Diseases of pearl oysters and other molluscs: a Western Australian perspective.
KEY WORDS: abalone, ciliates, parasites, Perkinsus, Thraustochytridea, scallops
The most valuable of the marine mollusc culture industries in Western Australia is pearling. The main species cultured is the golden-lipped pearl oyster Pinctada maxima (Jameson), but other pearl shells grown commercially include P. margaratifera (L), P. albina (Lamarck) and Pteria penguin (Roding). The pearling industry is supported by four operational hatcheries capable of supplying all of the spat requirements of industry. Western Australia also has a small mussel industry based on culture of Mytilus galloprovincialis; an abalone culture industry based on hatchery produced Haliotis spp. (greenlip abalone, H. laevigata (Donovan), brownlip abalone, H. conicopora Peron and Roe's abalone, 14. roei Gray); a rock oyster farm hatching and growing Saccostrea sp.; a hatchery producing scallops (Amusium balloti Bernardi) for reseeding and there is also some experimental reseeding of tropical reefs with hatchery produced trochus (Tectus niloticus [L]).
Because of this aquaculture activity, the disease status of these Western Australian molluscs is relatively well known. To date there have been few diseases of concern. The health status of the aquaculture industry in a State such as Western Australia is based on two factors. The first is the presence of a disease reporting procedure and a diagnostic capacity to identify causative agents. The second is the existence of passive and targeted surveillance programs aiming to identify endemic diseases not associated with morbidity or mortality. In Western Australia a dedicated fish disease diagnostic unit of the Department of Fisheries has worked with industry since 1988 to identify and solve disease problems. There have also been 3 major mollusc disease surveys in the State; one involved a wide range of molluscs around the coast (Hine & Thorne 2000), one on P. maxima (Humphrey et al. 1998) and an ongoing national survey of abalone diseases. Some additional information is available in isolated publications on individual parasites and further unpublished information is available from laboratory records. Parasites that are known from shellfish in Western Australia are listed in Table 1.
The disease that has caused most economic loss to the P. maxima industry is vibriosis (Pass et al. 1987). In the 1980s Vibrio spp. bacteria caused significant losses to the pearling industry that have now been overcome by improved management practices. The presence of a very rare haplosporidan in the digestive tubules is cause for concern (Hine & Thorne 1998) because very little is known about the biology of this parasite. The haplosporidan has been identified only three times and each time the oysters on the infected farm site have been destroyed. During the second occurrence, in late December 1995, 4.6% of a sample of 150 farmed juvenile pearl oysters was found infected within 6 weeks of being set at a remote site in Western Australia. By the time the oysters were destroyed 15 days later the infection prevalence had increased to 10% (n = 238). The notifiable disease Perkinsus olseni/ atlanticus has been reported from P. maxima in the Torres Strait (Norton et al. 1993a) but not from Western Australia, though the disease agent does occur here in bivalve molluscs other than P. maxima (Table 1).
Another parasite of concern in pearl oysters has only been detected in spat from the Exmouth Gulf and outlying islands since 2001, despite comprehensive monitoring of oysters in the State and the industry practice of harvesting and translocating shell to all other oyster growing areas for over 50 y. It is an intracellular ciliate similar in appearance to ciliates reported as nonpathogenic commensals in Spanish Mytilus galloprovincialis (Villalba et al. 1997) and in Mytilus edulis (L) on the east and west coasts of North America (Figueras et al. 1991a). The teardrop shaped basophilic ciliate (10.15 [micro]m x 5 [micro]m) has a dense polymorphic macronucleus and normally occupies an intraluminal or intraepithelial location within the digestive gland in P. maxima spat (Fig. 1). The ciliate is often associated with an inflammatory response in oysters smaller than 70 mm with 20-50 mm shell being most affected. A feature in smaller (20-40 mm) spat is the ciliate's capacity to penetrate the mucosal basal lamina and reside within hemolymph spaces or free within interstitial connective tissue. Translocation of infected pearl oysters beyond the Exmouth zone is not permitted.
[FIGURE 1 OMITTED]
The origin of the ciliate is unknown, and its relationship to similar organisms seen in northern hemisphere mussels is unknown. Either the ciliate has become much more prevalent in recent years or it has been introduced into the region. Its infectivity to other bivalves is also unknown.
