Perkinsus olseni of Abalone
Category 3 (Host Not in Canada)
Common, generally accepted names of the organism or disease agent
Perkinsus disease of abalone.
Scientific name or taxonomic affiliation
Widespread along the coast of South Australia. A species of Perkinsus, that is believed to be P. olseni, occurs in various molluscs from the Great Barrier Reef but was not detected in abalone from that area. Also, Perkinsus sp. has not been reported in molluscs from Tasmania (Goggin and Lester 1995). Molecular systematics demonstrated that P. olseni is a senior synonym of Perkinsus atlanticus in clams in Europe and eastern Asia (Murrell et al. 2002, Lester and Hayward 2005).
Haliotis rubra, Haliotis laevigata, Haliotis cyclobates and Haliotis scalaris. Molecular studies (nucleotide sequence of the internal transcribed spacers (ITS) in the ribosomal gene cluster (rDNA)) indicated that P. olseni occurs in many species of molluscs from Australia and is homologous to Perkinsus olseni (=atlanticus) in clams from Europe and eastern Asia. Perkinsus olseni was experimentally transmitted and highly infectious to a range of molluscs in the laboratory including two lamellibranchs, Pinctada sugillata and Anadara trapezia.
Impact on the host
Proliferates in tissues and may produce pustules (spherical brown abscesses up to 8 mm in diameter containing a caseous creamy-brown deposit) in the foot and mantle of H. rubra and H. laevigata thereby reducing market value. In two outbreaks of P. olseni in abalone culture facilities in Australia, about 30 to 40% mortality occurred among H. laevigata of 3 4 cm in shell length. In both cases H. rubra taken from an infected site had been introduced to the facility prior to the outbreaks (Goggin and Lester 1995). Apparently, P. olseni is not lethal to H. rubra (Lester and Hayward 2005). Suspected of causing mortalities in abalone and associated with devastated stocks of H. laevigata from the Gulf of St. Vincent side of the Yorke Peninsula, South Australia (Lester 1986; Lester et al. 1990; ODonoghue et al. 1991). After the epizootic had passed, two attempts were made to repopulate the area by transplanting adult H. laevigata. Eight months after the first attempt, many of the transplanted abalone had died and 15 of 20 survivors examined were heavily infected with P. olseni. Only the second attempt, conducted a year after the first attempt and three years following the epizootic event, appeared to be successful (Goggin and Lester 1995).
Transmission of this parasite occurs directly between individual molluscs. Prezoosporangia that escape from rupturing pustules (abscesses) or decaying dead abalone undergo further development to zoosporangia in seawater. Within nine days at 20 °C and three days at 28 °C, hundreds of motile, biflagellated zoospores (about 3 by 5 µm) exit from the zoosporangium. The zoospores are infective to abalone as well as other molluscs (Goggin et al. 1989). On Taylor Island, South Australia, field studies using molecular detection techniques indicated that infections of P. olseni in wild Haliotis rubra were positively correlated with both water temperature and size of abalone. Also, the parasite was being maintained by H. rubra with negligible contributions from other susceptible abalone species or other molluscs (Lester et al. 2001). Subsequent data and analysis by Hayward et al. (2002) indicated that the transmission of P. olseni among the wild H. rubra appeared to be reduced and infections were less severe in 2002. This apparent reduction in disease was attributed to lower maximum summer sea surface temperatures (cooling of almost 3 °C to below 20 °C).
Gross Observations: Macroscopic necrotic nodules (0.5 to 8.0 mm in diameter) in the adductor muscle and mantle.
Histology: Cross-section through tissues showing P. olseni in the connective tissue of the adductor muscle and mantle, and free in brownish masses in the haemolymph. Three stages of P. olseni are found in the connective tissues of the host: 1) immature trophozoites (2 - 3 µm in diameter), 2) mature trophozoites (3 - 18 µm in diameter) with large vacuoles (up to 10 µm diameter with some containing a weakly eosinophilic vaculoplast) commonly called the "signet-ring" stage, and 3) tomonts (dividing cells, 12 - 35 µm in diameter) containing 2 to 32 developing immature trophozoites. Abalone will respond to infection by an accumulation of haemocytes that eventually develops into the pustule. Parasites that do not die within the necrotic pustules develop further into the large prezoosporangium (60 - 95 µm in diameter, also called hypnospores). Although P. olseni appears to differ from P. marinus in some morphological features (size of mature trophozoites, occurrence and appearance of vacuoplasts, mean ratio of the length of the discharge tube to the diameter of the prezoosporangia, and size of zoospores) morphological characters do not consistently group or separate the various Perkinsus spp.
