Marteiliosis (Aber Disease) of Oysters
Category 1 (Not Reported in Canada)
Common, generally accepted names of the organism or disease agent
Aber disease, Digestive gland disease of the European oyster, Marteiliosis.
Scientific name or taxonomic affiliation
Marteilia refringens Grizel et al. (1974), Marteilia -like parasites, in the phylum Paramyxea. However, this parasite has been assigned to various higher taxa in the past (Berth et al. 2004, Renault 2008). The World Organisation for Animal Health (Office international des épizooties, OIE) Reference Laboratory for Infection with Marteilia refringens recognises two types of Marteilia refringens, types O and M (for oysters and mussels, respectively) as defined by Le Roux et al. (2001) (OIE 2009).
Atlantic Europe from southern United Kingdom to Spain (although a two year sampling program in the early 1990s indicated that O. edulis in Great Britain was free of this disease) and reported in O. edulis from Morocco by the OIE in August 2006; Marteilia sp. occur in O. edulis and M.galloprovincialis in the Gulf of Thermaikos, northern Greece (Virvilis et al. 2003, Virvilis and Angelidis 2006) and in M.galloprovincialis on the north coast of the Adriatic Sea (Zrnčić et al. 2001); Marteilia sp. (possibly another marteiliad), was found in Argopecten gibbus from the east coast of Florida (Moyer et al. 1993). Marteilia lengehi was described from Saccostrea (=Crassostrea) cuccullata in the Persian Gulf (Comps 1976) and a similar Marteilia sp. was reported from the same species of oyster in northern Western Australia (Hine and Thorne 2000).
Initially described from oystersOstrea edulis (Grizel et al. 1974), this parasite and close relatives have since been reported in other species of oysters, mussels, clams, scallops and other invertebrates. In oysters, M.refringens was reported in a single 1 yr old juvenile Crassostrea virginica introduced and reared in France (Renault et al. 1995); in Crassostrea gigas (possibly another marteiliad or only a transitory infection, see Cahour (1979) and Montes et al. (1998) because C. gigas appears resistant to infection (Berthe et al. 2004)), as well as Ostrea chilensis (=Tiostrea chilensis and =Ostrea lutaria), Ostrea angasi and Ostrea puelchana by experimental field trials in France (Grizel et al. 1982, Bougrier et al. 1986 and Pascual et al. 1991, respectively), and Ostrea denselamellosa (OIE 2009). In mussels, M.refringens was reported from Mytilus edulis and Mytilus galloprovincialis (Tige and Rabouin 1976, Comps and Joly 1980, Villalba et al. 1993, Le Roux et al. 2001, Balseiro et al. 2007) and an unidentified Marteilia sp. in the horse mussel Modiolis modiolis (Auffret and Poder 1983). In clams,M.refringens or Marteilia sp. were reported in Cerastoderma (=Cardium) edule (Comps et al. 1975), Tapes pullastra and Tapes rhomboides (Poder et al. 1983), Ruditapes decussatus, Venerupis (=Ruditapes) philippinarum, Ensis minor and Ensis siliqua (Berthe et al. 2004, OIE 2009), Solen marginatus (López-Flores 2008a) and Chamelea gallina (López-Flores 2008b). In scallops, a Marteilia sp. was reported from Argopecten gibbus (possibly another marteiliad, Moyer et al. 1993). Berth et al. (2004) speculated that in the future, other species of bivalves may be found infected with various species of Marteilia. In addition, M.refringens has been detected in other marine organisms that may serve as intermediate or alternate hosts (see below).
Two other named species morphologically similar to M.refringens have been described: Marteilia maurini in Mytilus galloprovincialis and Mytilus edulis from the Atlantic coast of Spain, France and the Persian Gulf; and Marteilia lengehi from Saccostrea (=Crassostrea) cuccullata from the Persian Gulf and possibly Australia. Due to the near impossibility of distinguishing between these marteilias and M.refringens, the validity of these two species is questionable. However, Le Roux et al. (2001) identified genetic dimorphism in the internal transcribed spacer region of the ribosomal RNA genes and the apparent link of the two genetic types to host species, indicating that two species of Marteilia existed in Europe; M.refringens in oysters (O. edulis) and M.maurini in mussels (M. edulis and M.galloprovincialis). Nevertheless, co-infections of oysters and mussels by both genetic types occurred but were rare and only in areas where the prevalence of both types was high (Le Roux et al. 2001). Contrary to the conclusions of Le Roux et al. (2001), López-Flores et al. (2004) and Novoa et al. (2006) suggested that the Marteilia from oysters and mussels may be two different strains of the same species based on the results of their molecular analysis and both strains appeared to readily infect both hosts. A third species, Marteilia christenseni from a pelecypod (Scrobicularia piperata) at Ronce-les-Bains, France, can apparently be differentiated from the others by the characteristics of the cytoplasmic contents of the sporangia and the morphology of the spore (Comps 1985b).