During surveys to determine the distribution of the ciliate, a single infected oyster was found with an enigmatic unidentified proctistan parasite (Fig. 2). The putative sporoblasts develop within the epithelial cells of the digestive tubules and fill the lumen of the tubules, with an associated basophilic hemocyte inflamma inflammatory response. With a prevalence of less than 0.005% it is not feasible to attempt follow-up sampling, but it does indicate that parasitic organisms can be present at extremely low prevalence in a population.
[FIGURE 2 OMITTED]
Another proctistan was detected at a low prevalence in P. maxima from the Exmouth Gulf during investigations into the ciliate parasite. The parasite was elongated (30 [micro]m x 20 [micro]m) and intimately associated with the digestive gland epithelial cells to which it appeared to have a sessile attachment (Fig. 3). The tubule epithelial cells beneath the site of attachment were multinucleated and ultrastructural examination indicated the multiple nuclei were of molluscan origin, suggesting the proctistan had induced this change within the host.
[FIGURE 3 OMITTED]
A proctistan tentatively attributed to the Thraustochytridea was identified in moribund, gaping P. maxima from a farm that had experienced losses after a cyanobacterial (Trichodesmium sp.) bloom. The affected oysters showed extensive necrosis of external epithelial surfaces of the palps and mantle with invasion of the underlying leydig tissues by brown pigmented and eosinophilic, segmented unicellular organisms 10-15 [micro]m in diameter, and smaller dense basophilic 5-[micro]m diameter cells that appeared to be embedded in a mucinous matrix (Fig. 4).
[FIGURE 4 OMITTED]
One of 411 Saccostrea glomerata (Gould) collected between Carnarvon to the Dampier Archipelago during 1995 was infected with a parasite histologically identical to Marteilia sydneyi Perkins and Wolf, the cause of QX disease in Saccastera commercialis (Iredale and Roughley) in Queensland and New South Wales. The same pathogen was also found in S. glomerata by Hine and Thorne (2000). On the eastern seaboard where QX occurs, it has caused losses of over 90% (Witney et al. 1988) and has resulted in a decline in New South Wales rock oyster production since the 1970s.
S. cucullata (Born) from Airlie Island, Rosily Island, Varanus Island, Hermite Island, King Bay and East Lewis Island in the north of Western Australia are infected by a second species of pathogenic haplosporidan protozoan, Haplosporidium sp. The parasite was identified as the cause of a significant mortality (up to 80%) of rock oysters at Airlie Island (Hine & Thorne 2000). The presence of this haplosporidan will threaten the viability of any rock oyster farm sited north of Exmouth.
The ova of mussels in Cockburn Sound, Western Australia are infected with the microsporidan Steinhausia mytilovum but the parasite has not been found in mussels from other growing areas. Heavily infected mussels are readily identified by the gross appearance of the mantle of cooked females. Infected mantle tissue has an uneven surface with depressed cream white patches and swollen tubercles forming spots against the orange-pink background color of healthy female gonad. Mussel samples in August and October 1995 from sites around the Sound showed a prevalence of Steinhausia sp. ranging from 22% to 57%, or 44.4% overall. Histology revealed that the patches and tubercles were associated with a marked infiltration of circulating granulocytes and large basophilic hemocytes into affected follicles with resorption of the germinal epithelium and ova (Fig. 5). Small basophilic hemocytes were not prominent in the response. Also present in 2% to 4% of the ova were parasitophorous vacuoles closely associated with the nucleus. These vacuoles measured 13.27 [+ or -] 2.37 [micro]m (n = 20) and contained over 30 small spores 2.5-[micro]m diameter. There was usually only one vacuole in an ovum, but occasionally up to three parasitophorous vacuoles could be seen. In histological sections the presence of resorbing follicles and focal accumulation of granulocytes among the developed ova was evidence of infection, but infected oocytes were also observed in apparently healthy follicles. Ova (normal, infected and degenerating), granulocytes, large basophilic granular hemocytes and cell debris occurred in the ciliated gonad ducts. Parasitophorous cysts containing spores also occurred free in the lumen of the ducts. The spore walls and contents stained negative to Feulgen, Ziehl-Neelsen and Grocotts, were Gram positive and Periodic-Acid-Schiffs negative.
