with Cothurnia in Italian crayfish was reported by Ninni (1864). Other lesser known ciliates of crayfish include the genera Podophyra (Order Exogenida), Acineta and Tokophyra (Order Endogenida), Discophyra (Order Evaginogenida) and Hyalophysa (Order Apostomatida) (Grimes, 1976; Morado, 1995; Romero and Jiménez, 2002; Vogelbein and Thune, 1988).\ 1.5.2. Phylum Apicomplexa Apicomplexans are pathogens of a wide range of animal hosts, including fish and shellfish (Carballal et al., 2001; Gestal et al.,

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2007; Steinhagen et al., 1997). In brief, the life cycle of members of the family Eimeriidae is divided into merogony in which the parasite invades the host cell and undergoes a transformation into a meront in which merozoites develop. This is followed by gamogony in which microgamonts and macrogamonts are formed, fertilization occurs and the zygote develops into an oocyst which contains sporocysts and sporozoites. Some coccidians have a two-host life cycle (Steinhagen, 1991; Steinhagen and Körting, 1990). To date two coccidians have been reported from crayfish. The eimerian coccidian Mantonella (=Yakimovella) potamobii described from the intestinal contents of A. leptodactylus, is characterised by oocysts with a single sporocyst which in turn contains four sporozoites (Gousseff, 1936). No pathology was described with this parasite. A second putative coccidian was noted in the ovaries of a single female marbled crayfish by Vogt et al. (2004). The parasite was tentatively considered to be a member of the family Eimeriidae with numerous unicellular stages present, with merogonous stages and putative sporogonic stages. Granulomas occurred in the hepatopancreas and hypodermal connective tissues of the infected animal although it is not clear if these were caused by the coccidian. Vogt et al. (2004) considered that the infection was lethal to crayfish. 1.6. Digenea Digenea have complex life cycles utilising at least two hosts, with adult stages normally occurring in vertebrates; metacercarial stages occur in crayfish, although several members of the genus Alloglossoides occur as adults in crayfish (Turner, 1984b). To date Digenea have been reported in crayfish from Europe, Asia, Australia and the USA. Digenea occur in most tissues within crayfish although this is dependent on the genus and species of digenean under investigation. Only two papers have described the pathology associated with digeneans in crayfish, both within the antennal gland of their hosts (Turner, 1984b, 1985). This perhaps reflects the emphasis on faunal surveys rather than lack of pathology associated with these parasites. 1.6.1. Family Allocreadiidae The life cycle of Crepidostomum cornutum was elucidated by Ameel (1937) who showed that both mother and daughter rediae occur in the freshwater bivalve Sphaerium sp. Free swimming cercariae released from the bivalve penetrates the host and encyst in the cardiac region, hepatopancreas and muscles of the cephalothorax of the crayfish host (Ameel, 1937; Sogandares-Bernal, 1965). Time from penetration to encystment is <2 h with metacercariae in crayfish attaining full size in 6–8 weeks (Ameel, 1937). Progenesis can occur in C. cornutum in crayfish, although the mechanisms for this are not understood (Sogandares-Bernal, 1965). Furthermore, eggs produced via this route are not viable, being uninfective to the clam host (Cheng, 1957; Lefebvre and Poulin, 2005). Transmission to the fish definitive host occurs following ingestion of infected crayfish. Sogandares-Bernal (1965) considered that C. cornutum was the most prevalent digenean in Louisiana crayfish, occurring in at least six species of crayfish; host specificity in the teleost host also appears to be very wide. Despite the apparent widespread nature of this parasite, its wide host range and lack of organ specificity, no studies have been conducted on pathogenesis of infection. 1.6.2. Family Choanocotylidae A new genus, Choanocotyle, was erected in a new family (Choanocotylidae) by Jue Sue and Platt (1998) to include two plagiorchiidan species from freshwater turtles collected near Brisbane, Australia. One species in the family, Choanocotyle elegans occurs in the small intestine of the freshwater turtles Chelodina expansa and Emydura macquarii. It uses the planorbid snail, Glyptophysa