Impact on the host
Since 1968, M.refringens has caused serious recurring mortalities with a significant negative impact on the European O. edulis industry (Comps 1970, Herrbach 1971, Alderman 1979, Grizel 1983). However, severe mortalities with similar pathology have been reported in O. edulis in Europe prior to the 1960s (Mackin 1959). Grizel et al. (1974) reported a 50–90% mortality rate during summer and autumn that was associated with sporulation of the parasite which occurs in the epithelial cells of the digestive tubules. More recently, M.refringens has been associated with the dramatic decrease in O. edulis production from the Gulf of Thermailos in northern Greece between 1994 and 1998 (Virvilis et al. 2003). Infection causes a poor condition index with glycogen loss (emaciation), discolouration of the digestive gland, cessation of growth, tissue necrosis, and mortalities. The digestive gland, in which M.refringens and other Marteilia species occur, is a site of intracellular food digestion and one of the main sites for storage of metabolic reserves (Berthe et al. 2004). Marteilia refringens may also interfere with glycogen storage in O. edulis (Robert et al. 1991). Severe infections may cause loss of condition as a consequence of reduced energy acquisition and the parasite may interfere with host feeding and absorption simply by its physical presence (OIE 2009). However, Marteilia can occur in some oysters without causing disease (Grizel et al. 1974). The factors triggering a pathogenic host response are not clearly established, but may be related to environmental stresses or stock differences in disease resistance.
Although Comps and Joly (1980) were able to infect mussels (Mytilus galloprovincialis) but not oysters (O. edulis) with M.refringens extracted from oysters (O. edulis), results of other experimental studies did not provide evidence to support direct horizontal transmission of M.refringens (Berthe et al. 1998). Audemard et al. (2001) indicated that intermediate or alternate hosts (unknown) or free-living stages (also unknown) were essential in the life cycle of this parasite, a hypothesis supported by Berthe et al. (2004). Audemard et al. (2002, 2004) employed molecular techniques to show that M.refringens occurs in the ovarian tissues of the Calanoida copepod Acartia (=Paracartia) grani and indicated that this copepod may be involved in the life-cycle of M.refringens, at least in French shallow-water oyster ponds (‘claires’). Marteilia refringens from oysters (O. edulis) and mussels (M. galloprovincialis) were infective to A. grani. However, attempts to transmit M.refringens from A. grani to O. edulis and mussels (Mytilus edulis) were unsuccessful (Audemard et al. 2002, Carrasco et al. 2008a). Also, Carrasco et al. (2008a) reported that the infection patterns of M.refringens/maurini in A. grani were different for copepods infected via M.galloprovincialis or via O. edulis with only early stages of infection found in the intestinal tract of A. grani infected from mussels compared to higher prevalence and intensity of infections in the intestinal tract and gonad of A. grani infected from oysters. Although, involvement of A. grani in the life cycle of M.refringens (at least in oysters) appears consistent with both the ecology of this copepod and the epidemiology of the disease, the possibility of A. grani serving as an alternate host and not part of the life cycle remains to be determined (Berthe et al. 2004). Marteilia DNA has been detected in other marine organisms by polymerase chain reaction (PCR). Specifically, Audemard et al. (2002) detected Marteilia DNA in 13 species including non-planktonic species, such as Cereus pendunculatus (Cnidaria) and Lineus gisserensis (Nematoda), from the French claire ponds for oyster aquaculture with the highest prevalence in A. grani. Carrasco et al. (2007a and b, 2008b) detected M.refringens DNA in other copepods (3 Calanoida, Acartia discaudata, A. clausi and A. italica; 1 Cyclopoida, Oithona sp.; and at least 1 Harpacticoida, Euterpina acutifrons and an unidentified Harpaticoida species) and in larval stages of decapod crustaceans (zoea larvae of Brachyura, probably Portumnus sp.) from the natural bays of the Ebro (Ebre) Delta (NW Mediterranean Sea, Spain) where mussels are the predominate farmed mollusc. The involvement of these organisms in the life cycle of M.refringens /maurini remains unknown.
