Marteilia refringens/maurini of Mussels


Category 1 (Not Reported in Canada)

Common, generally accepted names of the organism or disease agent

Marteiliasis of mussels.

Scientific name or taxonomic affiliation

Marteilia refringens and Marteilia maurini, in the phylum Paramyxea (Berthe et al. 2000). Unfortunately no morphological features are available to differentiate between M. refringens and M. maurini. 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 in some areas (Tige and Rabouin 1976, Le Roux et al. 2001) and common in others (Balseiro et al. 2007). From the results of molecular assays, López-Flores et al. (2004) and Balseiro et al. (2004) suggested that the Marteilia from oysters and mussels may be two different strains of the same species that appear to readily infect both hosts. The pathogenicity of each of the two species (if indeed both species are valid) is also unclear (Berthe et al. 2004). Balseiro et al. (2007) indicated that more research is required (e.g., into the molecular identity of new genes especially the coding genes, the relationship between the parasite and its different hosts, ecological studies to complete the life cycle, etc.) before the synonymy of the two species can be confirmed. 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).

Geographic distribution

Atlantic coast of Europe from southern United Kingdom to Portugal, Mediterranean Sea including the northern Adriatic Sea, Persian Gulf and the Gulf of Thermaikos in northern Greece; and maybe the east coast of Florida in the scallop Argopecten gibbus (possibly another marteiliad).

Host species

Mytilus edulis, and Mytilus galloprovincialis, as well as oysters, possibly clams and cockles, scallops and other bivalves (Berthe et al. 2004). Recently reported in the black pigmy mussel, Xenostrobus securis, a non-indigenous invasive species in the Ria de Vigo, Galicia, Spain where it cohabits with infected M. galloprovincialis (Pascual et al. 2010). Unpublished report in the horse mussel Modiolus modiolus (see Auffret and Poder 1983).

Impact on the host

This parasite is considered to be a potentially lethal pathogen. Mussels are usually not adversely affected by marteilioisis. However, in some areas, mortality attributed to this parasite is significant for the mussel farming industry. Berth (2002) indicated that mussels (M. edulis and M. galloprovincialis) from enzootic areas were susceptible to infection but not affected by (tolerant to) M. maurini, but naive M. edulis originating from an area free of Marteilia spp. experienced mass mortalities when transferred to the enzootic area. For example, mussel mortalities (up to 100%) associated with heavy infection by Marteilia maurini were reported in the past in France in Mytilus edulis bought in from Northern European countries for relaying in France. These mussels had no previous contact with M. refringens or M. maurini and were possibly highly susceptibility to the disease (Arzul and Joly 2011). Also, Fuentes et al. (2002) reported that hybrid crosses between M. edulis and M. galloprovincialis were more heavily parasitised than three M. galloprovincialis stocks cultured for one year under a commercial raft in northwest Spain. The average prevalence of infection in M. galloprovincialis from five rías in Galicia, NW Spain ranged up to 35% between 1985 and 1989 (Figueras et al. 1991, Villalba et al. 1997).

Figueras et al. (1991), Villalba et al. (1997) and Rayyan et al. (2006) noted that infection of M. refringens/maurini in mussels was associated with haemocyte infiltration of the digestive gland (connective tissue and epithelia) and extensive destruction of the digestive gland occurred in heavy infections. Pathologic disruption of the digestive gland tubule epithelia was most severe at the time of spore release from the sporangia. Heavy infections also caused reduction of absorption efficiency resulting in an inhibition of gonad and storage tissue development and thus a significant loss in condition of infected mussels (Robledo et al. 1995a). Infected M. galloprovincialis also had lower total carbohydrate concentrations in the haemolymph (Robledo et al. 1995a) and a significant increase in circulating haemocytes, especially hyalinocytes, (Carballal et al.1998) than uninfected mussels . The infection usually triggers a haemocytic reaction that may slow or even stop the infection at times. Association between mussel mortality and this parasite has been suggested (Villalba et al. 1993).

The complete life cycle of Marteilia refringens/maurini has not been fully described. Although Comps and Joly (1980) were able to infect mussels (Mytilus galloprovincialis) with M. refringens extracted from oysters (Ostrea edulis), Berthe et al. (1998, 2004) supported the hypothesis that intermediate or alternate hosts (unknown) or free-living stages (also unknown) were essential in the life cycle of this parasite. Research into potential intermediate hosts in French shallow-water oyster ponds (‘claires’), where species diversity is relatively limited, detected M. refringens DNA in various invertebrates with the most promising possibility being the Calanoida copepod Acartia (=Paracartia) grani (Audemard et al. 2001, 2002). Although A. grani could be infected with M. refringens/maurini from both oysters (O. edulis) and (M. galloprovincialis), 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. Subsequently, 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), 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 or gills. Carrasco et al. (2008c) detected initial (early) infections in the gill and mantle epithelium of M. galloprovincialis using the in situ hybridisation technique. 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 proliferated and eventually degraded. 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 occurs 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 mussel and infected mussels may shed large numbers of spores before death.