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gibbosa as first intermediate host. Metacercariae were found at low prevalence in naturally infected Cherax sp. 1.6.3. Family Cladorchidae Cercariae of Allassostomoides parvus (=parvum) encyst on the outer surfaces of crayfish species after release from the snail Planorbis trivolvis. The parasite develops in frogs and turtles following ingestion of parasitised crayfish (Beaver, 1929; Brooks, 1975). No pathology is associated with this infection; crayfish are not more susceptible to predation if infected with this parasite. 1.6.4. Family Gorgoderidae Adults of Gorgodera amplicava develop within the urogenital system of anurans with infections acquired via ingestion of infected crayfish. The first intermediate host is Musculium partumeium. Sogandares-Bernal (1965) described this parasite in the stomach wall near the gastric mill of Orconectes palmeri creolanus and P. clarkii. No pathology was reported. 1.6.5. Family Haematoloechidae Morrison (1966) reported, without details, metacercariae of Haematoloechus sp. in crayfish from Texas, USA. The parasite utilises snails as first intermediate hosts and matures in the lungs of various anurans. 1.6.6. Family Macroderoididae Two genera within the family Macroderoididae have been reported from crayfish, Macroderoides and Alloglossidium, most species occurring in the latter genus. Following phylogenetic analysis of several genera within the family, Alloglossoides was relegated to a junior synonym of Alloglossidium by Smythe and Font (2001). Although digeneans typically utilise three hosts in their life cycle (e.g. snails, arthropods and vertebrates), some members of the genus Alloglosidium are unusual in that they have a two host life cycle with leeches or crayfish as definitive hosts (Smythe and Font, 2001). Those in crayfish all occur either encysted or unencysted in the antennal gland. Three species within Macroderoididae which infect crayfish follow the typical three host life cycle: Macroderoides typicus, Alloglossidium corti and A. progeneticum (=Macroderoides progeneticus). M. typicus, reported from the cephalothoracic and antennal musculature of Procambarus blandingi acutus and P. clarkii in Louisiana, USA by Sogandares-Bernal (1965) utilises the snail Helisoma trivolvis lentum as first intermediate host, tadpoles and crayfish as second intermediate hosts, and the bowfin Amia calva as definitive host. Similarly, Heliosoma trivolvis has been recorded as the first intermediate host for A. corti, with crayfish and other invertebrates as second intermediate hosts and fish of the genera Noturus, Ictalurus and Micropterus as definitive hosts (Crawford, 1937; McMullen, 1935; Muzzall and Pracheil, 2007; Smythe and Font, 2001; Spall and Summerfelt, 1969). A. progeneticum described by Sullivan and Heard (1969) encysts on the antennary gland of Procambarus spiculifer and uses Ictalurus nebulosus as the final host (Font and Corkum, 1975; Sullivan and Heard, 1969). No disease or pathology has been reported for any of these species. Those members of the family which are suggested to follow a two host life cycle with crayfish as definitive host include Alloglossidium greeri, Alloglossidium cardicolum (=Alloglossoides cardicola) and Alloglossidium (=Alloglossoides) dolandi. A. greeri found unencysted in the antennary gland of Cambarellus shufeldtii has been described by Font (1994); its full life cycle has not yet been elucidated. Alloglossidium caridicolum occurs as an unencysted adult in the antennal gland of Procambarus acutus from Louisiana, Mississippi, Arkansas and Texas in the USA and shows strong host specificity to this crayfish (Corkum and Turner, 1977; Turner, 1999, 2007). Its complete life cycle has not yet been fully elucidated

