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Archived - Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish

Perkinsus of Clams and Cockles

Category | Common Name | Scientific Name | Distribution | Host Species
Impact on Host | Diagnostic Technique | Methods of Control | References | Citation


Category

Category 1 (Not Reported in Canada)

Common, generally accepted names of the organism or disease agent

Clam Perkinsus disease, Perkinsosis of Clams. The specific identity of some of these parasites has not yet been fully confirmed. Reports pertaining to the same parasite were assigned a letter code which is consistently applied to all available information on that parasite under each of the following headings.

Scientific name or taxonomic affiliation

a) Perkinsus olseni (=atlanticus). This parasite was initially described as Perkinsus atlanticus from clams in Europe (Azevedo 1989). Subsequently, a species of Perkinsus was found in Venerupis (=Tapes, =Ruditapes) philippinarum in Korea, China and Japan. Hamaguchi et al. (1998) found the nucleotide sequence of two internal transcribed spacers (ITS1 and ITS2) and the 5.8 S region of the RNA to be almost identical to P. atlanticus and Perkinsus olseni and suggested that the parasite in Japan may be P. atlanticus. Perkinsus olseni was originally described from abalone but cross-infection experiments and molecular studies suggested that only one species of Perkinsus occurs in a wide variety of molluscs, including clams, in Australia (Goggin et al. 1989, Goggin and Lester 1995). Further molecular studies identified considerable similarities between P. atlanticus and P. olseni (Berth 2004). Note: Because the synonymy of P. olseni and P. atlanticus proposed by Murrell et al. (2002) is upheld, the name P. olseni has priority. Using the same regions of the DNA and sequences of a non-transcribed spacer (NTS), Park et al. (2005) determined that the Perkinsus sp. from Korea was homologous to the species from Japan and identified it as Perkinsus olseni (=atlanticus). Elandaloussi et al. (2009) also found isolates from Spain homologous to P. olseni. Thus, P. olseni has a wide host range (infecting gastropods as well as bivalves) and a wide geographic range (including at least the coasts of Australia, New Zealand, Vietnam, Japan, Korea, China, Europe and Uruguay).
b) Perkinsus chesapeaki of Mya arenaria, identity based on minor differences in the morphology of the zoospore and molecular differences from Perkinsus marinus (McLaughlin et al. 2000b). At least two species of Perkinsus (P. marinus and P. chesapeaki) occur in M. arenaria (Kotob et al. 1999a, b; McLaughlin et al. 2000a). Crassostrea virginica, Macoma balthica and M. arenaria were experimentally susceptible to infection by mantle cavity inoculations of P. chesapeaki cultured isolates (Dungan et al. 2007a). Although P. chesapeaki was experimentally infective for oyster and clam hosts, a survey of wild bivalves in the Chesapeake Bay region revealed that P. chesapeaki infections predominated among members of at least 6 clam species and rarely (6%) occurred in C. virginica (Reece et al. 2008).
c) Perkinsus (=Labyrinthomyxa) andrewsi of Macoma balthica. Differentiated from Perkinsus marinus, Perkinsus atlanticusPerkinsus olseni and Perkinsus qugwadi based on sequence data from the rRNA locus (Coss et al 2001b). DNA analysis (using polymerase chain reaction (PCR) assays based on regions of the ribosomal RNA SSU loci (mainly ITS1 and ITS2)) indicated that this species can occur in other clams (Macoma mitchelli and Mercenaria mercenaria) as well as oysters (Crassostrea virginica) where it can coexist with Perkinsus marinus (Coss et al. 1999, 2001b). However, P. andrewsi was relatively rare (1.3%) in comparison to P. marinus detected in 58.4% of the 394 C. virginica examined during an investigation (Pecher et al. 2008). Analysis of the ITS regions of several species of Perkinsus (including several isolates of some species) consistently grouped P. chesapeaki and P. andrewsi (Murrell et al. 2002, Casas et al. 2002a,b). Also, analysis of the ITS sequence from cloned isolates of Perkinsus sp. from Mya arenaria and Tagelus plebeius from Chesapeake Bay suggested that the variations among ITS sequences of P. chesapeaki and P. andrewsi indicates true polymorphism within a single parasite species (Dungan et al. 2002). This variability was confirmed by Pecher et al. (2004) who reported a second rRNA gene unit in P. andrewsi in which all regions (except for the 5.8S) exhibited sequence differences from that initially described for this parasite and this second rRNA gene was similar to variations reported by Dungan et al. (2002). Note: If P. chesapeaki and P. andrewsi are synonymous, the name P. chesapeaki will have precedence over P. andrewsi (Burreson et al. 2003). A detailed study by Burreson et al. (2005) which included morphological (in vivo and in vitro), molecular (sequence analysis of three genetic loci: ribosomal RNA (rRNA) internal transcribed spacer (ITS) regions, rRNA large subunit (LSU) gene and actin gene) and experimental (reciprocal cross-infection) research, supported the synonymy of the two species. However, Pecher et al. (2008) considered this evidence to be too limited to support the synonymy. Specifically, Pecher et al. (2008) argued that P. chesapeaki was original described as a distinct morphotype by McLaughlin et al. (2000b) and the Perkinsus isolate that was analyzed to clarify the relationship of P. andrewsi and P. chesapeaki was designated as P. chesapeaki because it was isolated from the appropriate type host. Because this isolate appears to be morphologically identical to P. andrewsi, it may not be the P. chesapeaki that was originally described. Thus, the P. andrewsi designation was retained by Percher et al. (2008) who await additional evidence to support the synonymy despite the claim by Burreson et al. (2005) that the neohapantotype culture of P. chesapeaki was morphologically identical to the published morphology for P. chesapeaki and was obtained from the type host and from the same general area of Chesapeake Bay as the type locality.
d) Perkinsus sp. with a high level of sequence homology in the ITS-5.8S rRNA region to that of P. olseni and P. atlanticus (99.85% and 99.71%, respectively) and less homology (94.88% or greater) to that of other species (i.e., Perkinsus marinus, Perkinsus andrewsi and Perkinsus qugwadi). Like wise, the NTS rRNA region was only slightly different from that of P. olseni (1.31%) and P. atlanticus (3.73%) and highly different from that of P. marinus (24.62%) and P. andrewsi (53.45%). The species was not assigned in this case because the level of homology required to discriminate between species of Perkinsus had not been determined (Leethochavalit et al. 2003).
e) Perkinsus honshuensis (Dungan and Reece 2006).