The developmental stages of Marteilia spp. in bivalves were described by Grizel et al. (1974), Perkins (1976), Franc (1980), Grizel (1987) and Kleeman et al. (2002a), and summarized by Bower (2006) as follows. Infections by all Marteilia spp. are presumably initiated by a primary cell or stem cell (5 to 8 μm in diameter) in the epithelial cells of the gut (usually the stomach) and possibly the gills and labial palps. The primary uninucleate cell develops a secondary uninucleate daughter cell in a vacuole within its cytoplasm. The daughter cell divides by binary fission to produce four daughter cells within the enlarged primary (stem) cell and within each daughter cell a uninucleate cell develops by internal cleavage. The primary cell degenerates to release the daughter cells, which become new primary cells. In the gut, the parasite penetrates the basal membrane of the digestive gland tubules and becomes established as nurse cells at the base of the epithelial cells. Nurse cells containing daughter cells proliferate and eventually degrade. Daughter cells in the digestive gland tubules become sporangiosori called “primary cells” by Perkins and Wolf (1976) and pansporoblasts by Mialhe et al. (1985). Sporulation occurred within the sporangiosorus via a unique process of internal cleavages (endosporulation) to produce cells within cells (Fig. 1). At the initiation of sporulation, uninucleate segments become delimited within the cytoplasm of the sporangiosorus to form the sporangial primordia (secondary cells). Eventually, 8 to 16 sporangial primordia (each about 12 μm in diameter at maturity) form within the sporangiosorus that retains its nucleus and enlarges to about 30 μm in diameter. Each sporangial primordium matures into a sporont containing 2 to 4 spore primordia (tertiary cells) that mature into spores (Fig. 1). Each spore contains 3 uninucleate sporoplasms of graded sizes, with each of the smaller sporoplasms being enclosed within the cytoplasm of the next largest one (i.e., consecutive internal cleavage of two sporoplasms within the spore primordium) (Perkins, 1976). A continuous spore wall with no operculum occurs around each spheroid mature spore that measures 3.5 to 4.5 μm in diameter. As the spore matures, light refractile inclusion bodies appear in the sporont cytoplasm surrounding the spores. The specific name of M.refringens was derived from these 'refringent' inclusion bodies. Mature spores are shed into the tubule lumen for evacuation from the oyster and infected oysters may shed large numbers of spores before oyster death.
Initial stages of infection (without spore formation) was reported in the epithelium of the apical part of the stomach of C. gigas from Brittany, France (Cahour 1979) and Galicia, Spain (Montes et al. 1998). Montes et al. (1998) indicated that Marteilia sp. infections in C. gigas were transitory and not an obstacle for the culture of the Pacific oyster in areas enzootic for M.refringens.
Gross Observations: Clinical signs include dead or gaping molluscs (2 or more years old), especially at about 1 month after the water temperature exceeded 17°C (OIE 2009). Pale digestive gland, thin watery flesh, mantle retraction and reduced growth rate were reported for infected flat oysters and weak animals are particularly susceptible to any additional stress (Grizel et al. 1974). However, these clinical signs are not specific to infection with M.refringens and could be indicative of other infections (Berthe et al. 2004).
Wet mounts: In advanced infection, mature sporangia with refringent granules can be observed in wet mounts from gaping oysters or freshly dead oysters or faeces of live oysters. Squash a piece of digestive gland or faeces from suspect oysters on a glass slide. Observations are then made at x400 and can potentially show refringent granules in mature sporangia. A positive result is the presence of large (20–30 μm) spherical bodies containing spherical thick wall structures (spores). In susceptible species, within the known geographical range of infection with M.refringens/maurini, a positive result is indicative of infection with this parasite. In other species, or outside the known geographical range of infection with M.refringens/maurini, a positive result is indicative of infection with a Marteilia species that needs to be confirmed by the OIE Reference Laboratory (OIE 2009).