but is likely to involve a snail host (Turner, 2000). The clear seasonality of infection in P. acutus provides evidence for a second host (Turner, 2000). The parasite causes localised damage within the antennal gland, using its tegumental spines to abrade the tubule epithelium and feeds on the abraded tissue (Turner, 1985). Nodules associated with these abraded areas and feeding by the parasite leads to loss of tissue in the nephridial tubule. In contrast to A. caridicolum, A. dolandi has wide host specificity, occurring in 6 species of Procambarus, including its type host Procambarus epicyrtus (Turner, 2007; Turner and McKeever, 1993). Although it occurs sympatrically with A. caridicolum, it occurs unencysted in the lumen of the labyrinth portion of the antennal gland (Turner and McKeever, 1993). Edgerton et al. (2002a) erroneously listed crayfish as a host for the freshwater shrimp specific parasite, Alloglossidium renale described by Font and Corkum (1975). 1.6.7. Family Microphallidae Attempts to resolve the convoluted taxonomy of the family were made by Deblock (1973). Sogandares-Bernal (1965) reported Maritrema (Atriospinosum) obstipum in the central shaft of the gill filaments and hepatopancreas of C. shufeldti and P. clarkii. Although Etges (1953) suggested that this parasite utilised the gastropod Amnicola pilsbryi, the isopod Asellus communis and various bird and mammalian hosts in its life cycle, it is possible that M. obstipum represents a group of cryptic species. Overstreet et al. (1992) described Microphallus fonti from the hepatopancreas of P. clarkii from Louisiana. They considered that the M. opacus from the hepatopancreas of C. puer and P. clarkii of Sogandares-Bernal (1965) was conspecific with M. fonti. Furthermore, there is a contention that M. opacus may be conspecific with M. ovatus (Caveny and Etges, 1971). Therefore, there is a need to reevaluate the taxonomy of members of this genus. M. opacus and M. ovatus have been recorded in the hepatopancreas of Cambarus sp. by Stafford (1931) and Osborn (1919) respectively. Microphallus minutus has been recorded in Cherax dispar and C. destructor from Australia (Shimazu and Pearson, 1991). Sheldon (1938) confirmed that Quasimaritremopsis medius (=Microphallus medius = Maritrema medium = Maritreminoides medium) utilised Orconectes (=Cambarus) virilis and C. propinquus in its life cycle but was unable to ascertain the definitive host for this parasite which occurs specifically within crayfish gill filaments. The progenetic digenean Sogandaritrema progeneticum (=M. progeneticus) was originally described from the cephalothoracic cavity in C. puer by Sogandares-Bernal (1962a) and has also been noted in C. shufeldtii and P. clarkii. Its life cycle was elucidated by Lotz and Corkum (1983) who showed that the parasite produced infective cercariae in the gastropod Amnicola peracuta. 1.6.8. Family Opecoelidae Utilising the prosobranch snail Posticobia brazieri as the first intermediate host, Opecoelus variabilis infects the Australian crayfish C. depressus and C. dispar and other freshwater crustaceans. Following ingestion of infected crayfish, the parasite develops into adults in the intestine of at least 17 species of freshwater fish (Cribb, 1985). No disease in crayfish has been reported for this species. 1.6.9. Family Orchipedidae Alderman and Polglase (1988) suggest that Orchipedium (=Distoma) isostomata is found in all species of European crayfish. It occurs in most tissues, including hepatopancreas, near nerve cords and gonads, heart, muscle and free among other organs. Despite the apparent ubiquity of this species in Europe, its life cycle has not been fully elucidated.

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1.6.10. Family Paragonimidae Few Digenea of crayfish have been reported as being of zoonotic concern. One of the more important digeneans in this respect are those in the genus Paragonimus spp., in particular Paragonimus westermani in Asia, Africa and some South American countries and P. kellicotti in North America (Liu et al., 2008). Infections in humans are normally characterised by fever and coughing. However, few cases of P. kellicotti (or P. westermani) have been reported in the USA, in part due to the limited ingestion of raw crayfish there. Paragonimus spp., in common with other digeneans, use a snail as the first intermediate host, a crustacean as the second intermediate host and mature to adults in a mammal (Healy, 1970). In the USA, several Cambarus spp., Procambarus blandingi acutus and P. clarkii have been reported as hosts for P. kellicotti with infections occurring in the heart and surrounding tissues and in the musculature (Sogandares-Bernal, 1965). P. westermani infections in Asia have been well recognised for several years with Cambaroides similis being important hosts for the parasite in Korea (Hong et al., 1986; Kim et al., 2009; Park, 1962; Shin and Min, 1999). Chai et al. (1996) suggest that despite loss of suitable habitat for crayfish in Korea, the incidence of P. westermani in crayfish remains high. 1.6.11. Superfamily Plagiorchioidea The monotypic genus Allocorrigia was erected in the family Dicrocoeliidae by Turner and Corkum (1977) for digenean parasites of the antennal gland of crayfish. Subsequently, Pojmanska et al. (2008) considered that Allocorrigia was a genus incertae sedis within the superfamily Plagiorchioidea. Allocorrigia filiformis from P. clarkii matures within the antennal gland and releases eggs containing active miracidia via the excretory gland (Turner and Corkum, 1977). The remainder of its life cycle remains unknown and the parasite appears to be host specific to P. clarkii (Turner, 1984a, 2004, 2006). Unlike the macroderoidid A. caridicolum which abrades the antennal gland tubule epithelium causing localised pathology, adults of A. filiformis do not appear to elicit a host response (Turner, 1984b, 1985). However, eggs released from the worm are sometime trapped in interstices below the epithelium leading to the production of melanised nodules containing ova. 1.6.12. Family Psilostomatidae Warren (1903) described the life cycle of Astacotrema cirrigerum from the musculature of A. astacus and whilst he did not provide an adequate generic description of the parasite, considered that it should be placed in the Family Psilostomatidae. However, it is highly probable that the detailed study involved a composite of two species (one being A. cirrigerum sensu stricto and the other being O. isostomata from the family Orchipedidae). As a result of a number of discrepancies, Kostadinova (2005) considers that A. cirrigerum should not be placed in the Family Psilostomatidae and regard it as a genus incertae sedis. Astacatrematula macrocotyla encysts as metacercaria externally on the sternites and gills of Astacus trowbridgi from North America. The parasite uses the psorobranch gastropod, Flumenicola virens, as a first intermediate host and was transmitted experimentally from crayfish to birds by Macy and Bell (1968) who provide a full description of all life stages of the digenean. Spatio-temporal variations in prevalence and intensity of this parasite on crayfish were attributed to absence of a suitable final host by Macy and Bell (1968). 1.6.13. Family Reniferidae Renifer (=Ochetosoma) sp. occurs within the musculature of P. clarkii in Lousiana and completes its life cycle in snakes (Sogandares-Bernal, 1965). No further records of this parasite in