NOTE 1: A Perkinsus atlanticus-like protist isolated in vitro from the clam Ruditapes decussatus in Galicia, Spain had a small subunit (SSU) ribosomal RNA gene sequence unlike that published for P. olseni (= atlanticus) in GenBank. This parasite was tentatively named "Pseudoperkinsus tapetis" and affiliated with fungus-like protists in the recently named Mesomycetozoa (Figueras et al. 1992, 1996, 2000). Like Perkinsus spp., this isolate developed large prezoosporangia (hypnospores) that stained dark blue with Lugol’s iodine stain after incubation in FTM (Figueras et al. 2001). Thus, the Ray's FTM assay can not be used to differentiate between the two species (Novoa et al. 2002). Also, the protease activity of Pseudoperkinsus tapetis extracellular products was different from those described for Perkinsus marinus (Ordás et al. 2001b).

NOTE 2: Perkinsus spp. cannot be discriminated on the basis of morphology, host species or geographic location. Even with molecular data, there can be problems recognizing true species boundaries if multiple clonal cultures are not established, if too few DNA clones are sequenced and if cultures are from only a restricted host or geographic range (Burreson et al. 2005). Recommendations for describing new species of Perkinsus were presented by Burreson et al. (2005).

Geographic distribution

a) France (Atlantic and Mediterranean coasts (Miossec et al. 2006) including at least four different French marine areas; Morbihan gulf, Arcachon bay, Leucate and Thau lagoons (Arzul et al. 2009) ), Portugal, Spain (including Galicia and Delta de l'Ebre, Catalonia (NW Spain), Huelva coast (SW Spain), Andalucia (S. Spain), Balearic Islands and Mediterranean coast), and Italy (the Mediterranean Sea and the NW Adriatic Sea); Great Barrier Reef, South Australia, northern Western Australia, northern New Zealand and in an Vietnamese ornamental clams (Tridacna crocea) imported into the U.S.A. (Sheppard and Phillips 2008, Sheppard and Dungan 2009); west and south coasts of South Korea, in Kumamoto, Hiroshima and Mie Prefectures of Japan, and Bohai Sea, along the northern coast of the Yellow Sea and the southern coast of China; and the coast of Uruguay. Perkinsus olseni (=atlanticus) was also tentatively identified from Macoma balthica in a tributary of Chesapeake Bay, USA (Kleinschuster et al. 1994).
b) Chesapeake Bay (McLaughlin and Faisal 2000) and Delaware Bay (Bushek et al. 2008), USA. Unidentified Perkinsus species were detected by Ray's FTM and genus-specific PCR (as described by Robledo et al. 2002) in Mercenaria mercenaria from the Gulf of Mexico coast of Florida (McCoy et al. 2007).
c) Virginia and Maryland (Chesapeake Bay) to Maine, USA (Pecher et al. 2008). Unidentified Perkinsus species were detected by Ray's FTM and genus-specific PCR (as described by Robledo et al. 2002) in Mercenaria mercenaria from the Gulf of Mexico coast of Florida (McCoy et al. 2007).
d) Gulf of Thailand in Chonburi Province, Thailand.
e) Gokasho Bay, Mie Prefecture, Japan.