Smears/Tissue Imprints: In advanced infection, parasites ranging in size up to 30–40 μm can be observed in digestive gland imprints from gaping oysters or freshly dead oysters. Prepare digestive gland imprints of suspect oysters by drying excised tissues on absorbent paper and make several imprints on a glass slide. Air-dried slides are then fixed in methanol or in absolute ethanol and stained using a commercially available blood-staining kit, in accordance with the manufacturer’s instructions (e.g., Hemacolor, Merck; Diff-QuiK, Baxter). After rinsing in tap water and drying, the slides are mounted with a cover-slip using an appropriate synthetic resin. Slides are observed first at x200 magnification and then under oil immersion at x1000 magnification. Note that because infections may be focal and because the early and late stages of infection targets different tissues, imprints might miss early and low levels of infection. A positive result is the observation of cells ranging in size up to 30–40 μm in diameter with basophilic cytoplasm, eosinophilic nucleus, pale halos around large, strongly stained (refringent) granules and, in larger cells, cell within cell arrangements may be evident (Grizel et al 1974, Berthe et al. 2000, Berthe et al. 2004, for colour image see Arzul and Joly 2011). In susceptible species, within the known geographic range of infection with M.refringens/maurini, a positive result is strongly indicative of infection with this parasite. In other species, or outside the known geographic range of infection with M.refringens/maurini, a positive result is indicative of infection with a Marteilia species that needs to be confirmed by the OIE Reference Laboratory (OIE 2009).
Histology: Cross-sections of infected oysters show the parasite ranging in size from 4 up to 40 μm. Young plasmodia (uninucleate) are mainly found in the epithelium of labial palps and stomach. Sporulation involves divisions of cells within cells and takes place in the digestive gland tubules and ducts. Refringent granules appear during the course of sporulation, but are not observed in early stages. In late phases of infection, sporangia are observed free in the lumen of the digestive tract. Tissue sections stained with haematoxylin and eosin stain show the cytoplasm stained basophilic, the nucleus stained eosinophilic and the granules can range from deep orange to deep red. The unique feature of internal cleavage to produce cells within cells during sporulation differentiates Paramyxea including Marteilia spp. from all other protista. Values of sensitivity and specificity for histology were estimated at 70% and 99%, respectively (Thébault et al. 2005). A modified staining technique described by Gutiérrez (1977) may enhance the detection of the parasite in paraffin embedded histological sections.
Electron Microscopy: Bonami et al. (1971) described the osmophilic (refringent) bodies in the cytoplasm of M.refringens.Cytoplasmic organelles called haplosporosomes similar to those describe from Haplosporidia were report in M. refirngens (Perkins 1979, Comps et al. 1980). Comps et al. (1981) indicated that M.refringens could be differentiated from Marteilia maurini in Mytilus galloprovincialis by subtle differences in haplosporosome shape and the 'existance of a multimembraneous envelope next to the spore wall'. However, Longshaw et al. (2001) concluded that these criteria were invalid. Haplosporosomes in mature Marteilia from oysters and mussels were similar in shape (sphaeroid and oblate), although those from mussels were marginally smaller in size, and spore wall morphology was found to vary depending on the state of maturity of the parasite. Nevertheless, ultrastructural criteria are not sufficient to discriminate between Marteilia refringens and M.maurini (Arzul and Joly 2011).
Comps et al. (1979) described a Microspora hyper-parasite (Nosema ormieresi) in M.refringens from O. edulis collected in the Bay of Arcachon, France. This hyper-parasite caused necrotic changes in the primary cells and sporangia such as degeneration, membrane alteration, cytoplasm condensation and reduction in the number of spores. Although suggested as a potential biological control, this hyper-parasite has not been reported since it was first described making it apparently rare in the natural environment and with an unknown mode of transmission, the possibility of using it for biological control seems remote (Berthe et al. 2004).
Isolation and Purification: Mialhe et al. (1985) described a procedure for the isolation and purification of Marteilia sp. from oysters and mussels by the application of density gradients to a homogenate of heavily infected digestive gland.