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crayfish have been made and no assessment of its potential impact in the host is available. 1.6.14. Family Troglotrematidae The life cycle of Macroorchis spinulosus was experimentally completed by Chai et al. (1996) who also redescribed the adult and metacercarial forms of the parasite. First reported in freshwater crabs, Chai et al. (1996) noted the parasite within the muscle of C. similis. The authors considered that it was theoretically possible for zoonotic infections to occur following ingestion of raw crayfish in areas endemic for the parasite. 1.7. Cestoda To date there have been few records of tapeworms within crayfish. O’Donoghue et al. (1990) reported unsuccessful attempts to feed metacestodes of Vampirolepis diminuta (Family Hymenolepididae) found in the intestinal mucosa of C. destructor to laboratory rats. Reports of the cysticercoids of Hymenolepis collaris and Hymenolepis tenuirostris (Family Hymenolepididae) in the body cavity of European crayfish were considered incorrect by Alderman and Polglase (1988) and Edgerton et al. (2002a). Hymenolepis species mature in the intestine of ducks and other birds. The only other cestode reported from crayfish is Austramphilina elongata. The parasite belongs to the order Amphilinidea, a group of monozoic cestodes which are primitive to the eucestodes. Generally, Amphilinidea infect primitive freshwater fish, freshwater turtles and more advanced marine teleosts. Eggs of A. elongata are released from adults in the coelom of turtles and following maturation, infective larvae are released (Rohde and Georgi, 1983). These larvae penetrate the cuticle of the crayfish C. destructor and migrate through to the abdominal muscle near the anus (Rohde and Watson, 1989). Transmission occurs when infected crayfish are ingested by turtle definitive hosts. Despite obvious penetration of the parasite through the cuticle and inherent damage caused by this, no data exists on the potential pathology associated with this mode of entry. 1.8. Acanthocephala Acanthocephalans are obligate parasites with complex life cycles generally involving at least two hosts, usually an invertebrate and a vertebrate. Typically, the invertebrate host is an insect or a crustacean, while the vertebrate hosts can include mammals, birds and fish. There is some controversy regarding their phylogenetic position and although some workers have considered them to be sister taxa to the Rotifera (Near, 2002), others have expressed caution (Nickol, 2006). To date only five species of acanthocephalans have been reported from crayfish, including larval stages of Polymorphus biziurae in C. destructor from Australia (Johnston and Edmonds, 1948; O’Donoghue et al., 1990), P. minutus (=boschadia) in Orconectes (Faxonius) limosus (originally reported as Cambarus affinis) by Golvan (1961), Fillicollis anatis in A. astacus by Golvan (1961), Southwellina dimorpha in P. clarkii and C. shufeldtii from the USA (Richardson and Font, 2006; Schmidt, 1973) and Neoechinorhynchus rutili in Pacifastacus trowbridgi from the USA (Merritt and Pratt, 1964). However, Merritt and Pratt (1964) considered that crayfish were a paratenic host for N. rutili since it normally utilises ostracods as an intermediate host. It should be noted that the listing of ‘‘crayfish” as a host for Polymorphus formosus by Richardson and Font (2006) is incorrect as the original report by Schmidt and Kuntz (1967) used the term ‘‘crayfish” in relation to the freshwater shrimp Macrobrachium sp. All acanthocephalans so far reported in crayfish are normally found encysted on the outside of the intestine and no pathological changes have been reported in crayfish infected by Acanthocephala (Schmidt, 1973).