Immunological Assays: Polyclonal antibodies raised against Perkinsus marinus bound to some but not all Perkinsus sp. isolated from various species of Australian molluscs.
DNA Probes: The internal transcribed spacers ITS1 and ITS2 and the connecting 5.8S region of the riboromal RNA was sequenced. There is 13% difference in the ITS1 plus ITS2 with that of Perkinsus marinus but only 0.8% difference with that of Perkinsus olseni (=atlanticus) from clams in Portugal and two other isolates from Australia; Perkinsus sp. from the jewel box and blood cockle. Perkinsus olseni species-specific primers have been designed for polymerase chain reaction (PCR) and in-situ hybridization (ISH) assays (Moss et al. 2006).
Culture: Examination of tissues placed in Fluid Thioglycollate Medium (FTM) as described for Perkinsus marinus. After approximately 3 to 14 days of incubation in FTM at 25 °C, infected samples should contain Lugol positive prezoosporangia (diameter of 56-94 µm). This diagnostic procedure is frequently referred to as Ray's thioglycollate test (or technique). Partial positive (non-infectious) controls for this test can be made by inactivating the prezoosporangia (via freezing, ethanol or formalin immersion) which will retain their iodinophilic properties (Moore et al. 2002). Note: the Ray's thioglycollate test is not specific for P. olseni.
Methods of control
No known methods of prevention or control. For H. rubra and H. laevigata, stress such as high temperature (e.g., 20 °C) or temporary food shortage may predisposes the abalone to disease. However, infected H. rubra but not H. laevigata appear to contain and possibly eliminate the infection at 15 °C and during the winter (Lester and Davis 1981, Goggin and Lester 1995). Lester and Hayward (2005) concluded that a reduction in the number of abalone with abscesses (i.e., removal of older abalone from a hotspot) should result in a drop in the prevalence of infection in an area if other reservoir alternate host(s) do not occur in that area.
Disease outbreaks in abalone culture facilities were controlled by isolating infected tanks, removal of infected abalone, followed by washing equipment in fresh water (Goggin and Lester 1995). Commercial UV irradiation equipment that gives a dose of 60,000µWs/cm2 would almost totally prevent the passage of viable P. olseni in incoming water (Lester and Hayward 2005). Prezoosporangia and trophozoites can tolerate a wide range of salinities and temperatures and when enclosed in tissue, were also resistant to chlorine. Also, because this parasite can survive freezing (-60 °C) in abalone tissue for 197 days, its potential for being spread is high in relation to processing plants (Goggin et al. 1990). Abalone from areas with records of the disease should not be imported into areas with no record of P. olseni unless suitable precautions are taken not to return potentially contaminated materials to the marine environment.
Bower, S.M. 2000. Infectious diseases of abalone (Haliotis spp.) and risks associated with transplantation. In: Campbell, A. (Editor), Workshop on Rebuilding Abalone Stocks in British Columbia. Canadian Special Publication of Fisheries and Aquatic Sciences 130: 111-122.
Bower, S., E. Burreson and K. Reece. 2003. Annex 10: Review of molecular techniques used to differentiate the various species/isolates of Perkinsus. Report of the Working Group on Pathology and Diseases of Marine Organisms, Aberdeen, UK, 11-15 March 2003. Mariculture Committee, ICES CM 2003/F:03, Ref. ACME, pg. 54-60 (for electronic version see: http://www.ices.dk/products/CMdocs/2003/F/F0303.PDF (pg 60 of 101)).
Goggin, C.L. 1994. Variation in the two internal transcribed spacers and 5.8S ribosomal RNA from five isolates of the marine parasite Perkinsus (Protista, Apicomplexa). Molecular and Biochemical Parasitology 65: 179-182.
Goggin, C.L. and R.J.G. Lester. 1995. Perkinsus, a protistan parasite of abalone in Australia: a review. Marine Fisheries Research 46: 639-646.
Goggin, C.L., K.B. Sewell and R.G.J. Lester. 1989. Cross-infection experiments with Australian Perkinsus species. Diseases of Aquatic Organisms 7: 55-59
Goggin, C.L., K.B. Sewell and R.J. Lester. 1990. Tolerances of Perkinsus spp. (Protozoa, Apicomplexa) to temperature, chlorine and salinity. Journal of Shellfish Research 9: 145-148.
Hayward, C., R. Lester, S. Barker, H. McCallum, A. Murrell and S. Kleeman. 2002. Transmission of Perkinsus olseni among wild blacklip abalone in South Australia. (Abstract). Handbook and Abstracts, Fifth Symposium on Diseases in Asian Aquaculture, Queensland, Australia, 24-28 November 2002. Pg. 139.
Lester, R.J.G. 1986. Abalone die-back caused by protozoan infection? Australian Fisheries 45: 26-27.
Lester, R.J.G. and G.H.G. Davis. 1981. A new Perkinsus species (Apicomplexa, Perkinsea) from the abalone Haliotis ruber. Journal of Invertebrate Pathology 37: 181-187.
Lester, R.J.G. and C.J. Hayward. 2005. Control of Perkinsus disease in abalone. Final Report for FRDC Project no. 2000/151, Fisheries Research and Development Corporation, Australian Government, and The University of Queensland - Marine Parasitology, Brisbane. Pg. 50.
Lester, R.J.G., C.L. Goggin and K.B. Sewell. 1990. Perkinsus in Australia. p. 189-199. In F.O. Perkins and T.C. Cheng [ed.]. Pathology in Marine Science. Academic Press Inc., San Diego. CA.
Lester, R.J.G., S.N. Kleeman, S.C. Barker and H.I. McCallum. 2001. Epidemiology of Perkinsus olseni, pathogen of abalone. (Abstract). Book of Abstracts, European Association of Fish Pathologists, Tenth International Conference "Diseases of Fish and Shellfish. Trinity College Dublin, Ireland, 9 - 14 September 2001. pg. O-006.
Moore, B.R., S.N. Kleeman and R.J.G. Lester. 2002. The development of a positive non-infectious control for the detection of Perkinsus using the Ray test. Journal of Shellfish Research 21: 871-873.
Mortensen, S., I. Arzul, L. Miossec, C. Paillard, S. Feist, G. Stentiford, T. Renault, D. Saulnier and A. Gregory. 2007. Molluscs and crustaceans, 5.3.18 Perkinsosis due to Perkinsus olseni. In: Raynard, R., T. Wahli, I. Vatsos, S. Mortensen (eds.) Review of disease interactions and pathogen exchange between farmed and wild finfish and shellfish in Europe. VESO on behalf of DIPNET, Oslo. p. 389-396. (For electronic publication see www.dipnet.info under "Documents", subgroup "Reports and project deliverables").
Moss, J.A., E.M. Burreson and K.S. Reece. 2006. Advanced Perkinsus marinus infections in Crassostrea ariakensis maintained under laboratory conditions. Journal of Shellfish Research 25: 65-72.
Murrell, A., S.N. Kleeman, S.C. Barker and R.J.G. Lester. 2002. Synonymy of Perkinsus olseni Lester & Davis, 1981 and Perkinsus atlanticus Azevedo, 1989 and an update on the phylogenetic position of the genus Prekinsus. Bulletin of the European Association of Fish Pathologists 22: 258-265.
O'Donoghue, P.J., P.H. Phillips and S.A. Shepherd. 1991. Perkinsus (Protozoa: Apicomplexa) infections in abalone from South Australian waters. Transactions of the Royal Society of South Australia 115: 77-82.
Bower, S.M. (2010): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Perkinsus olseni of Abalone.
Date last revised: November 2010
Comments to Susan Bower
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