[FIGURE 5 OMITTED]
The relationship between this Steinhausia sp. and the one previously described by Anderson et al. (1995) from cysts within ova of the rock oyster S. commercialis in Queensland, Australia is unknown. The biology of Steinhausia sp. is not well understood. Field (1923) reported that infected eggs were shed along with normal eggs. Sparks (1985) suggested that infected eggs do not seem to become necrotic or degenerate though Rybakov and Kholodkovskaya (1987) noted that Steinhausia sp. clearly distorts the nucleus of the ovum and can also cause the destruction of the egg, as has been seen in Western Australian mussels. It is likely that the loose spores are released along with intact eggs or through phagocytosis and subsequent diapediasis. Figueras (1991) reported that the presence of the parasite is always accompanied by a strong hemocyte response and the impact on the host has been described as severe (Rybakov & Kholodkovskaya 1987) to negligible (Maurand & Loubbs 1979). In the case of the S. mytilovum infection seen here, there is an absence of the typical bivalve inflammatory response, which involves invasion of the site of trauma primarily by small agranular hyalinocytes (90%), granular basophils (8%) and granular acidophils (2%) (Bayne et al. 1979, Brereton & Alderman 1979). Instead the major components are the phagocytic hemocyte and the granular acidophil and the process closely resembles gonad resorption. Prevalence of Steinhausia mytilovum does not increase with the size of the mussel, suggesting that infection is annual (Table 2).
The proctistan observed in the ova of Western Australian blue mussel has the same measurements and appearance under the light microscope as S. mytilovum from both European and American M. galloprovincialis. The taxonomy of the blue mussel in Western Australia is disputed. Still referred to as M. edulis planulatus (L), electrophoretic studies have shown that the species is M. galloprovincialis and that M. galloprovincialis from Australasia, eastern Asia, Western Europe, the Mediterranean and California, and M. edulis from eastern North America and Western Europe are electrophoretically distinct species with an overlapping distribution (Koehn 1991, McDonald et al. 1991, Geller et al. 1993). There is a fossil record of mussels in Australian waters and Koehn (1991) hypothesized that M. galloprovincialis may have been an early introduction into the Northern Hemisphere as a hull-fouling organism. Distribution of parasites often reflects the distribution of their primary hosts so S. mytilovum may also be an introduction from the Southern Hemisphere. Steinhausia mytilovum is reported to infect the ova of M. edulis along the Atlantic coast of the USA (Field 1923, Figueras et al. 1991a). Sparks (1985) reported that, based on examination of thousands of mussels, the parasite was absent from California, Oregon and Washington and it was unreported from Europe. It now occurs in M. galloprovincialis from Spain (Figueras et al. 1991b), Italy (De Vincentiis & Renzoni 1963), Greece (Rayyan et al. 2004), the Black Sea (Rybakov & Kholodkovskaya 1987, Gayevskaya & Machkevskiy 1991), northern France (Comtet et al. 2004) and the west coast of the USA (Hillman 1990, 1991). Hillman (1990) noted that M. galloprovincialis had been accidentally introduced into southern California and suggested that S. mytilovum had been introduced with the mussels.
There are two nematode larvae found in scallops (Amusium balloti) in the Shark Bay area of Western Australia. The "common" nematode in scallops is Sulcascaris sulcata. This was reported by Lester et al. (1980) to infect up to 64% of the landed catch in Shark Bay and occurs in a brownish capsule 3-7 mm dia. The adult nematodes of S. sulcata live in the loggerhead turtle Caretta caretta (L) and have a wide geographic distribution and range of molluscan hosts. The second species, Echinocephalus sp., forms small yellow-brown cysts 2-3 mm dia. Lester et al. (1980) reported that only 2 of 10 scallops he examined were infected, but in recent years the nematode has been much more common and in 2001 was the dominant nematode in A. balloti. The genus Echinocephalus occurs widely in molluscs in warm waters, and probably matures in marine skates or rays.
The Western Australian component of a national health survey of abalone has recently been completed. In this survey, up to 25% of wild-caught green-lipped abalone (Haliotis laevigata) were infected with trematode metacercariae. Low prevalences of proctistans in the lumen of the stomach and digestive gland, apicomplexans and putative viral inclusions in the intestinal tract were observed. Abalone in Western Australia are free of the disease perkinsosis, found in South Australia and New South Wales, but the organism does exist on the south coast of Western Australia. Perkinsus is a primitive fungus-like organism of uncertain taxonomic status, probably in the phylum Labrinthulomycota. A worldwide effort to understand the taxonomy of this organism (or group of organisms) is underway. In South Australia and New South Wales Perkinsus olseni/atlanticus affects abalone with yellow-green pus filled blisters (0.5-8 mm dia.) containing a creamy brown deposit. Once processed the lesions appear as pale brown circles. Perkinsosis occurs in a variety of shellfish in the north of the State (Hine & Thorne 2000), however, a survey of 300 abalone from six sites along the south and west coasts of Western Australia in 1995 were negative for Perkinsus sp. by the thioglycolate media method. Subsequently, in 2003 Perkinsus sp. was cultured from the gill tissue of one clinically normal abalone (14. laevigata) from the south coast.