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1.9. Nematoda Parasitic nematodes have complex life cycles involving several hosts, normally consisting of invertebrates and vertebrates as intermediate hosts and vertebrate definitive hosts in which sexual reproduction takes place. Although several ‘‘external” nematodes have been reported on crayfish, these can be considered to be free-living epibionts with limited impact on host survival (Edgerton et al., 2002a; Jones and Lawrence, 2001; Schneider, 1932). Truly parasitic nematodes are normally found internalised within crayfish, either encysted in the muscle or encapsulated on the intestinal wall; to date few parasitic nematodes have been reported in crayfish, with crayfish acting as paratenic hosts in all cases. These include the human pathogen Gnathostoma spinigerum in Japan and the rat lungworm, Angiostrongylus cantonensis, using Cambarus sp. from the USA (Miyazaki, 1954; Moravec, 2007; Rachford, 1975). The giant kidney worm Dioctophyme (=Dioctophyma) renale infects a wide range of fish-eating mammals, including humans, throughout the world (Dyer, 1998; Measures and Anderson, 1985). The first intermediate host of the parasite is the annelid ‘‘Lumbriculus variegatus” which was recently shown to be a species complex by Gustafsson et al. (2009); several fish species act either as intermediate or paratenic hosts (Mace and Anderson, 1975; Measures and Anderson, 1985). However, confusion has arisen regarding the role of crayfish in the transmission of this nematode due in part to erroneous reporting of the branchiobdellid Cambarincola philadelphica on the gills of crayfish as the first intermediate host (Myers, 1970; Olsen, 1974; Woodhead, 1950). Larvae identified by Woodhead (1950) were likely horsehair worms (Gordiacea) and therefore crayfish should not be considered as hosts for this parasite (Mace and Anderson, 1975). Attempts to experimentally transmit Dracunculus insignis to the crayfish Orconectes propinquus from infected copepods were unsuccessful leading the authors to suggest that crayfish may not be a suitable host for this nematode (Crichton and Beverley-Burton, 1977). Quaglio et al. (2006b) reported unidentified nematodes encysted in the intestines of P. clarkii but were unable to ascertain the role of crayfish in the life cycle of the parasite; no pathology was reported for this infection. 1.10. Branchiobdellida Branchiobdellids are annelids which are generally considered to be ectocommensals or ectosymbionts of crayfish. There has been some debate regarding their taxonomic position and although they are clearly members of the Clitellata, have at various times been assigned to the Oligochaeta, Hirudinea and as an independent taxon equal in rank to both the oligochaetes and the leeches (Apakupakul et al., 1999; Gelder and Siddall, 2001). Through use of several methodologies, including morphology, molecular tools and sperm characteristics, it is apparent that branchiobdellids are derived oligochaetes (Erséus et al., 2008). An estimated 150 species of branchiobdellids in 21 genera exist, with most species in the genera Cambarincola and Branchiobdella (see Gelder, 1996). Branchiobdellids appear to be restricted to the northern hemisphere, occurring in central and North America, Europe and Asia, but not apparently in Australia where the niche is partly occupied by temnocephalids (Alderman and Polglase, 1988; Edgerton et al., 2002a). Overall, the genus Branchiobdella is restricted to Europe and the genus Cambarincola to the USA (Duris et al., 2006; Füreder et al., 2009; Holt, 1973; Holt and Opell, 1993; Klobucar et al., 2006; Oberkofler et al., 2002; Quaglio et al., 2006a; Rogers et al., 2003; Subchev et al., 2007; Timm, 2005; Williams et al., 2009). However, subsequent to anthropogenic movements of crayfish, there are an increasing number of reports of exotic species of branchiobdellids

