Marteiliosis (Aber Disease) of Oysters

On this page

• Category
• Common, generally accepted names of the organism or disease agent
• Scientific name or taxonomic affiliation
• Geographic distribution
• Host species
• Impact on the host
• Diagnostic techniques
• Methods of control
• References
• Citation information

Category

Category 1 (Not Reported in Canada)

Common, generally accepted names of the organism or disease agent

Maladie des Abers, "Aber disease", Digestive gland disease, Marteiliosis.

Scientific name or taxonomic affiliation

The parasite Marteilia refringens Grizel et al. (1974a) described from Ostrea edulis and Marteilia-like parasites, are in the phylum Cercozoa and order Paramyxida (Cavalier-Smith and Chao 2003, Feist et al. 2009). However, these parasites have been assigned to various higher taxa in the past (Perkins 1976, Ormières and Grizel 1979, Lauckner 1983, Berth et al. 2004, Renault 2008). Although the taxonomic identity of M. refringens has been clearly established, a morphologically similar parasite in mussels (Mytilus galloprovincialis) from the Adriatic coast of Italy was named Marteilia maurini, (Comps et al. 1981 (1982), 1982). Subsequently, M. maurini was reported in M. galloprovincialis and Mytilus edulis as well as oysters (Ostrea edulis) from other locations in the Mediterranean basin and on the Atlantic coasts of Europe (for details see the webpage on Marteilia spp. of Mussels). Following the application of molecular tools, genetic differences were reported between the Marteilia isolates from oysters and mussels. However, the molecular signatures of each was detected in both oysters and mussels (Le Roux et al. .2001). López-Flores et al. (2004) suggested that the form most commonly found in oysters and the other most commonly found in mussels to be separate strains of one species M. refringens. Others have supported this hypothesis that types M and O (for oysters and mussels, respectively) represent the same unique species, M. refringens (Novoa et al. 2005; Balseiro et al. 2007; Arzul et al. 2014; Carrasco et al. 2015, 2017). 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 as defined by Le Roux et al. (2001) (OIE 2012, ICES 2012). Alfjorden et al. (2017) argued that they should conservatively be regarded as a single species, M. refringens, for most effective health management. However, based on molecular analysis, Kerr et al. (2018) provided a basis for reinstating two separate species and named the species, that they detected mainly in M. edulis and a few O. edulis from northern Europe, Marteilia parafringens. Although no morphological differences were reported between M. refringens and M. pararefringens, they described a means of discriminating these taxa based upon a specific molecular diagnostic assay (Kerr et al. 2018).

Other species of Marteilia occur in various species of oysters. During their revision of the order Paramyxida, Feist et al. (2009) proposed that the genus Marteilioides Comps, Park et Desportes (1986) be suppressed and the type species of the genus, M. chungmuensis Comps, Park et Desportes (1986) from oysters in Korea and Japan, be transferred to Marteilia. This proposal was supported by Itoh et al. (2014), Carrasco et al. (2015) and Alfjorden et al. (2017) but refuted by Ward et al. (2016). Because of difference in morphology and location of infection in oysters, M. chungmuensis will continue to be described on another webpage and not mentioned in the following text. Likewise, Marteilia sydneyi, from oysters mainly in Australia , is described on a separate webpage. Another species, Marteilia lengehi was described from Saccostrea (=Crassostrea) cucullata from the Persian Gulf and possibly Australia (Comps 1976, Hine and Thorne 2000, Carrasco et al. 2015) but little is known about this parasite. Another little known Marteilia-like species was reported in the epithelium of the digestive system of Saccostrea forskali from Chonburi Province on the east coast of Thailand with a low prevalence of infection (less than 7%) and no apparent association to mortalities (Taveekijakarn et al. 2002). However, haemocytic infiltration was found surrounding the infected area and extra sporogonic proliferation was observed in the gill of some of the infected oysters. This Marteilia sp. was characterised as having each sporangiosorus typically containing 2 to 6, or possibly more, sporonts and each sporont contained two spores (Taveekijakarn et al. 2002, 2008).

Geographic distribution

Marteilia refringens has been reported in Ostrea edulis from the Atlantic coast of Europe from southern United Kingdom to Spain and in the Mediterranean Sea (Alfjorden et al. 2017). However, a two year sampling program in the early 1990s indicated that O. edulis in Great Britain was free of this disease and although M. refringens was inadvertently introduced into coastal Netherlands after importation of infected oysters from France in 1974 (van Banning 1979a), it was not observed in the Netherlands after 1978 (Engelsma and Hine 2009b; Carrasco et al. 2017). Similarily, Kerr et al. (2018) did not detect O-type M. refringens in any northern European samples, including the United Kingdom, Norway and Sweden, by molecular and histological assays. Only M-type Marteilia were detected in a few O. edulis that were co-habiting with mussels infected by the M-type Marteilia, which they named M. pararefringens (Kerr et al. 2018).

In the Mediterranean Sea, Martielia refringens was reported in O. edulis from Morocco by the OIE in August 2006, in O. edulis and M.galloprovincialis from Diana Lagoon, Corsica (Arzul et al. 2014) and in Ostrea stentina from Tunisia (Elgharsalli et al. 2013). Marteilia sp. occurred in O. edulis, M.galloprovincialis and Modiolus barbatus in the Gulf of Thermaikos (Thessaloniki), northern Greece (Virvilis et al. 2003, Virvilis and Angelidis 2006).

Marteilia refringens has also been detected, using molecular assays (nested PCR), in Crassostrea gigas and Crassostrea corteziensis from the Gulf of California, Mexico (Grijalva-Chon et al. 2015). Marteilia pararefringens was detected in 1 of 150 O. edulis  from Tamar estuary, UK and 10 of 30 O. edulis from Bemlo, Norway where M. edulis was heavily infected (Kerr et al. 2018). Marteilia lengehi was described from Saccostrea (=Crassostrea) cucullata 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, Meyers 2006).

Host species

Initially described from oysters Ostrea edulis (Grizel et al. 1974a), Marteilia refringens and close relatives have since been reported in other species of oysters, mussels, clams, cockles, scallops (see Table 1 in Carrasco et al. 2015) and other non-bivalve invertebrates (as described below). In oysters, M.refringens was found in Ostrea chilensis (=Tiostrea chilensis =Ostrea lutaria), Ostrea angasi and Ostrea puelchana during experimental field trials in France (Grizel et al. 1982, Bougrier et al. 1986 and Pascual et al. 1991, respectively), in Ostrea denselamellosa (OIE 2012) and in Ostrea stentina (Elgharsalli et al. 2013). In addition, M.refringens was detected in a single 1 year old juvenile Crassostrea virginica introduced and reared in France (Renault et al. 1995) and in adult Crassostrea gigas  (Cahour 1979, Montes et al. 1998). There is speculation that the infection in C. gigas may be another marteiliad or only a transitory infection because C. gigas appears resistant to infection (Berthe et al. 2004) and M.refringens rarely develops to mature stages in C. gigas (Engelsma and Hine 2009a, Carrasco et al. 2015). Grijalva-Chon et al. (2015) detected M. refringens in C. gigas and Crassostrea corteziensis from the Gulf of California, Mexico. However, in both species of oysters the prevalence of infection was low, there was no associated mortality and the only assay used was nested PCR. As indicated by the authors, histology, in situ hybridization and transmission electron microscopy studies are necessary to determine if M. refringens has become established in the Gulf of California oyster (Grijalva-Chon et al. 2015).

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, ICES 2012). Marteilia refringens was also detected in a non-indigenous invasive mussel species Xenostrobus securis in the Ria de Vigo, Spain (Pascual et al. 2010). In addition to oysters and mussels, M. refringens has been reported in various species of clams and cockles as described on another webpage. Marteilia refringens has also been detected in other marine organisms that may serve as intermediate or alternate hosts (see below).

Impact on the host

Since 1968, M. refringens has caused serious recurring mortalities with a significant negative impact on the European Ostrea edulis industry (Comps 1970, Herrbach 1971, Balouet 1978, Alderman 1979, Grizel 1983, Englesma and Hine 2009b). However, severe mortalities with similar pathology have been reported in O. edulis in Europe prior to the 1960s (Mackin 1959). Grizel et al. (1974a) 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 2012). In most cases, M. refringens infection is associated with severe haemocyte infiltration surrounding the parasite in the primary and secondary digestive tubules of O. edulis  (Carrasco et al. 2015). However, Marteilia can occur in some oysters without causing disease (Grizel et al. 1974a, van Banning 1979b, Carrasco et al. 2015). The factors triggering a pathogenic host response are not clearly established, but may be related to environmental stresses, stock differences in disease resistance and/or pathogen virulence (Carrasco et al. 2015).

Development of M. refringens infections is directly related to high water temperature and low salinity (ICES 2012). Studies showed that 17 °C seems to be a trigger for parasite multiplication and transmission, followed by the disease outbreak (Audemard et al. 2001, Carrasco et al. 2015). Tige et al. (1979) reported that O. edulis in Britany, France were infected by M. refringens from the beginning of July to the end of August. Grizel and Tige (1979) determined that the period of infection varied between May and August and during the winter, the parasite seems to be eliminated from the tissues of O. edulis. However, recent studies revealed that, at least in Mediterranean waters where temperatures reach high values, M. refringens has two periods or cycles, for optimal multiplication and transmission during the year: spring / summer and autumn (Boyer et al. 2013, Carrasco et al. 2015). The highest prevalence appears to be in enclosed farming areas and M. refringens may have been transmitted around Europe in transfers of oysters and mussels via the commercial movements of stocks among growers in different regions (Guo and Ford 2016). Because this pathogen is not directly transmitted among oysters and mussels, potential, unknown, alternative host(s) would have had to be already present in the new locations or moved along with the bivalves (Guo and Ford 2016).

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 Paracartia (= Acartia) 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"). Arzul et al. (2014) also detected M. refringens in the ovarian tissue of Paracartia latisetosa from Diana Lagoon, Corsica. Marteilia refringens from oysters (O. edulis) and mussels (M. galloprovincialis) were infective to P. grani. However, attempts to transmit M. refringens from P. 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 P. 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 P. grani infected from mussels compared to higher prevalence and intensity of infections in the intestinal tract and gonad of P. grani infected from oysters. Although, involvement of P. 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 P. grani serving as an alternate host and not part of the life cycle remains to be determined (Berthe et al. 2004). Boyer et al. (2013) provided additional evidence that P. grani may be involved in the life cycle of M. refringens. Specifically, M. refringens was detected in the copepodid stages of P. grani between June and November by molecular assays (PCR and ISH) when the disease was active in associated M. galloprovincialis. In the third copepodid stage of P. grani, M. refringens was observed in the digestive tract and gonad suggesting that the parasite could infect a copepod by ingestion and be released through the gonad. Indeed, M. refringens DNA was detected in P. grani eggs obtained from PCR-positive females, which suggests that eggs could contribute to the parasite spreading in the water and could allow overwintering of M. refringens. Also, retention efficiency (number of copepods retained by a mussel) for all developmental stages of P. grani (especially eggs and nauplii) by M. galloprovincialis was up to 90%, further incriminating the involvement of P. grani in the life cycle of M. refringens (Boyer et al. 2013). However, experimental studies demonstrating the transmission of Marteilia spp. from infected copepods to uninfected molluscs is required in order to corroborate the involvement of copepods in the life cycle of these parasites (Carrasco et al. 2015).

Marteilia DNA has been detected in other marine organisms other than bivalves 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 P. grani. Carrasco et al. (2007a, 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. Burreson (2008) indicated that the PCR assay used by Carrasco et al. (2007a, b) was unvalidated to detect Marteilia spp. Because the PCR-only results from unvalidated assays were not confirmed by histology, they should be interpreted with caution and 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. (1974a, b), Perkins (1976), Grizel (1976, 1987), Franc (1980), Lauckner (1983) 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 epithelial cells of 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, endogenous budding) to produce cells within cells (Fig. 1). At the initiation of sporulation in M. refringens, 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, C. gigas, in areas enzootic for M. refringens.

Diagnostic techniques

Summary

There are no known morphological features that can be used to differentiate between M. refringens, M. maurini and M. pararefringens.Tissue imprints and histology of juvenile and adult oysters are recommended by the OIE (2012) as standard diagnostics methods especially for moderate to heavy infections in enzootic areas. However, Darriba Couñago (2017) emphasized that it is not always possible to identify species of Marteilia just by light microscope observation; unless specific identification is already stated with other techniques. Therefore, molecular analysis using specific polymerase chain reaction (PCR) assays that target the ITS1 region, of M. refringens is the method of choice in light infections (Le Roux et al. 2001), or under quarantine before the release of introduced stocks. PCR can also be used for targeted surveillance and presumptive diagnosis (OIE 2012, Aranguren and Figueras 2016). However, Burreson (2008) cautioned that a positive PCR result verifies an infection in a tested host only if the assay was properly validated for the geographic area and for the hosts examined. Also, molecular tools detect DNA sequences of the pathogen which does not imply that pathogen is viable in the host cell and the infection is established (Aranguren and Figueras 2016). Confirmatory diagnosis can be made by sequencing PCR products which can further be supported by in situ hybridisation (ISH) (Le Roux et al. 1999) or transmission electron microscopy (Englesma and Hine 2010a). However, ISH and/or ultrastructural criteria alone are not sufficient to discriminate between M. refringens/maurini (if they prove to be separate species) and various other Marteilia spp. (Arzul 2011, Carrasco et al. 2015).

Gross Observations

Clinical signs include dead or gaping molluscs (2 or more years old), especially at water temperature exceeding 17 °C (OIE 2012). Pale digestive gland, thin watery flesh, mantle retraction and reduced growth rate were reported for infected O. edulis and weak animals are particularly susceptible to any additional stress (Grizel et al. 1974a). However, these clinical signs are not specific to infection with M.refringens and could be indicative of other infections (Berthe et al. 2004, ICES 2012).

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 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 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 2012).

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 removing excess fluid from 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. 1974a, b; Berthe et al. 2000; Berthe et al. 2004; ICES 2012; for colour image see Arzul 2011 and Elgharsalli et al. 2013). 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 2012).

Histology

Cross-sections of infected oysters show the parasite ranging in size from 4 up to 40 μm (Darriba Couñago 2017). Young plasmodia (uninucleate) are mainly found in the epithelium of the stomach and gill lamellae or possibly the connective tissue of the mantle, labial palps or gills (Carrasco et al. 2015). Sporulation involves divisions of cells within cells and takes place in the epithelial cells of 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 (ICES 2012, Elgharsalli et al. 2013). The unique feature of internal cleavage to produce cells within cells during sporulation differentiates Paramyxida 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 and Grizel et al. (1974b) reported on the cell within cell arrangement of this parasite. 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 (1982)) indicated that M.refringens could be differentiated from Marteilia maurini in Mytilus galloprovincialis by subtle differences in haplosporosome shape and the 'existence of a multi-membraneous 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 2011). Elgharsalli et al. (2013) illustrated the ultrastructure of various stages of M. refringens in Ostrea stentina.

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). Microsporidian hyper-parasites have also been reported in M. refringens infecting M. galloprovincialis in Galicia, Spain (Villalba et al. 1993) and in Marteilia cochillia infecting Cerastoderma edule also from Galicia, Spain (Villalba et al. 2014).

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. Rodriguez (1997) used Percoll density gradients to obtain Marteilia sp. isolates for molecular analysis.

DNA Probes

In 1997, Rodriguez (1997) reported his success at the initial identification of a contig of 244 bp containing a segment of the small subunit ribosomal RNA (SSU rDNA or 18S rDNA) gene from Marteilia sp. Since then, the nucleotide sequence of the 18S rDNA gene was compared with that of various eukaryotic organisms and polymerase chain reaction (PCR) primers were designed (Le Roux et al. 1999). At that time, 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. Subsequently, this PCR assay was found to detect M. refringens, M. cochillia , M. sydneyi and probably other Marteilia spp. and may be better suited to detecting parasites in the genus Marteilia (Carrasco et al. 2015). Therefore, the World Organisation for Animal Health (OIE) recommends PCR primers that target the ITS1 (internal transcribed spacer) region described by Le Roux et al. (2001) to amplify M.refringens (OIE 2012). Le Roux et al. (2001) subjected the ITS1 region PCR products to restriction fragment length polymorphism (RFLP) with the endonuclease HhaI which generated two different profiles used to distinguish type "O" and type "M" of M. refringens. However, type "M" shows the same profile as M. cochillia from cockles (Cerastoderma edule) possibly resulting in misidentifications using this methodology (Carrasco et al. 2012). Carrasco et al. (2017) developed a competitive real-time PCR assay based on the ITS1 of M. refringens for rapid and sensitive detection of M. refringens which discriminated between "M" and "O" genotypes of M. refringens as well as the closely related M. cochillia. This real-time PCR assay was shown to be analytically sensitive and specific and has a high repeatability and efficiency (Carrasco et al. 2017). Kerr et al. (2018) designed and tested new Marteilia-specific PCR primers amplifying from the 3' end of the 18S rRNA gene through to the 5.8S gene, which specifically amplified the target region from both tissue and environmental samples and would differentiate between M. refringens and M. pararefringens.

In addition to analysis of ribosomal gene (rDNA) sequences, a nested PCR assay, that has been tested only with M.refringens from O. edulis and M.galloprovincialis from Huelva, SW Spain and two locations in NW Italy, was developed using primers targeting the rDNA intergene spacer (IGS) (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 2012). The RFLP-PCR on the IGS with BglII enzyme showed different profiles for M. refringens and M. cochillia (Carrasco et al. 2012). 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. False negative results can also occur due to the presence of inhibitors that may be present in the sample and could affect the activity of the DNA polymerase (Aranguren and Figueras 2016). 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 rDNA and 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 (18S) region of the rRNA gene complex have been validated against histology (Le Roux et al. 1999, Thébault et al. 2005). However, the probe, named SMART 2, was shown to cross react with Marteilia sydneyi and Marteilioides chungmuensis (Kleeman et al. 2002b). In addition, an ISH assay was developed using a probe targeting the rDNA intergene spacer (IGS) (López-Flores et al. 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 2011, ICES 2012).

Methods of control

Currently, the main method of control is the restriction of movements from infected areas to non-infected areas, as well as the harvesting or destruction of infected shellfish crops (Berthe et al. 2004, Carrasco et al. 2015).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 such as outer coast waters) to limit the development of M.refringens (Berthe et al. 2004, Alfjorden et al. 2017). The threshold temperature for parasite sporulation and transmission is 17 °C (Grizel 1987). This and warmer temperatures are 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 2012). 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, may be effective in controlling the disease (Bodoy et al. 1991, OIE 2012). However, Bodoy et al. (1991) and ICES (2012) claimed that any significant production of O. edulis in tidal saline ponds where M.refringens occurs could only been obtained with very short cycles of production, so as to reach a commercial size, before the contamination by M. refringens caused heavy mortalities. Nevertheless, 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. Few attempts to develop disease resistance to M. refringens by genetic selection have been made, although observations suggest that the biological basis exists (Berthe et al. 2004, Carrasco et al. 2015).

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Citation information

Bower, S.M. (2019): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Marteiliosis (Aber Disease) of Oysters

Date last revised: December 2020

Comments to Susan Bower