Whereas, overall, disease has not been a problem for the molluscs industry in Western Australia, it is certain that many more pathogenic organisms remain to be discovered, particularly as molluscs become subject to aquaculture or are subject to environmental stresses associated with economic activity. Because of the age of the Australian continent and the relative isolation of the coastal fauna it is likely that many of these will prove to be unique to Western Australia. Strict controls are therefore imposed to limit translocation of parasites by aquaculture. Whereas these pathogens may represent an economic threat, it is probable that they will also provide new insights on the zoogeography and derivation of the Western Australian mollusc fauna.
The authors thank the shellfish industries in Western Australia for their assistance in providing specimens and Paul Hillier and Tina Thorne for help in collecting mussels. Melanie Crockford, Greg Maguire and Fran Stephens provided editorial comment.
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J. B. JONES * AND J. CREEPER
Department of Fisheries, Government of Western Australia, P.O. Box 20, North Beach, WA 6920, Australia
* Corresponding Author. E-mail: email@example.com
TABLE 1. Molluscan disease-causing agents (other than bacteria) reported from Western Australia. Associated Agent Host with Epizootic? Virus Virus-like inclusions Pinctada maxima No Haliotis laevigata No Isognomon isognomum No Pinna bicolor No Saccostrea cucullata No Haliotis laevigata No Papova-like virus Pinctada maxima No Rickettsia-like organisms Barbatia helblingii No Haliotis laevigata No Isognomon isognomum No Pinctada maxima No Pinna bicolor No Pinna deltoides No Pteria penguin No Saccostrea glomerata No Saccostrea cucullata No Stavilia horrida No Haplosporidia Bonamia sp. Ostrea sp. Yes Haplosporidium sp. Saccostrea cucullata Yes Haplosporidium sp. Pinctada maxima No Marteilia sydneyi Saccostrea cucullata No Saccostrea glomerata No Mikvocytos roughleyi Saccostrea sp. No Microspora Steinhausia mytilovum Mytilus galloprovin- No cialis Marteilioides sp. Saccostrea echinata No Apicomplexa Heart apicomplexan Pinctada maxima No Perkinsus olseni Barbatia helblingii No Isognomon isognomum No Malleus meridianus No Pinctada albina No Pinna deltoides No Saccostrea cucullata No Saccostrea glomerata No Septifer bilocularis No Spondylus sp. No Haliotis laevigata No Ciliates Ciliates on gills Pinctada spp. No Saccostrea sp. No Ciliates in gut Pinctada maxima No Haliotis laevigata No Ancistrocomids Pinctada albino No Pinna bicolor No Saccostrea cucullata No Saccostrea echinata No Saccostrea glomerata No Haliotis laevigata No Sphenophyra-like ciliates Saccostrea glomerata No Trematoda Proctoeces sp. Malleus meridianus No Sporocysts Haliotis laevigata No Isognomon isognomum No Pinctada albina No Pinctada maxima No Malleus malleus No Saccostrea echinata No Saccostrea glomerata No Cestoda Tylocephalum sp. Dendostrea folium No Isognomon isognomum No Malleus malleus No Malleus meridianus No Pinctada spp. No Pinna bicolor No Pinna deltoides No Pteria penguin No Saccostrea cucullata No Saccostrea glomerata No Saccostrea echinata No Stavilia horrida No Tetrabothriate cestodes Pteria penguin No Nematoda Nematode larvae Saccostrea cucullata No Sulcascaris sulcata Amusium balloti No Echinocephalus sp. Amusium balloti No Gregarines Nematopsis sp. Saccostrea cucullata No Unidentified gregarine Pinctada maxima No Copepoda Anthessius pinctadae Pinctada maxima No Agent Host Reference Virus Virus-like inclusions Pinctada maxima Pass et al. (1988); Humphrey et al. (1998) Haliotis laevigata Laboratory records Isognomon isognomum Hine & Thorne (2000) Pinna bicolor Hine & Thorne (2000) Saccostrea cucullata Hine & Thorne (2000) Haliotis laevigata Laboratory records Papova-like virus Pinctada maxima Norton et al. (1993b); Humphrey et al. (1998) Rickettsia-like organisms Barbatia helblingii Hine & Thorne (2000) Haliotis laevigata Laboratory records Isognomon isognomum Hine & Thorne (2000) Pinctada maxima Humphrey et al. (1998) Pinna bicolor Hine & Thorne (2000) Pinna deltoides Hine & Thorne (2000) Pteria penguin Hine & Thorne (2000) Saccostrea glomerata Hine & Thorne (2000) Saccostrea cucullata Hine & Thorne (2000) Stavilia horrida Hine & Thorne (2000) Haplosporidia Bonamia sp. Ostrea sp. Laboratory records Haplosporidium sp. Saccostrea cucullata Hine & Thorne (2000) Haplosporidium sp. Pinctada maxima Hine & Thorne (1998) Marteilia sydneyi Saccostrea cucullata Hine & Thorne (2000) Saccostrea glomerata Hine & Thorne (2000) Mikvocytos roughleyi Saccostrea sp. Laboratory records Microspora Steinhausia mytilovum Mytilus galloprovin- Laboratory records cialis Marteilioides sp. Saccostrea echinata Hine & Thorne (2000) Apicomplexa Heart apicomplexan Pinctada maxima Humphrey et al. (1998) Perkinsus olseni Barbatia helblingii Hine & Thorne (2000) Isognomon isognomum Hine & Thorne (2000) Malleus meridianus Hine & Thorne (2000) Pinctada albina Hine & Thorne (2000) Pinna deltoides Hine & Thorne (2000) Saccostrea cucullata Hine & Thorne (2000) Saccostrea glomerata Hine & Thorne (2000) Septifer bilocularis Hine & Thorne (2000) Spondylus sp. Hine & Thorne (2000) Haliotis laevigata Laboratory records Ciliates Ciliates on gills Pinctada spp. Humphrey et al. (1998) Saccostrea sp. Laboratory records Ciliates in gut Pinctada maxima Laboratory records Haliotis laevigata Laboratory records Ancistrocomids Pinctada albino Hine & Thorne (2000) Pinna bicolor Hine & Thorne (2000) Saccostrea cucullata Hine & Thorne (2000) Saccostrea echinata Hine & Thorne (2000) Saccostrea glomerata Hine & Thorne (2000) Haliotis laevigata Laboratory records Sphenophyra-like ciliates Saccostrea glomerata Hine & Thorne (2000) Trematoda Proctoeces sp. Malleus meridianus Hine & Thorne (2000) Sporocysts Haliotis laevigata Laboratory records Isognomon isognomum Hine & Thorne (2000) Pinctada albina Hine & Thorne (2000) Pinctada maxima Humphrey et al. (1998); Pass (1987); Hine & Thorne (2000) Malleus malleus Hine & Thorne (2000) Saccostrea echinata Hine & Thorne (2000) Saccostrea glomerata Hine & Thorne (2000) Cestoda Tylocephalum sp. Dendostrea folium Hine & Thorne (2000) Isognomon isognomum Hine & Thorne (2000) Malleus malleus Hine & Thorne (2000) Malleus meridianus Hine & Thorne (2000) Pinctada spp. Hine & Thorne (2000); Humphrey et al. (1998) Pinna bicolor Hine & Thorne (2000) Pinna deltoides Hine & Thorne (2000) Pteria penguin Hine & Thorne (2000) Saccostrea cucullata Hine & Thorne (2000) Saccostrea glomerata Hine & Thorne (2000) Saccostrea echinata Hine & Thorne (2000) Stavilia horrida Hine & Thorne (2000) Tetrabothriate cestodes Pteria penguin Hine & Thorne (2000) Nematoda Nematode larvae Saccostrea cucullata Hine & Thorne (2000) Sulcascaris sulcata Amusium balloti Lester et al. (1980) Echinocephalus sp. Amusium balloti Lester et al. (1980) Gregarines Nematopsis sp. Saccostrea cucullata Hine & Thorne (2000) Unidentified gregarine Pinctada maxima Humphrey et al. (1998) Copepoda Anthessius pinctadae Pinctada maxima Humphrey et al. (1998) TABLE 2. Prevalence of Steinhausia sp. in female Mytilus galloprovincialis from Cockburn Sound, Western Australia. Number Total number Size of host (mm) infected examined % infected <50 16 34 47.0 50-59 33 65 50.7 60-69 23 80 28.7 70-79 93 189 49.2 80-89 51 114 44.7 >90 7 10 70.0 Overall 223 492 45.3 There is no increase in prevalence of infection with mussel size.
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|Publication:||Journal of Shellfish Research|
|Date:||Apr 1, 2006|
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