throughout their range (Gelder et al., 1994, 1999, 2002; Ohtaka et al., 2005). Branchiobdellids occur externally on the host, including gills and carapace, although their site specificity can vary depending on season and presence of other species of branchiobdellids (Edgerton et al., 2002a). Life cycles of branchiobdellids are poorly understood with cocoons being laid on the external surfaces of the crayfish host where development of young branchiobdellids takes place. Transmission of branchiobdellids between crayfish is likely through host to host contact. There have been suggestions that branchiobdellids are pathogenic to their host; some species occurring in the gill chambers elicit a host response such as melanisation at the site of attachment and grazing (Alderman and Polglase, 1988). However, there has only been one report of a mortality allegedly associated with excessive numbers of branchiobdellids; dead animals were also found to have high levels of bacteria and other pathogens which are more likely to be the primary cause of the mortality (Hubault, 1935). Branchiobdellids engage in cleaning symbiosis with the crayfish host, removing fouling organisms and thus improving growth rates of the host (Brown et al., 2002; Keller, 1992; Lee et al., 2009). 1.11. Temnocephalida Temnocephalids are ectosymbiotic rhabdocoeles in the phylum Platyhelminthes, occurring mainly on crayfish, and also on other freshwater crustaceans, molluscans, insects and chelonians (Amato et al., 2003, 2006; Damborenea and Brusa, 2008, 2009; Jones and Lester, 1993; Volonterio, 2007; Williams, 1994). They are characterised by the presence of eyespots, a posterior sucker and usually 5, 6 or 12 digitate processes on the anterior end of the body (Edgerton et al., 2002a; Sewell and Whittington, 1995). The anterior processes and the posterior sucker are predominately used in attachment to and locomotion around the host (Sewell and Whittington, 1995). Temnocephalids are predominantly found in the southern hemisphere although they can occur in the northern hemisphere, usually resulting from anthropogenic movements of the crayfish host (Gelder, 1999; Volonterio, 2009). Typically, temnocephalids have a direct life cycle, depositing eggs on the host carapace. Following development in the eggs, ciliated juveniles hatch and remain attached to the host (Jones and Lester, 1992). They are able to leave the host and survive for a period; some species may be able to lay eggs off the host (Hickman, 1967; Jennings, 1971), although for others, this does not occur (Jones and Lester, 1992). Both juvenile and adult temnocephalids browse across the surface of the host feeding on other fouling organisms and on crayfish eggs (Jones and Lester, 1993). Temnocephala chilensis from a freshwater crab can act as an intermediate host for the digenean Echinoparyphium megacirrus; the possibility that crayfish temnocephalids can harbour digeneans is unconfirmed (Viozzi et al., 2009). Additionally, Rickettsia-like organisms have been reported in the testes of Temnocephala novaezealandiae and Troglocaridicola mrazeki with limited pathology and no apparent transmission to progeny of temnocephalids (Williams, 1991). Its relationship to the Rickettsia-like organisms identified as C. cheraxi in crayfish remains unknown. 1.12. Other fouling organisms Additional to harbouring pathogens, crayfish can act as hosts for fouling organisms which readily attach to most of their external surfaces where they generally do not elicit a host response or cause any problems for the host. This includes cyanobacteria and chlorophyta algae, corixid eggs, mites, ostracods, oligochaetes, polychaetes, rhabdocoel flatworms, rotifers, Argulus eggs, bryozoans and

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zebra mussels (Alderman and Polglase, 1988; Amato, 2001; Brazner and Jensen, 2000; Cannon and Jennings, 1987; Duris et al., 2006; Edgerton et al., 2002a; Lamanova, 1971; Romero and Jiménez, 2002). The latter three are probably most important with regards to potential disease transmission through the anthropogenic and natural movements of crayfish between water bodies. Argulus spp. are parasitic branchiurans of fish that are disease causing agents and have a minor role in the passive transmission of spring viraemia (SVC) of carp (Ahne, 1985); bryozoans are hosts for the myxozoan parasite Tetracapsuloides bryosalmonae, the causative agent of proliferative kidney disease of salmonids (Feist et al., 2001). Zebra mussels transmit several parasites to naïve hosts (Molloy et al., 1997; Mühlegger et al., 2009). The role of crayfish in dissemination and enhancement of disease transmission through epibionts is likely to be limited. 1.13. Idiopathic conditions Limited reports exist of idiopathic conditions of crayfish, in part because of a lack of clear case definitions, and sometimes because disease investigations of crayfish involve case reports of single infections. Thus idiopathic conditions are grossly underreported. Dexter (1954) described a cyst-like growth under the carapace of a single O. propiquus in the USA. The mass inside the cyst was described as rubber-like and whilst it was promulgated that it may have been due to an infectious agent, no evidence was apparent to support this proposition. Black to dark blue coloured spots have been reported on the exoskeleton of C. quadricarinatus from Australia and Ecuador and although they were not associated with poor health they rendered the crayfish unmarketable (Edgerton, 2000; Edgerton and Owens, 1999; Jiménez and Romero, 1997). Histopathology of crayfish affected by haemocytic enteritis was described by Edgerton (2000) and Edgerton and Owens (1999). Histologically, there was loss of epithelium of the midgut with concomitant thickening of the basement membrane. In addition, necrotic changes and rounding up of hepatopancreatocytes occasionally occurred in the hepatopancreas. Other idiopathic conditions noted in crayfish by Edgerton (2000) included needle shaped crystals in the nephridial canal and giant cells in the labyrinth epithelium. Nodules and granulomas were reported from the hepatopancreas of crayfish and other than some melanosis, no pathology was noted with this condition (Romero and Jiménez, 2002). Additionally, iron granules have been reported in the hepatopancreas of C. quadricarinatus from Ecuador (Romero and Jiménez, 2002).

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much emphasis was being placed on crayfish plague, little has changed. Thus actual causes for mortalities may go underreported and transmission of new and novel disease agents may not be recognised (Alderman and Polglase, 1988; Edgerton et al., 2002a, 2004; Edgerton, 2002; Edgerton and Jussila, 2004; Edgerton and Owens, 1999; Romero and Jiménez, 2002). Additionally, reliance on molecular methods may lead to underreporting of disease causes and provide erroneous results (Ballesteros et al., 2009; Claydon et al., 2004). Future studies on crayfish should consider an integrated approach to sampling that ensures that samples are taken from individual animals for electron microscopy, histopathology, classical taxonomy, molecular biology and so on. Only by utilising this integrated approach can disease experts hope to unequivocally identify pathogens of concern in crayfish and to understand the role of disease in structuring crayfish populations in conjunction with epidemiologists, population biologists and ecologists (Feist and Longshaw, 2008). The use of – omics technologies and microarrays may well prove useful in disease diagnosis but clearly need to be linked with confirmation of a pathological response to infection to unequivocally link the infection with disease. The loss of expertise in classical taxonomy and the lack of taxonomic rigour applied to some studies of crayfish disease should be addressed – it is critical that both novel and traditional approaches to understanding crayfish pathogens and their impact on overall biodiversity are utilised (Boero, 2010). Previously reported and poorly described pathogens should be revaluated using appropriate methods to confirm their identity and impact. Future studies should focus on the role of the underlying immune status of crayfish in response to synergistic and multiplicative factors, including disease agents which lead to lethal and sublethal effects being noted in crayfish (Cárdenas and Dankert, 1997; Cerenius et al., 2003; Jiravanichpaisal et al., 2004; Johansson and Söderhäll, 1985; Shi et al., 2005; Unestam and Weiss, 1970). This holistic approach may aid further understanding of methods to boost immune function or to mitigate against mortalities by removal of those intrinsic factors which lead to death or debilitation of the host. Few studies have been conducted on differences in disease status in different age cohorts of crayfish and this should be addressed. Finally, given the importance of crayfish in ecosystems, it seems somewhat counterintuitive that despite several studies on crayfish health, there remains a paucity of health data for many species of crayfish and for many crayfish populations. It is hoped that this review provides the impetus to continue looking at crayfish health and to finding and describing new parasites and diseases whilst mindful of the need to maintain taxonomic rigour and to consider the whole biota, not just one or two selected diseases.

2. Conclusions and future directions

References

Crayfish throughout their range contain a wide range of infectious and non-infectious agents that have the potential to cause mortalities in individuals and affect populations. Notwithstanding the focus on A. astaci, mortalities of crayfish appear to be associated mainly with bacteria, viruses, other fungi, protists and occasionally other ‘‘unexplained” reasons. In addition, mortalities due to underlying environmental conditions occur and the interplay between host, pathogen and environment need to be considered more fully (Bitner Anderson et al., 1997; Davies, 1989; France and Graham, 1985; Gil-Sánchez and Alba-Tercedor, 2006; Royo et al., 2002). The movement of crayfish for aquaculture purposes and, to a lesser extent the aquarium trade, has led to disease outbreaks and new geographical and host records for disease conditions. Unfortunately, despite repeated calls for incorporation of histopathology into studies of crayfish diseases and a concern that too

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