Host species

a) Ruditapes (=Tapes, =Venerupis) decussatus, Ruditapes (=Tapes) semidecussatus, Venerupis rhomboides, Venerupis aurea, Venerupis (=Ruditapes) pullastra, Venus verrucosa and imported/cultured Venerupis (=Tapes, =Ruditapes) philippinarum introduced into France in the mid 1970's (Flassch and Leborgne 1992). In France, epizootiological surveys in 2004 and 2005 found prevalence and parasite burden higher in R. decussatus compared to V. philippinarum with no associated mortality (Arzul et al. 2009). In the North-Western Adriatic Sea (Italy), Perkinsus sp. (probably P. olseni) has also been detected in Cerastoderma edule, Chamelea gallina, Callista chione and other bivalves (Da Ros and Canzonier 1985, Canestri-Trotti et al. 2000) and in the area of Sardinia from Cerastoderma glaucum (Culurgioni et al. 2006). Many species of molluscs including Tridacna gigas, Tridacna maxima, Tridacna crocea, Anadara trapezia, Hippopus hippopus, Chama iostoma, Chama pacificus, Acrostengma unicolor and Katelysia rhytiphora from the southwestern Pacific Ocean including Macomona liliana, Barbatia novaezealandiae, and Austrovenus stutchburyi from northern New Zealand were infected (Goggin and Lester 1987, Hine 2002, Hine and Diggles 2002, Murrell et al. 2002, Dungan et al. 2007b). In northern Western Australia, a survey detected Perkinsus sp. in a wide diversity of bivalve molluscs including species of clams such as Barbatia helblingii (Hine and Thorne 2000). Venerupis (=Tapes, =Ruditapes) philippinarum and Protothaca jedoensis but not observed in 10 other molluscs (including Crassostrea gigas and Pinctada fucata martensii) from enzootic areas in South Korea (Choi and Park 1997, Park et al. 2001, Park et al. 2006). However, DNA of P. olseni was detected in Crassostrea ariakensis in its native range (China, Japan and Korea) and in Crassostrea hongkongensis from the south coast of China via molecular diagnostic screening using PCR based assays (Moss and Reece 2005, Moss et al. 2007). Also found in the commercial clam Pitar rostata from Uruguay (Cremonte et al. 2005). Crassostrea virginica and Mercenaria mercenaria were found susceptible to P. olseni via experimental exposure (inoculations and bath challenges with cultured or directly harvested parasites) in the laboratory (Moss et al. 2008).
b) Originally described from Mya arenaria which is designated as the type host. Also occurs in Macoma balthica, Tagelus plebeius, Macoma mitchelli, Mercenaria mercenaria, Mulinia lateralis, Rangia cuneata, Cyrtopleura costata and Crassostrea virginica (Burreson et al. 2005, Reece et al. 2008).
c) Macoma balthica designated as type host but also occurs in Macoma mitchelli, Mercenaria mercenaria and Crassostrea virginica (Coss et al. 2001b, Pecher et al. 2008).
d) Paphai undulata (Leethochavalit et al. 2003, 2004).
e) Venerupis philippinarum. Note that at least two Perkinsus spp. (P. olseni and P. honshuensis) infect Japanese Manila clams (Dungan and Reece 2006).

Impact on the host

In most clam species, the parasite frequently induces the formation of visible milky white cysts or nodule on the gills, foot, gut, digestive gland, kidney, gonad and mantle of heavily infected clams. The sometimes massive aggregation of Perkinsus sp. and haemocytes form lesions that may interfere with respiration and other physiological processes such as reproduction (fertility/fecundity, when large lesions occur in the gonads), growth and/or survival and thus have an impact on fishery productivity. Infection in Ruditapes decussatus has been associated with extensive mortalities in clam breeding areas located on the south coast of Portugal (Azevedo et al. 1990). However, on the Galician coast of Spain, perkinsosis did not appear to affect the energetic physiology of infected R. decussatus at about 15 °C but, Villalba and Casas (2001) speculated that higher temperatures may impact on disease severity. Also, Villalba et al. (2005) detected an annual pattern of infection with lowest mean intensity and prevalence of infection during the winter with the peak in spring significantly associated with seawater temperature at about 15 °C and clams from a perkinsosis-affected area had a significantly higher mortality than clam from a non-affected area (e.g., 7.0% and 2.8%, respectively, in early September, the time of year with the highest mortality). However, Elandaloussi et al. (2008) observed no obvious seasonality in the prevalence of P. olseni in V. philippinarum and R. decussatus from the NW Mediterranean Sea and there was no significant correlation between the intensity of infection in these clams and either seawater temperature or salinity. Also, Dang et al. (2010) detected no seasonal cycle of P. olseni in V. philippinarum at 34 stations throughout Arcachon Bay (SW France) where the prevalence of infection was high (on average between 70 and 100%). A two-year survey for P. olseni in V. philippinarum and R. decussatus along the coasts of France revealed a lower prevalence of infection along the Atlantic coast and English Channel than in the Mediterranean Sea but no abnormal mortalities were detected (Miossec 2006, Miossec et al. 2006). Goggin (1996) reported that P. olseni did not cause tridacnid clams (specifically, T. crocea), up to 100 mm shell length, to lose wet tissue weight. Montes et al. (2001) determined that P. olseni (=atlanticus) parasitism favours the development of opportunistic infections by bacteria and viruses which have detrimental effects in clam (R. semidecussatus) populations from the northern Mediterranean coast of Spain.

Haemocytes, especially granulocytes, of R. decussatus in vitro were able to phagocytose trophozoites but not zoospores of (López et al. 1997). In addition, secretion products from cultured of P. olseni (=atlanticus), which contained high protein concentration, acid phosphatase and protease activity, inhibited the phagocytosis of various particles, specifically zymosan, Escherichia coli and Vibrio tapetis (Ordás et al. 1999). Hégaret et al. (2007) found that V. philippinarum infected with P. olseni maintain haemocyte function but their immune system response to harmful or toxic algal exposure was modified by parasite infection. In addition, da Silva et al. (2008) determined that the prevalence and intensity of P. olseni decreased in clams exposed to the same toxic algae (Karenia selliformis), and K. selliformis cells were toxic to P. olseni during in vitro tests (altered morphology and increased percentage of dead P. olseni). Overall, initial exposure of P. olseni-infected clams to K. selliformis appeared to modify the host–parasite interaction by causing effects in both the host and its parasite.

Perkinsus sp. was considered as the cause of epizootic mass mortalities and decline in commercial harvest for the decade prior to 2005 of Venerupis philippinarum in Korea (Park et al. 1999, Choi and Park 2005) and China (Liang et al. 2001) and the cause of populations declines of this clam in Japan (Hamaguchi et al. 1998). Also, heavy infections observed in older clams in Korea appeared to cause retarded growth and delayed gamete maturation resulting in altered population dynamics and stability (Park and Choi 2001). In laboratory studies, P. olseni caused direct mortality in V. philippinarum juveniles (3 to 10 mm shell length) and the lethal level of infection was estimated at approximately 107 pathogen cells/g soft tissue weight (Shimokawa et al. 2010). However, results of clam disease surveys in Korea by Lee et al. (2001) indicated that caution should be applied when determining a causal relationship between Perkinsus sp. infections and V. philippinarum mortalities. Also, in Korea, results of surveys for Perkinsus sp. in V. philippinarum indicated that the spatial distribution of this parasite is in some way controlled by temperature, salinity and substrate type (Park and Choi 2001). Nago and Choi (2004) found that the prevalence and intensity of Perkinsus was lowest in September and highest in March in V. philippinarum from Jeju, an island off the south coast of Korea. However, Choi and Park (2005) reported that infection intensity was highest in October when most clams completed spawning and mass mortalities were observed in the clam beds on the west and south coast of South Korea.

Gills appear to be the main target tissues for most Perkinsus sp. in clams. In Mya arenaria, the most commonly observed lesions were clusters of trophozoites encapsulated in well-circumscribed walls forming a cyst-like structure. In advanced infections, the plethora of cysts in the branchial connective tissue was accompanied by loss of underlying tissue structures and gill lamellae (McLaughlin and Faisal 1998a). Similar observation were made by Park and Choi (2001), Lee et al. (2001) and Choi and Park (2005) for Perkinsus sp. in Venerupis philippinarum from Korea. Ruditapes decussatus and R. semidecussatus developed a defensive response to P. olseni (=atlanticus) which included differentiation of recruited granular haemocytes in the vicinity of the parasite and the de novo secretion of the polypeptide p225 (Montes et al. 1996, Montes et al. 1997). Ordás et al. (2000) determined that advanced infections in R. decussatus had a measurable effect on defence parameters, especially anti-bacterial activity and agglutination titres (lectins) in the haemolymph. Kim et al. (2006) and Kang et al. (2008) also detected lectin production in V. philippinarum infected with P. olseni in South Korea. However, in contrast, da Silva et al. (2008) found a low impact of P. olseni on the immune system of V. philippinarum, from Brittany, France, with no induction of lectin production. Immunofluorescence staining (using polyclonal monospecific IgG antiserum produced in rabbits) revealed that the lectin from V. philippinarum bound to the surfaces of purified (hypnospores) of Perkinsus sp. from Korea (Bulgakov et al. 2004). McLaughlin et al. (2000a) and McLaughlin and Faisal (2001) reported a difference in the production of extracellular proteins by P. chesapeaki and P. marinus that may help to explain the difference in pathology observed in infected Mya arenaria and Crassostrea virginica, respectively.

After reviewing epidemiological methods currently used in field studies to evaluate the occurrence of P. olseni in clams, Miossec et al. (2005) proposed recommendations for a basic methodological design to conduct an epidemiological survey. These recommendations included: clearly defining the target population, identifying sampling methodology (preferably as probabilistic as possible), calculate sample size according to survey objectives, evaluate the quantitative values of sensitivity and specificity for the diagnostic test used, and apply additional tools to fully characterize (identify) the pathogen (Miossec et al. 2005).

Diagnostic techniques

Gross Observations: Infected clams may have whitish nodules or cysts on the surface of the gill, digestive gland and mantle tissues due to a haemocytic response by the clam to Perkinsus spp.

Wet Mounts: Spherical bodies containing an eccentric vacuole (signet-ring) in cysts from moribund clams.

Histology: Systemic proliferation of haemocytes in response to immature trophozoites (=aplanospores), mature trophozoites (="signet-ring" or aplanospore with large eccentric vacuole that displaces the nucleus to the periphery of the cell), and tomonts (="rosette", sporangium, schizont or palintomic cells) stages of parasite development. In comparison to Perkinsus marinus from Crassostrea virginica, P. olseni (=atlanticus) from clams in Europe can have larger trophozoites (30-40 µm in diameter) but generally diameter varies from 3 to 15 µm and the trophozoites of Perkinsus sp. from V. philippinarum in Japan vary in size from 2 to 32.5 µm with average diameters of 12 to 15 µm. In most clams, infection is usually associated with an infiltration of numerous haemocytes into the surrounding tissues (Ordás et al. 2001a). Typical lesions were haemocyte-encapsulated granulomatous cysts in haemocyte-infiltrated connective tissues, containing Perkinsus cells that were often enrobed in an amorphous eosinophilic matrix apparently secreted by the clam haemocytes. Occasionally, the Perkinsus cells occur within granular haemocytes and Sheppard and Phillips (2008) reported a radiating corona pattern in the eosinophilic material surrounding some trophozoites. Lesions usually occur in gill and digestive system connective tissues and less frequently in gonad, mantle, kidney, and heart connective tissues (Dungan and Reece 2006).

Haemocytes

Figure 1. Extensive infiltration of haemocytes around clusters of Perkinsus sp. (arrows) causing congestion of the gills in Venerupis (=Ruditapes) philippinarum from Korea. Haematoxylin and eosin stain.

Haemocytes

Figure 2. Two clusters of many Perkinsus sp. (arrows) within an accumulation of haemocytes in the connective tissue of the gill of V. philippinarum from Korea. Haematoxylin and eosin stain.

Haemocytes

Figure 3. Two clusters of mature trophozoites (arrows) surrounded by haemocytes in the gills of V. philippinarum from Korea. Haematoxylin and eosin stain.
Images in Figs. 1-3 obtained from histological section kindly provided by Dr. K.-S. (Albert) Choi, Cheju National University, South Korea.

Haemocytes

Figure 4. Another example of several clusters and single trophozoites (arrows) surrounded by haemocytes in the gills of V. philippinarum from Korea. Haematoxylin and eosin stain.

In lightly infected M. arenaria, Perkinsus sp. usually occur in the gill lamellae, either free or surrounded (often encapsulated) by haemocytes, which appears to result in fusion between adjacent lamellae. Often clusters of trophozoites embedded in amorphous eosinophilic material and tissue debris formed cysts (17.8 ± 7.9 µm, range of 8 to 44 µm) in the gills. Entrapped trophozoites were circular or oval (3.8 ± 1.4 µm in diameter), uninucleate and each contained a large vacuole that occupied most of the cell. In more heavily infected M. arenaria, the cysts increased in number and in size (47.6 ± 12.8 µm, range of 24 to 68 µm) and their amorphous outer wall became more demarcated from the surrounding tissue. In advanced infections, Perkinsus sp. cells predominated the internal structure of the gill lamellae and subepithelial connective tissue. Cysts were also observed in the connective tissue between the tubules of the digestive gland, in the gonads and kidneys and were often associated with large lesions consisting of free and encapsulated Perkinsus sp. cells and haemocytes within an eosinophilic matrix. Perkinsus sp. propagated by schizogony with tomonts (7.9 ± 1.6 µm in diameter, range of 6 to 12 µm) containing up to 4 daughter cells (McLaughlin and Faisal 1998). Burreson et al. (2005) reported that host reactions to P. chesapeaki infections varied considerably in the three clam species (M. arenaria, T. plebeius and M. balthica) that have been studied histologically and this resulted in some differences in parasite sizes and morphologies observed in these different hosts.

Histopathological examination of clams often detect fewer infected clams than Ray's FTM assay (Rodríguez and Navas 1995, Almeida et al. 1999, Leethochavalit et al. 2004, Dungan et al. 2007b). To date, no well defined morphological features have been identified for differentiating between the various species of Perkinsus reported from clams and other molluscs. Also, trophozoite morphology does not have taxonomic value because it can be influenced by the host, the time of the year, and nutrient availability (Villalba et al. 2004).

Electron Microscopy: Chagot et al. (1986) and Comps and Chagot (1987) provided a brief description and a few ultrastructural images of a Perkinsus in R. decussatus from Portugal and Sagrista et al. (1996) provided a detailed account of zoosporulation of a Perkinsus in V. philippinarum from the Mediterranean coast of Spain. Also, the fine structure of clonally propagated in vitro life stages of Perkinsus andrewsi were described (Coss et al. 2001a). However, all features observed were consistent with those of other species of Perkinsus described from various molluscs.

Immunological Assay: An antiserum prepared against Perkinsus marinus (polyclonal antibodies, prepared by C.F. Dungan, Cooperative Oxford Laboratory, Oxford, MD, USA) cross reacted will trophozoites of Perkinsus sp. in histological sections of V. philippinarum from Japan (Maeno et al. 1999). Montes et al. (2002) used immunological techniques to localize a main proteinaceous component of the cell wall of P. olseni (=atlanticus) and determined that polyclonal antibodies to the protein cross-reacted with Perkinsus marinus. Rabbit anti-P. olseni IgG prepared by Park et al. (2010) was specific to all life stages, including the prezoosporangium, trophozoite, and zoospore by an immunofluorescent assay and was used to isolate P. olseni prezoosporangium-like cells from marine sediment collected from the west coast of Korea where P. olseni–associated clam mortality had recurred for the past decade. To date, no diagnostic assay based on anti-Perkinsus sp. antibodies has been rigorously validated, and antibodies that have been produced may exhibit cross-reactivity with dinoflagellates (Villalba et al. 2004).

DNA Probes: The internal transcribed spacers (ITS region including ITS1 and ITS2 and the connecting 5.8S region), the small subunit (SSU or 18S), the large subunit (LSU) and/or the non-transcribed spacer (NTS) of the ribosomal RNA locus and/or the actin gene of some Perkinsus isolates from various clams (and other bivalves) have been sequenced. Sequences have been compared between various isolates from various molluscs and species synonymy were proposed based on sequence similarities. For example, only slight differences (0.8%) were found between the ITS region of Perkinsus from clams and cockles and Perkinsus olseni from abalone in Australia (Goggin 1994). Hamaguchi et al. (1998) found this sequence in Perkinsus sp. from V. philippinarum in Japan to be almost identical (99.9%) to that reported by Goggin and Barker (1993) and Goggin (1994) from P. atlanticus, P. olseni and other Perkinsus sp. isolates. Casas et al. (2002a, b) also reported that the ITS sequence from 13 isolates of Perkinsus sp. from Ruditapes decussatus in Galicia (NW Spain) were closely matched with equivalent sequences from P. atlanticus, P. olseni, and Perkinsus sp. from Chama pacificus and A. trapezia. This synonymy was supported by Park et al. (2005) for Perkinsus from V. philippinarum in Korea and by Elandaloussi et al. (2009) for Perkinsus from clams (V. philippinarum and R. decussatus) in Spanish Mediterranean waters. In these publications, the gene sequences of the P. olseni (=atlanticus) isolates were distinct from those of P. marinus and also different from a cluster consisting of P. chesapeaki and P. andrewsi. Thus, the synonymy of P. olseni and P. atlanticus proposed by Murrell et al. (2002) is well supported by molecular evidence. Molecular sequences have also been used to identify new species. For example, the nucleotide sequences (ribosomal DNA internal transcribed spacer region, the large subunit rRNA gene, and actin genes) of one of four Perkinsus isolates from V. philippinarum in Gokasho Bay, Japan, that was morphologically unique, differed from those of all described Perkinsus species and was named Perkinsus honshuensis where as similar sequences from the other three isolates were consistent with those reported for P. olseni (Dungan and Reece 2006). Because Perknisus spp. tend not to be host specific and are morphologically similar, species identification based on rRNA sequence differences has resulted in controversy between investigators. For example, P. chesapeaki and P. andrewi initially described from different bivalves from Chesapeak Bay are now reported to occur in a similar range of host bivalves. Also, isolates from these host species are now known to have multiple polymorphic sequences in the rRNA gene with sequence similarities occurring in isolates identified as P. chesapeaki and P. andrewsi (Dungan et al. 2002, Percher et al. 2004). To further complicate the issue, corresponding type material was not deposited for each species (i.e., P. chesapeaki is represented by a type histological slide but no holotype in vitro culture isolate is available while P. andrewsi is represented by a holotype clonal culture but no type histological slide was deposited) making it difficult to compare the two species from original material. In an attempt to address the problem, Burreson et al. (2005) demonstrated that both parasites were indistinguishable based on molecular, morphological and experimental evidence and supported the synonymy of the two species. However, Pecher et al. (2008) retained the P. andrewsi designation because the neohapantotype culture isolate of P. chesapeaki presented by Burreson et al. (2005) appeared to be morphologically identical to P. andrewsi. Thus, Pecher et al. (2008) stated that the neohapantotype culture isolate may not be the P. chesapeaki originally described and await additional evidence to support the synonymy.

Various polymerase chain reaction (PCR) based diagnostic assays proposed to be genus-specific and/or species-specific have been developed (Reece et al. 2001, Robledo et al. 2002, Casas et al. 2002a, Park et al. 2005). For example: Hamaguchi et al (1998) designed a PCR method for the diagnosis of the Perkinsus sp. from V. philippinarum in Japan; Elston et al. (2003) used in situ hybridization to verify that the parasite in V. philippinarum from Korea was Perkinsus sp.; Kotob et al (1999a, b) used sequence analysis of the ITS regions of two isolates of Perkinsus sp. from Mya arenaria to suggest that the two isolates were different species of Perkinsus and Balseiro et al. (2010) identified a nested PCR assay for P. olseni to improve on the PCR assay described by Kotob et al. (1999b); Robledo et al. (1999), Cross et al. (2001b) and Percher et al. (2008) developed PCR-based diagnostic assays for P. andrewsi in the USA; Moss et al. (2006) designed P. olseni species-specific primers for polymerase chain reaction (PCR) and in-situ hybridization (ISH) assays; Robledo et al. (2000) and De la Herrán et al. (2000) developed a PCR-based diagnostic assay for P. olseni (=atlanticus) from Spain; Elandalloussi et al. (2004) developed a PCR-enzyme-linked immunosorbent assay (ELISA) for amplification of an IGS sequence region and rapid detection of Perkinsus species in which the specific hybridisation of DIG-labelled amplified products to species-specific capture probes was detected colourimetrically; and Abollo et al. (2006) developed a species-specific polymerase chain reaction-restriction fragment length polymorphism (PCR-RFLP) assay of the rRNA ITS region that used a single restriction enzyme (Rsa I) to discriminate P. chesapeaki and P. marinus and a combination of two endonucleases (Rsa I plus Hinf I) to discriminate P. olseni and P. mediterraneus. Arzul et al. (2009) employed the PCR procedures described by Casas et al. (2002a) and the RFLP of Abollo et al. (2006) followed by sequencing of products to identify the species of Perkinsus detected in clams for France. However, as cautioned by Burreson (2000), more research is necessary to compare PCR assays with standard diagnostic techniques before PCR can be recommended as the method of choice for perkinsosis diagnosis. For example, primers that target the NTS, a region with high inter-specific variation, have demonstrated good species specificity (Coss et al. 2001b). However, intra-specific variations within the NTS region has not been broadly assessed creating a risk of false negatives due to polymorphism within a species if the PCR primers do not bind the target sequence of all strains of that species (Villalba et al. 2004). Also, further research is required to resolve which DNA sequence differences are consistently significant in the identification of species and how these differences relate to biological parameters that can be used to describe and differentiate between closely related species. This is especially important if some species and/or strains of Perkinsus prove to be non-pathogenic for some or all host species. Nevertheless, molecular sequence data is playing an increasingly important role in the identification of Perkinsus species and requires adequate DNA sequence data at the targeted loci from the same and related species over a wide geographic area in order to develop reliable, accurate and sensitive molecular diagnostic tools  (Villalba et al 2004).

Culture: Examine tissues which have been placed in Fluid Thioglycollate Medium (FTM) for approximately 7 days for Lugol's iodine positive prezoosporangia (hypnospores), up to 250 µm in diameter (Ray's FTM, see Ray (1966) for details of this technique and Nickens et al. (2002) for an alternative formulation). Although not true propagating cultures, this procedure is used for the diagnosis of many species of Perkinsus but may also detect other organisms (Villalba et al. 2004) and thus can not be used to distinguish Perkinsus to species (Elandaloussi et al. 2008). The usual diameter of Perkinsus sp. prezoosporangia from FTM were reported as 30 to 40 µm from T. decussatus from Portugal (Azevado 1989) and 25 to 75 µm from P. undulata from Thailand (Leethochavalit et al. 2004). False negative diagnosis have occurred in 46%, 22% and 13% of infected clams (T. decussatus and T. philippinarum in Spain) in which only haemolymph, gills and the remaining body, respectively, were assayed by Ray's FTM (=thioglycollate diagnosis, Rodriguez and Navas 1995). However, Villalba et al. (2005) claimed that the examination of two gill lamellae from T. decussatus processed by Ray’s FTM was more sensitive, quicker and cheaper than examination of histological sections and although the examination of the whole-clam soft tissues by Ray's FTM allowed for the detection of very light infections, the correlation between the infection intensity estimated by both assays was high. For Perkinsus sp. in M. arenaria, Ray's FTM of gill and palp tissue is more sensitive than assaying either rectal tissue or haemolymph in light infections and more sensitive than histological examination. But, the use of both rectal and gill tissues in Ray's FTM was recommended (McLaughlin and Faisal 1999). Bushek et al. (2008) indicated that for P. chesapeaki, the difference in detection capabilities between Ray's FTM and PCR based detection assays related to the quality and type of tissues processed rather than assay sensitivity per se and further cautioned to use care when applying and interpreting diagnostic assays used on novel species.

Moore et al (2002) determined that trophozoites of P. olseni from A. trapezia made nonviable with formalin, irradiation or colchicine failed to swell in FTM and did not differentially stain in Lugol's iodine. However, trophozoites that had already developed into prezoosporangia in FTM and subsequently rendered inactive by freezing, ethanol or formalin retained their iodinophilic properties and could be used as a partial control for the FTM test. Choi and Park (1997) described a method of determining the number of Perkinsus sp. in V. philippinarum by digesting clam tissues from FTM in sodium hydroxide (2M NaOH), followed by washing (via centrifugation) and counting a subsample, stained with Lugol's iodine, using a haemocytometer. Almeida et al. (1999) also determined that lysis in 2M NaOH at 60 °C for 1 to 3 hours after incubating an entire minced or homogenized clams in FTM for 4 days followed by centrifugation to remove the NaOH supernatant and staining with Lugol's iodine solution was a quantitative diagnostic procedure. This procedure was more sensitive than histology for detecting low levels of infection (Almeida et al. 1999).

Optimal conditions for zoosporulation of prezoosporangia was in samples washed free of FTM and incubated in sea water at 24-28 °C, 25-35 ppt salinity, and pH 7-8. Ahn and Kim (2001) determined that temperature and salinity had significant effects on zoosporulation of Perkinsus sp. in V. philippinarum in Korea. Prezoosporangia isolated from clams during the winter sporulated and released motile zoospores at 10 °C and 5 parts per thousand salinity but, prezoosporangia isolated during the summer did not sporulate at 10°C and low salinities (10 parts per thousand or less) had a significant negative impact on development. Virus-like particles were observed in trophozoite-like cells isolated from gill tissues of R. decussatus that had been incubated in FTM (Azevedo 1990).

Perkinsus olseni (= atlanticus) in the haemolymph from the adductor muscle of R. decussatus was propagated in vitro and found to adapt to very different culture media, salinity (tolerance for 15 to 40 ppt) and temperature conditions (within extremes of 5 and 37 °C and optimums of 16 to 26 °C), and the inoculum density did not affect final cell concentrations attained (Ordás and Figueras 1998). La Peyre et al. (2008) determined that P. olseni declined in metabolic activity and proliferation from 28 °C to 15 °C and was viable after 30 days incubation at 4 °C, but had limited metabolic activity and no proliferation.  Robledo et al. (2002) developed an in vitro clonal culture of this parasite. The culture media described by Robledo et al. (2002) has been used to investigate the influence of specific drugs on the metabolic pathways of P. olseni (Elandalloussi et al. 2005). La Peyer et al. (2006) identified another nutrient medium that they used to determine the effects of salinity on viability, metabolic activity and proliferation of P. marinus, P. olseni and P. chesapeaki. Perkinsus sp. in haemolymph from M. balthica could also be propagated in nutrient medium as for P. marinus from Crassostrea virginica (Coss et al. 2001a, Arzul et al. 2009). Casas et al. (2002b) reported a low frequency of zoosporulation (<1% of dividing cells) in continuous cultures of P. olseni (=atlanticus) isolated from Tapes decussatus. Also, their cultured cells enlarged in FTM and stained blue-black with Lugol's iodine which is characteristic for this parasite from infected clams. Alternatively, Burreson et al. (2005) obtained in vitro isolates of Perkinsus sp. from prezoosporangia produced in FTM and reported that P. chesapeaki proliferated in cultures by both schizogony (multiple internal divisions in trophozoites enlarged to about 15 µm in diameter to produce clusters of sibling daughter trophozoites that subsumed the mother cell biomass, ~75% of dividing cells) and zoosporulation (a series of reductive divisions to yield hundreds of motile zoospores within a zoosporangium that were between 25 to 85 µm in diameter, ~25% of dividing cells). Zoospores released into the culture medium were motile for about 24 hours, shed their flagella, enlarged into typical trophozoites (about 9 µm in diameter with vacuolated, "signet-ring" morphology including an eccentric nucleus bearing a prominent nucleolus) and underwent subsequent schizogony or zoosporulation (Burreson et al. 2005). Dungan and Reece (2006) also used the same procedure and culture medium (850 mOsm/kg (29 ppt) DME:Ham’s F-12 Perkinsus sp. culture medium containing 3% (v/v) fetal bovine serum) as Burreson et al. (2005) to obtain isolates of Perkinsus spp. from V. philippinarum from Japan. Cryopreserved isolates of Perkinsus spp. are available at the American Type Culture Collection (ATTC, Rockville, MD, USA, www.atcc.org).

Note: Balseiro et al. (2010) compared three diagnostic techniques to detect P. olseni in various species of clams from Galicia, Spain. They determined that nested PCR was appropriate for rapidly screening large numbers of clams. It showed high sensitivity and good correlation between research groups, was faster than histopathology and Ray's FTM and less expensive than histopathology. Also, nested PCR required less specialized training for technicians than histology. Although Ray's FTM lacked analytical specificity and gave divergent results between research groups, particularly in the case of low levels of infection, it was useful for disease-monitoring purposes.

Methods of control

No known methods of prevention in enzootic areas. Areas of the world where Perkinsus sp, is not known to occur (e.g., the Pacific coasts of north and central America) must be especially diligent in screening seed from enzootic regions prior to importation for grow-out (Elston et al. 2003). Park et al. (2010) found that the fecal discharge (feces and pseudofeces) and decomposition of infected clam tissue could be the two major P. olseni routes of transmission. In France (Arcachon Bay), infection acquisition of P. olseni by V. philippinarum appeared to be episodic within spatially defined areas (Dang et al. 2010). In the management of perkinsosis, it must be kept in mind that at least for P. olseni (=atlanticus) in Spain, zoosporulation can occur in a wide range of temperatures (15 to 32 °C) and salinity (10 to 35 ppt) (with optimum values of 19 to 28 °C and 25 to 35 ppt), prezoosporangia survive up to 129 days at 10 °C, and zoospores survived for more than 20 days at various temperatures between 10 and 28 °C (Villalba et al. 2000). Goggin et al. (1990) determined that trophozoites of Perkinsus sp. in the tissues of blood cockles (Anadara trapezia) from Australia survived for at least 197 days at -60 °C. Cigarría et al. (1997) indicated that clam mortalities could be minimized by avoiding stressful conditions such as high densities, harvesting stress or overcrowding in depuration plants during the warmer months (water temperatures greater than 20 °C; apparently temperatures below 15 °C prevent Perkinsus sp. propagation in T. decussatus), as well as implementing two prophylactic measures of removing sets with parasitized clams and putting out (planting) unparasitised juveniles (seed) clams in aquaculture areas.

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Citation Information

Bower, S.M. (2011): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Perkinsus of Clams and Cockles.

Date last revised:  February 2011
Comments to Susan Bower