DNA Probes: The nucleotide sequence of the small subunit ribosomal RNA (SSU rDNA or 18S rDNA) gene was compared with that of various eukaryotic organisms and polymerase chain reaction (PCR) primers were designed (Le Roux et al. 1999). The specificity of the amplified fragment for Marteilia sp. was confirmed by Southern blotting with an oligoprobe. Berthe et al. (2000) reported that the SSU rDNA gene sequence of Marteilia sp. that they isolated from O. edulis and M.edulis collected in different location in France were identical. Therefore, the World Organisation for Animal Health recommends PCR primers that target the ITS1 (internal transcribed spacer) region described by Le Roux et al. (2001) to amplify M.refringens (OIE 2009). Balseiro et al. (2007) analysed the ITS1 region subjected to restriction fragment length polymorphism (RFLP) with the endonuclease HhaI of Marteilia spp. from oysters and mussels from various locations in Europe to support the synonymy of M.maurini with M.refringens. In addition to ITS1 region analysis, a nested PCR assay, that has been tested only with M.refringens from O. edulis from Huelva, SW Spain and M.galloprovincialis also from Huelva, SW Spain and two locations in NW Italy, was developed using primers targeting the rDNA intergene spacer (López-Flores et al. 2004). This assay was demonstrated to be more sensitive than ITS1 PCR assay but needs to be tested more thoroughly for specificity (OIE 2009). As infection may be focal and also because infection targets different tissues in the early and late stages, the sensitivity of PCR detection may be lower than the expected theoretical PCR performance. A nested PCR reaction was developed for the diagnosis of M.refringens in Mytilus galloprovincialis from Galicia, Spain (Pernas et al. 2001). Sequences from regions of the SSU rDNA, ITS1 and intergenic spacer (IGS) are available in the public gene banks.
In situ hybridisation (ISH) protocols have been developed and published (Le Roux et al. 1999, Berthe et al. 2000). Probes that target the SSU of the rRNA gene complex and have been validated against histology (Le Roux et al. 1999, Thébault et al. 2005) are recommended by the World Organisation for Animal Health (OIE 2009). However, the probe, named Smart 2, was shown to cross react with Marteilia sydneyi and Marteilioïdes chungmuensis (Kleeman et al. 2002b). In addition, an ISH assay was developed using a probe targeting the rDNA intergene spacer (IGS) (Lopez-Flores 2008a, 2008b). This assay was demonstrated to be more specific than the SSU ISH assay but needs to be thoroughly validated. In situ hybridization can help to detect early infections which are more difficult to detect in traditional histological sections (Arzul and Joly 2011).
Analysis of segments of the small subunit ribosomal RNA genome in comparison with that of other eukaryotic organisms indicates that the phylum Paramyxea should continue to be recognized (Berthe et al. 2000). Le Roux et al. (2001) indicated that M.refringens and M.maurini are different species where as López-Flores et al. (2004) and Novoa et al. (2005) suggested that they were different strains of the same species.
Methods of control
Oysters from areas known to be infected (currently or historically) should not be transferred to areas with no record of M.refringens. Oysters from areas where M.galloprovincialis or M.edulis are infected by Marteilia spp. should be treated with similar caution. In enzootic areas, control is attempted by curtailing the planting of O. edulis seed during the period of transmission (July and August) and by growing O. edulis in areas with high salinities (35-37 ppt) and deeper water (subtidal/sublittoral areas) to limit the development of M.refringens (Berthe et al. 2004). The threshold temperature for parasite sporulation and transmission is 17°C (Grizel 1987). This temperature is common in estuaries or bays where prevalence is usually higher in the upper parts of the water column (Audemard et al. 2001, Berthe et al. 2004). Infection with M.refringens is seldom observed in open sea waters (OIE 2009). High salinity and water renewal could be detrimental to M.refringens development and transmission, although these parameters appear to be less significant than temperature (Audemard et al. 2001). Stocking at low density or in association with resistant mollusc species, such as the introduced Pacific oysters Crassostrea gigas, has been shown to be effective (OIE 2009). In addition, the increased production of C. gigas took the place of O. edulis farming (Grizel 1983). Comps (1975) reported that holding infected oysters out of water for 5 days seemed to reduce the intensity of infection when the oysters were returned to the water.
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Bower, S.M. (2011): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Marteiliosis (Aber Disease) of Oysters
Date last revised: July 2011
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