Figure 1.Schematic drawing to scale, of the sporulation process of Marteilia spp. with the cytoplasm of each stage colour coded for easy recognition. S = sporangiosorus (or primary cell, pink coloured cytoplasm), NS = nucleus of sporangiosorus, SP = sporangial primordium the matures into a sporont (the secondary cell, green cytoplasm), NSP = nucleus of sporangial primordium, St = sporont, SpP = spore primordium that matures into the spore (the tertiary cell, blue cytoplasm), MSp = mature spore, R = refringent bodies, N1 = nucleus of outer most sporoplasm, N2 = nucleus of middle sporoplasm, N3 = nucleus of inner most sporoplasm.

Diagnostic techniques

Gross Observations: Clinical signs include dead or gaping molluscs (2 or more years old), especially when temperature is at a maximum for mussels (Carrasco et al. 2007b, OIE 2009). Reduced growth rate and inhibition of gonad development were reported for infected mussels (Villalba 1993). However, these clinical signs are not specific to infection with M. refringens/maurini 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 mussels, freshly dead mussels or faeces of live diseased mussels. Squash a piece of digestive gland or faeces from suspect mussels on a glass slide. Observations are then made at ×400 magnification 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 walled structures (spores). In susceptible species, within the known geographic range of infection with M. refringens/maurini, a positive result is 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).

Smears/Tissue Imprints: In advanced infection, parasites ranging in size up to 30–40 μm can be observed in digestive gland imprints from gaping mussels or freshly dead mussels. Prepare digestive gland imprints of suspect mussels 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 ×200 magnification and then under oil immersion at ×1000 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: Various stages of the parasite (as described above) can be observed in the epithelial cells of the digestive gland ducts (basophilic stages, mainly nurse cells (plasmodia)) and the epithelial cells of the digestive tubules (acidophilic stages, mainly sporangiosori at various stages of development). Parasites in the primary and secondary digestive tubules are slightly smaller (mean 8.4 µm in diameter) than those in the major ducts and stomach (mean 9.9 µm in diameter with internal primary cells about 4.1 µm in longest axis). The sporangiosori, containing eight sporangial primordia (sporont), enlarge up to 19.6 µm in diameter and each sporangial primordia eventually contains 3-4 spores (mean 2.6 µm in diameter). Refringent granules appear during the course of sporulation and can range from deep orange to deep red in colour in tissues stained with haematoxylin and eosin stain.

Figure 2. Sporangiosori at different stages of development (arrows) and developing spores (arrowheads) of Marteilia maurini in digestive gland tubule epithelium of Mytilus edulis. Image from histological material supplied by I. Arzul of the OIE reference laboratory for Marteilia refringens. Haematoxylin and eosin stain.

Electron Microscopy: Only the outer sporoplasm of the spore contains haplosporosomes (130-400 × 130-200 nm). 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, Auffret and Poder (1983) and Longshaw et al. (2001) concluded that these criteria are 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).

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. Further details on the isolation of various developmental stages from the digestive gland of M. galloprovincialis are detailed by Robledo et al. (1995b).

Immunological Assay: Tiscar et al.(1993) prepared polyclonal antibodies to Marteilia sp. isolated from Mytilus galloprovincialis from southern Italy that was used to clearly mark the parasite by direct immunoperoxidase staining in smears of the digestive gland of infected mussels. Monoclonal antibodies from six clones obtained from mice (Balb/c) against Marteilia sp. from Mytilus edulis in Brittany, France were specific for Marteilia spp. and cross reacted with Marteilia refirngens from Mytilus galloprovincialis in Ria de Vigo, Spain. Four of the monoclonal antibodies reacted with the spore wall and two with the spore cytoplasm (Robledo et al. 1994a). These monoclonal antibodies were found to produce various results when used in different immunological tests (Pernas et al. 2000).

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 internal transcribed spacer 1 (ITS1) 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 that targete regions of the SSU rDNA, ITS1 and intergene 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 targets 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). Nevertheless, Zrnčić et al. (2001) used this probe to detect Marteillia sp. in Mytilus galloprovincialis in Croatia and Carrasco et al. (2008c) used it to detect the initial infective stages of M. refringens in M. galloprovincialis. In addition, an ISH assay was developed using a probe targeting the rDNA intergene spacer (IGS) (López-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 and developed a protocol of restriction fragment length polymorphism (RFLP) analysis applied to PCR products obtained using ITS1 primers that differentiates between Marteilia refringens and M. maurini. However, López-Flores et al. (2004) and Novoa et al. (2005) suggested that theses parasites were different strains of the same species.

Methods of control

Do not transfer mussels from areas known to be infected (currently or historically) to areas with no record of M. refringens/maurini. Mussels from areas where Ostrea edulis is known to carry Marteilia refringens should be treated with similar caution. Mussels from the inner part of two rías in Galicia, Spain, and those held at less depth (2 m rather than 5 m) in one ría had higher mean prevalence of infection. Thus, culture rafts located in the outer zones of the rías contribute to minimizing the impact of this parasite on the mussel culture industry in Galicia (Fuentes et al. 1995). In the Thermaikos Gulf, the prevalence of Marteilia sp. was significantly greater in mussels cultured on tables than on long-lines (Karagiannis and Angelidis 2007). Robledo et al. (1994b) suggested that the collection of mussel seed from areas free of Marteilia sp. may contribute to a reduction in the prevalence of the parasite in cultured stocks.


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Citation Information

Bower, S.M. (2011): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Marteilia refringens/maurini of Mussels.

Date last revised: July 2011
Comments to Susan Bower

Date modified: