Haplosporidium pinnae sp. nov., a haplosporidan parasite associated with mass mortalities of the fan mussel, Pinna nobilis, in the Western Mediterranean Sea

Accepted Manuscript Haplosporidium pinnae sp. nov., a haplosporidan parasite associated with mass mortalities of the fan mussel, Pinna nobilis, in the Western Mediterranean Sea Gaetano Catanese, Amalia Grau, Jose Maria Valencia, Jose Rafael GarciaMarch, Elvira Alvarez, Maite Vazquez-Luis, Salud Deudero, Susana Darriba, María J. Carballal, Antonio Villalba PII: DOI: Reference:

S0022-2011(18)30103-4 https://doi.org/10.1016/j.jip.2018.07.006 YJIPA 7110

To appear in:

Journal of Invertebrate Pathology

Received Date: Revised Date: Accepted Date:

22 March 2018 5 July 2018 7 July 2018

Please cite this article as: Catanese, G., Grau, A., Maria Valencia, J., Rafael Garcia-March, J., Alvarez, E., VazquezLuis, M., Deudero, S., Darriba, S., Carballal, M.J., Villalba, A., Haplosporidium pinnae sp. nov., a haplosporidan parasite associated with mass mortalities of the fan mussel, Pinna nobilis, in the Western Mediterranean Sea, Journal of Invertebrate Pathology (2018), doi: https://doi.org/10.1016/j.jip.2018.07.006

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Haplosporidium pinnae sp. nov., a haplosporidan parasite associated with mass mortalities of the fan mussel, Pinna nobilis, in the Western Mediterranean Sea GAETANO CATANESE1,2,*, AMALIA GRAU1,2, JOSE MARIA VALENCIA1,2, JOSE RAFAEL GARCIA-MARCH3, ELVIRA ALVAREZ4, MAITE VAZQUEZLUIS4, SALUD DEUDERO4, SUSANA DARRIBA5, MARÍA J. CARBALLAL6, ANTONIO VILLALBA6,7,* 1

Laboratori d’Investigacions Marines i Aqüicultura, (LIMIA) - Govern de les Illes Balears, Av.

Gabriel Roca 69, 07158 Port d’Andratx - Mallorca (Spain). 2

Instituto de Investigaciones Agroambientales y de Economía del Agua, (INAGEA) (INIA-CAIB-

UIB). Ctra. Valldemossa km. 7,5 Ed. Edifici Guillem Colom Casasnoves, 07122 Palma de Mallorca - Illes Balears 3

Instituto de Investigación en Medio Ambiente y Ciencia Marina (IMEDMAR). Universidad

Católica de Valencia. Avenida Puerto Pesquero s/n, 03710 Calpe (Spain) 4

Instituto Español de Oceanografía (IEO). Centro Oceanográfico de Baleares. Muelle de Poniente

s/n, 07015 Palma de Mallorca, Spain, 5

Instituto Tecnóloxico para o Control do Medio Mariño de Galicia (INTECMAR), Consellería do

Mar, Xunta de Galicia. Peirao de Vilaxoán s/n, 36611 Vilagarcía de Arousa, Spain 6

Centro de Investigacións Mariñas (CIMA), Consellería do Mar, Xunta de Galicia, Pedras de

Corón s/n, 36620 Vilanova de Arousa, Spain, 7

Departamento de Ciencias de la Vida, Universidad de Alcalá, 28871 Alcalá de Henares, Spain

G. Catanese and A. Grau contributed equally to this work

* Corresponding authors: Laboratori d’Investigacions Marines i Aqüicultura, (LIMIA) Email: [email protected] Centro de Investigacións Mariñas, Consellería do Mar, Xunta de Galicia. Email: [email protected]

Keywords: , endangered species, histology, electron microscopy, phylogenetic analysis, mass mortality

Abstract This study provides morphological and molecular characterization of a new species, Haplosporidium pinnae, very likely responsible for mass mortality of fan mussels, Pinna nobilis, in the Western Mediterranean Sea. The parasite was found in dead or moribund P. nobilis but did not occur in healthy fan mussels from locations that were not affected by abnormal mortality. Histological examination of infected fan mussels showed uninucleate cells of a haplosporidan parasite throughout the connective tissue and hemolymph sinuses of the visceral mass and binucleate cells and, rarely, multinucleate plasmodia were also detected in the connective tissue. Additionally, stages of sporulation occurred in the epithelium of the host digestive gland tubules. Spores were slightly ellipsoidal with a hinged operculum in one pole. Typical haplosporosomes were not found with TEM but vesicles with two concentric membranes resembling haplosporosomes were abundant in the cytoplasm of the multinucleate plasmodia occurring in host digestive gland tubules. SEM analysis showed multiple structures on the spore surface; some spores had two or four long tape-like filaments attached to the spore wall. Phylogenetic analysis based on the SSU rDNA sequence placed this parasite within a large clade including species of the order Haplosporida, not in the Bonamia/Minchinia subclade or the subclade containing most Haplosporidium species, but within a subclade of Haplosporidium sp. from Penaeus vannamei. Our results suggested that H. pinnae and the parasite of P. vannamei may represent a distinct new genus within the order Haplosporida.

1. Introduction The fan mussel Pinna nobilis (Linnaeus, 1758), is the largest endemic bivalve in the Mediterranean Sea. It occurs at depths between 0.5 and 60 m, mostly on soft-bottom areas overgrown by seagrass meadows, but also occasionally on bare sandy substrate and maërl beds (Garcia-March et al., 2002; Katsanevakis, 2007; Zavodnik et al., 1991). P. nobilis filters large amounts of detritus and retains a high percentage of organic matter, an important ecological role in the system (Trigos et al., 2014), contributing to water clarity. It also provides a hard substrate in soft-bottom areas, increasing habitat variability and providing a surface that can be colonized by other benthic species. P. nobilis populations greatly declined in the 20th Century due to anthropogenic activities, including recreational and commercial fishing, ornamental harvesting, and accidental killing by anchoring, bottom nets and trawlers (Vicente and Moreteau, 1991; Richardson et al., 2004; Katsanevakis, 2007; Hendriks et al., 2013; Deudero et al., 2015; Vázquez-Luis et al., 2015). This species is legally protected under Annex II of the Barcelona Convention (SPA/BD Protocol 1995), Annex IV of the EU Habitats Directive (European Council Directive 92/43/EEC), and the Spanish Catalogue of Threatened Species (Category: Vulnerable, Royal Decree 139/2011). A mass mortality event (MME) impacting Pinna nobilis was detected across a wide geographical area of the Spanish Mediterranean Sea (Western Mediterranean Sea) in early autumn 2016, affecting specimens of all sizes, depth ranges and habitat types. Mortality reached 100% in the south and central Mediterranean coasts of the Iberian Peninsula populations while the northern coasts of the Spanish Mediterranean Sea seemed to be unaffected at that time (Vazquez-Luis et al., 2017). Unfortunately, severe mortality has been observed recently in Pinna nobilis from Catalonia (Spain), Ischia (Italy), Sicily (Italy) and Corsica (France) coasts, which suggests the arrival of an MME to these localities (Elisabet Nebot, Francesco Patti, Salvatore Giacobbe, Christine PergentMartini pers. comm. and citizen science/ http://www.observadoresdelmar.es). In consequence, 17th July 2017, the Spanish Sectoral Environmental Conference approved changing the species status at the national level from “Vulnerable” to “Critically Endangered”. This declaration requires that the research and projects aimed at the recovery of this species are of general interest and implementation will be of an emergency nature (Article 60.2 of Law 42/2007, Natural Heritage and Biodiversity. The congeneric species Pinna rudis, shares habitat with P. nobilis in many of the areas affected by the MME but has not been affected by abnormal mortality (Vázquez-Luis et al., 2017). A haplosporidan parasite has been reported as the likely cause of the fan mussel mass mortality (Darriba, 2017; Vázquez-Luis et al., 2017). Haplosporidans parasitize marine and

freshwater invertebrates globally and can be highly pathogenic. Among them, Haplosporidium nelsoni was the responsible for mass mortalities of Crassostrea virginica on the east coast of the USA (Ford and Tripp, 1996) and two species, Bonamia ostreae and Bonamia exitiosa, have caused extensive mortality in populations of various oyster species (Engelsma et al., 2014). Both Bonamia species are pathogens listed by and notifiable to the World Organization for Animal Health (OIE) and the European Union (EU). In addition, unnamed haplosporidan parasites were responsible for high mortality of cultured shrimps Penaeus vannamei in the Caribbean Sea (Nunan et al., 2007) and Indonesia (Utari et al., 2012). The protist order Haplosporida includes over 50 described species in four genera: Haplosporidium, Minchinia, Bonamia and Urosporidium (Arzul and Carnegie, 2015; Azevedo and Hine, 2017). Life stages include uninucleate cells, plasmodia, and spores. Spore morphology, particularly the external ornamentation, is used to differentiate among genera (McGovern and Burreson, 1990). Haplosporidan spores are ovoid with an orifice at one end. The orifice is closed by internal plugs of spore wall material in the species of genus Urosporidium, whereas the spores of genera Haplosporidium and Minchinia, as well as the species Bonamia perspora, have hinged caps or opercula covering the orifice. Spore ornamentation originates from the spore wall in Haplosporidium spp. and B. perspora but, in Minchinia spp., originates from epispore cytoplasm, then disappears in the mature stage (Burreson and Ford, 2004; Arzul and Carnegie, 2015). Phylogenetic studies showed that Minchinia spp., Bonamia spp. and Urosporidium spp. form monophyletic clades and the genus Haplosporidium constitutes a paraphyletic clade (Burreson and Reece, 2006; Hartikainen et al., 2014). Fan mussel sampling was performed along the Mediterranean coast of the Iberian Peninsula, including Balearic Islands, during the MME to search for its cause; standard histology and molecular procedures (PCR and DNA sequencing) were used to detect the haplosporidan parasite. This article describes a new haplosporidan species, Haplosporidium pinnae, based on the characterization of the morphology and molecular phylogeny of this fan mussel parasite.

2. Materials and Methods 2.1 Sampling Due to status as an endangered and protected species, sampling of P. nobilis and P. rudis was carried out under permission of regional and national Authorities (Govern de les Illes Balears, Generalitat Valenciana, Generalitat de Catalunya, Junta de Andalucía, Región de Murcia; and Ministerio de Agricultura y Pesca, Alimentación y Medio Ambiente-MAPAMA). Weak or

moribund specimens (lacking response to stimuli and slow valve closing) and healthy specimens were collected by scuba divers over a depth range of 0.3-36 m in the selected areas, which cover large parts of Spanish Mediterranean coastal areas where MME has been detected (Fig. 1). Both diseased and apparently healthy fan mussel samples were collected in the affected areas, with a goal of detecting the etiological agent of the MME by histological and molecular analyses. In addition, one individual was taken from each of two areas (Mar Menor and Alfacs) that had not been affected by abnormal mortality at the sampling time. A total of 43 specimens of P. nobilis were collected from October 2016 to November 2017 for this study: Marina Real (Valencia; n = 7), Port d’Andratx (Balearic Islands; n = 23), Cabrera (Balearic Islands; n = 2), El Calón (Almeria; n = 3), Son Saura and Sa Farola (Minorca-Balearic Islands; n = 2), Tossa de Mar (Girona; n. = 4), Mar Menor (Murcia; n = 1) and Alfacs (Tarragona; n = 1). Four fan mussels collected from Calpe (Alicante) that were used in studies by Darriba (2017) were also included in this study for molecular characterization of the parasite. (Fig. 1; Table 1). Additionally, two specimens of the spiny fanmussel, Pinna rudis, from the Cabrera National Park, were collected in the same zone of high P. nobilis mortality to determine if this species could be carrier of the parasite. One specimen was apparently healthy while the other could not close its valves, an unspecific indication of disease. Visceral tissues of the individuals included in these samples were fixed for histological purposes, and small portions of different tissues were fixed in absolute ethanol for molecular studies (see below). In addition, tissue samples were collected from 19 apparently healthy crustacean symbionts, Pontonia pinnophylax (both male and female), 3 Nepinnotheres pinnotheres sheltered within fan mussels, 1 healthy abalone, Haliotis tuberculata, and 2 common limpets, Patella vulgate, found adhered to the shells of two affected fan mussels. Tissue samples of two predators, the snail Hexaplex trunculus and the starfish Echinaster sepositus, both captured devouring moribund fan mussels at the time of the sampling in Sa Farola and in Port d’Andratx, respectively, were processed as well. Individuals with abnormal appearance or behavior at first inspection, namely those that were gaping and presenting mantle retraction, lacked response to stimuli, and valve closing was slow, were transported to the laboratory under cool conditions within 24 hours or were dissected and sampled “in situ”. All specimens were examined in order to evaluate vitality, and were measured and weighed. The presence or absence of symbionts and other information relevant to the search for the etiological agent of the MME were recorded and samples were taken for histopathological, SEM, TEM and molecular characterization.

2.2 Histological analysis

The 43 fan mussels were fixed in phosphate buffered formalin or in Davidson’s fixative for routine histological purposes (Table 1). A cross section of the visceral mass of each processed bivalve mollusk was taken at the level of the digestive gland, dehydrated in an increasing ethanol gradient, cleared with Microclearing, embedded in paraffin wax, sectioned at 3-4 μm, and stained with Mayer’s hematoxylin and eosin (MHE) for routine light microscopic examination. Some additional sections were stained with Harris’ hematoxylin and eosin (HHE) (Howard et al., 2004) and others with Mayer’s hematoxylin and Verde Luz-orange G-acid fuchsine (MH-VOF) (Gutiérrez, 1967). Gram staining was also performed to detect bacteria. A longitudinal section of crustacean symbionts (16 Pontonia pinnophylax, both male and female), two Nepinnotheres pinnotheres) and the two predators also were processed following the same procedure.

2.3 Ultrastructural analysis For transmission electron microscopy (TEM) analysis, small fresh samples of infected tissue from the digestive gland of P. nobilis were fixed in 4% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) for 2 h at 4°C, washed for 24 h at 4°C in the same buffer and post-fixed in buffered 2% OsO4 (0.1 M) for 2 h at room temperature. After washing in cacodylate buffer, samples were dehydrated in a graded ethanol series, washed in propylene oxide and embedded in Epon resin. Ultrathin sections were double stained with uranyl acetate and lead citrate, and observed using a JEOL JEM1010 TEM operating at 80kV. For scanning electron microscopy (SEM) analysis, a piece of digestive gland of an infected specimen that had been preserved in 96% ethanol for molecular analysis was used; it was rehydrated and sonicated to release spores. Sonication was performed with a 450 Sonifier (Branson), holding the tissue piece in distilled water within a vial on crushed ice, operating at duty cycle 30 (ultrasonic pulses of 0.3 seconds, one pulse per second) and the amplitude control at position 2, for 2 minutes; then a drop of liquid was examined with a light microscope and the process was repeated until satisfactory spore separation was attained. Isolated spores were dehydrated in a graded ethanol series, critical point-dried and sputter-coated with gold. Observations and image acquisitions were made using a FEI Quanta 200 SEM operating at 15 kV.

2.4 Molecular analysis For molecular analyses, organs commonly infected by the parasite (digestive gland, adductor muscle and mantle), as observed in early histological analysis, were dissected and homogenized. We analyzed 33 samples of P. nobilis, representatives of all sampling sites (Table 1), 2 of Pinna rudis, 3 of Pontonia pinnophylax and 1 of Nepinnotheres pinnotheres. Fourteen samples of a total

of 23 from Port d’Andratx were used only for histological analysis because tissues had been fixed with formaldehyde. Total genomic DNA was purified using the Macherey-Nagel DNA Tissue extraction kit following the manufacturer’s instructions. A fragment of about 300 bp of the small subunit ribosomal DNA (SSU rDNA) gene was initially amplified using the primers HapF1 and HapR2 (Table 2) and PCR conditions described by Renault et al. (2000) for Haplosporidium spp. We then designed two new forward primers HPNF1 and HPNF3, using as reference the sequence of conserved regions of similar genera and a specific reverse primer, HPNR3, designed from the sequences obtained by previous amplification with HapF1/HapR2. PCR using primer pairs HPNF1/HPNR3 and HPNF3/HPNR3 generated amplicons with length of about 1.2 kb and 600 bp, respectively. We used the primer HPNF3 paired with the reverse 16Sb (Medlin et al., 1998) to amplify a fragment of about 1 kb, corresponding to the 3’ end of the SSU rDNA. All PCR fragments were bi-directionally sequenced using ABI 3130 Genetic Analyzer (Applied Biosystems, Carlsbad, CA, USA). PCR reactions were performed in a total volume of 20 μl containing 0.8 μl 10 mM dNTP mix, 1 U Taq DNA polymerase (Biotools), 2 μl PCR Reaction Buffer (+Mg) 10×, 0.5 μl of each primer (stock 20 uM) and 1 μl of DNA between 60 and 200 ng/μl. The temperature profile followed an initial denaturation at 94 ºC for 10 min; 35 cycles of 94 ºC for 1 min, 49-56 ºC for 1 min (depending on the primers pairs; Table 2) and 72 ºC for 1 min, and a final extension at 72 ºC for 10 min. PCR products were separated on 1% agarose in TAE 1× buffer gels (w/v), stained with GelRed (Biotium) including a LowRanger 100-bp DNA ladder size standard (Norgen) and visualized on UV transilluminator. Obtained sequences were edited and aligned using the BioEdit v7.2.5 software (Hall, 1999) and MEGA 6.0 (Tamura et al., 2013). The pairwise nucleotide differences and the detection of different haplotypes were obtained by DNAsp v5.0 software (Librado and Rozas, 2009). Sequences of the SSU rDNA from GenBank of other haplosporidan species as well as other protozoa within Rhizaria and Alveolata protozoan groups were added to the alignment and employed to build Maximum likelihood (ML), neighbor-joining (NJ) and Bayesian Inference (BI) trees, using a representative species of Mikrocytos mackini as an outgroup. JModelTest v2.1.7 (Darriba et al., 2012) using the Akaike Information Criterion (AIC; Posada and Buckley, 2004) was used to select the appropriate model of evolution, as a guide to determine the best-fit maximum likelihood model. MEGA 6.0 software (Tamura et al., 2013) was adopted to reconstruct the phylogenetic relationships building ML and NJ trees, with 1,000 bootstrap replicates. In addition, a tree with Bayesian inference methods was created using MrBayes v. 3.2 (Ronquist et al., 2012) with 2,000 replicates. Pairwise Nei's genetic distances (Nei, 1972) were calculated in MEGA 6.0.

3. Results 3.1 Gross observations The total length of 43 fan mussels examined in this study ranged between 22-71 cm and total weight 82-1290 g (Table 1). A wide range of epibionts were observed attached to the valves: polychaetes, bryozoans, red coralline and green algae, sponges, small common limpets (Patella vulgata), empty oyster shells, and one live abalone Haliotis tuberculata. At necropsy, disease indications were not specific: i) the specimens were emaciated, with abundant watery large vesicles on the visceral mass in acute clinical cases; ii) the gills were collapsed and appeared with a pale brownish coloration; iii) the digestive glands were darker and softer than the normal appearance and sometimes presented liquefaction. Typically, in heavily affected individuals the crystalline style as well as the byssus were not present. In moribund specimens, crustacean symbionts were absent indicating imminent death of the host. Crustacean symbionts were observed in 16 out of 32 samples processed at laboratory. Pontonia pinnophylax were found in pairs (n=26 individuals), whilst the crabs Nepinnotheres pinnotheres were observed individually (n=3). Crustacean symbionts were absent in healthy fan mussels collected in areas with lower and higher salinities than the Mediterranean Basin (Alfacs and Mar Menor, respectively) as well as in the two sampled spiny fan-mussels P. rudis. Four individuals were dead when they were dissected for necropsy as deduced from advanced degradation signs, with marked watery aspect of the visceral mass.

3.2 Histological analysis Histological examination revealed the presence of a haplosporidan-like parasite in 37 out of 43 analyzed P. nobilis, particularly within the digestive gland. Uninucleate stages were the most abundant stage of this parasite in the connective tissue throughout all the visceral mass, including hemolymph sinuses. Their morphology varied from spherical to elongated, 1.8–4.6 μm maximum length (mean =3.2; SE = 0.02; n = 50) (Fig. 2A); they showed eosinophilic cytoplasm and the nucleus appeared usually condensed, although nuclei with ring shape (clear center) were less frequently observed. Binucleate stages (Fig.2B) and, rarely, plasmodia with more (up to 6) nuclei were also observed in the connective tissue. Some parasite cells were seen within host hemocytes, without degradation signs (Fig. 2C) and all these stages were occasionally observed in the gut epithelium (Fig. 2D). Additionally, sporulation stages occurred in the epithelium of digestive gland tubules; the following parasite stages were observed. Uni- and binucleate cells, similar to those observed in the connective tissue, accumulated in high numbers in the epithelia of digestive gland

tubules (Fig. 3A). Plasmodia with light eosinophilic cytoplasm and up to 10 nuclei/section were observed (Figs. 3A, 3B). Larger, deeply stained plasmodia with nuclei sometimes greater than 50/section were most abundant (Fig. 3B); compartmentalization of the cytoplasm was more or less evident, demarcating uninucleate areas in the clearer cases. Abundant sporocysts enclosing numerous (up to 50 or more in section) uninucleate sporoblasts with a more or less distinguishable wall, depending on the degree of maturity. The size of sporocysts was approximately 30 µm but estimating individual size was difficult because they frequently appeared tightly packed (Figs. 3C3E). Relatively mature spores with a patent wall, an operculum visible in those with favorable position, and one central nucleus were observed in sporocysts or less frequently free within the epithelium of the host digestive gland tubules; their shape varied from ellipsoidal to angulated, 3.6 – 5.4 µmin length (mean = 4.3; SE = 0.12; n = 30) and 2.7 – 4.5 µmin width (mean = 3.5; SE = 0.11; n = 30) (Fig. 3E). The endosporoplasm shrank as spores matured leaving a growing halo between the wall and the sporoplasm. The progression of sporulation was easily observed with VOF staining, because the transition from early sporoblast to mature spore was associated with color change from blue to yellowish-orange; the more mature stage, the darker the orange color (Fig. 3E). In advanced infections, the epithelium of all digestive gland tubules was almost completely occupied by stages of the parasite. The epithelium height of digestive gland tubules was reduced, resulting in a wider lumen (Figs. 3D, 3E); large vacuoles enclosing amorphous eosinophilic material were frequently observed in the epithelium of infected digestive gland tubules (Figs. 3A, 3B). A heavy inflammatory host response was associated with infection, with serious infiltration of the connective tissue of the digestive gland by host hemocytes (Fig. 4A) and abundant brown cells (Fig. 3D). Abnormal hemocytic infiltration did not occur in the digestive gland of non-infected fan mussels (Fig. 4B) and brown cells were less abundant. Their digestive gland tubules showed taller epithelium with numerous cytoplasmic brownish granules (Fig. 4C), both typical features of the absorptive phase. Sporulation stages were observed from October 2016 to March 2017 in specimens from different localities affected by the outbreak at that time (Almeria, Valencia, Mallorca and Cabrera), but not in the rest of the samples (Valencia, Menorca, Tossa de Mar). In the latter samples, the parasite occurred in the connective tissue and the gut epithelium, where the uninucleate and the less frequent plasmodial stages again were associated with a heavy inflammatory response. Remarkably, heavily sick individuals showed digestive epithelium damage although sporulating stages were not observed in the digestive epithelia. No other parasites, prokaryotes or eukaryotes, were detected that would produce the observed lesions. Ciliates were more or less abundant, depending on the specimen, in the gills of weak and dying individuals.

The haplosporidan-like parasite was not detected in 6 out of 43 fan mussels histologically analyzed in this study (Table 1). Four included the mussels that were dead at necropsy, and with a high degree of post-mortem autolysis and gram-negative bacterial contamination observed by histology, thus hampering diagnosis. The parasite was not detected in the two P. nobilis collected from healthy populations located in areas of higher and lower salinity than in the affected areas (Mar Menor and Alfacs Bay, respectively). Moreover, the parasite was not detected in the two spiny fan mussels P. rudis, which showed a normal morphology of their digestive gland and adjacent connective tissues, nor was it observed in any of the examined symbionts, epibionts and predators. All epibionts and predators had normal gross morphological appearance, as well as a normal histological structure of their digestive glands and connective tissues, without any apparent lesions.

3.3 Ultrastructural observations The earliest stage of the parasite detected with TEM corresponded to multinucleate plasmodia, round to elongated in shape with 10-20 µm maximum length, located in the epithelium of digestive gland tubules (Fig. 5A). Nuclei appeared frequently paired, with up to 3.5 µm in diameter, occasionally with nucleolus. The observation of an evagination of the envelope of one paired nucleus from which the outer membrane appeared inflated (inset in Fig. 5A) located near vesicles with similar size to that of the inflated evagination end (210 nm in diameter), suggested the release of vesicles from the nuclei. Numerous vesicles occurred in the cytoplasm; among them, round to ovoid vesicles with an outer membrane and a concentric inner membrane and ranging in diameter from 260-480 nm (mean = 340.1; SE= 17.45; n = 30) were particularly abundant (Figs. 5A-5C). The marginal area between the outer and the inner membrane was electron-dense while the inner core was lighter, even electron-lucent (Fig 5B). A typical Golgi apparatus with piled flattened cisternae was not observed, although numerous tubular cisternae (and their circular sections) of smooth endoplasmic reticulum with dense content, were observed anastomosing in areas with abundant vesicles with two concentric membranes (Fig. 5C). Lipid droplets were frequent and inflated mitochondria with few tubular cristae and free ribosomes also occurred in the cytoplasm (Fig. 5A). Sporonts were observed enclosing sporoblasts in which two areas were distinguished, the inner developing sporoplasm enclosing one nucleus, and the outer episporoplasm without nucleus, both areas separated by two membranes. The wall of the developing spore was produced by deposition of dense material in nodes on the outer of those two membranes (Fig. 6A) followed by growing of the nodes until they fused (Fig. 6B). The spore wall formed a flange on one of the poles and the portion of the wall becoming the operculum rested on the flange, with the exception of the hinge

area. The sporoplasm contained one nucleus, a spherule, mitochondria, lipid droplets and vesicles of variable size; in maturing spores, the sporoplasm showed abundant oval to elongated, membrane bound, electron-dense bodies, up to 850 x 280 nm, and without identifiable internal substructure typical of haplosporosomes (Figs. 6C-6E). Some maturing spores (Figs. 6C and 6D) showed a portion of the sporoplasm not covered by wall, which could be the result of opercular opening, thus suggesting possible spore excysment. The mature spore wall consisted of an inner electron-dense layer 40 nm thick, next a lighter layer 25 nm thick, a dense layer 50 nm thick and a denser outer layer with irregular thickness. The outer layer showed protrusions, with ca. 170 nm of diameter in section and up to 420 nm long, at variable distance among them, consisting of an electron-dense outer layer, ca. 30 nm thick, continuous with the spore wall outer layer and a light content; these protrusions formed variable angles with the spore wall; some were curved and others straight (Figs. 6C-6G). Additionally, longitudinal sections of filaments composed of spore wall material were observed close (Fig. 6C) or tangential (Fig. 6D) to spores; in a favorable section, two filaments appeared to project from a spore wall thickening or knob (Fig. 6G). The episporoplasm degraded as spore matured and the sporoplasm became reduced, detaching from the wall, thus leaving an empty space between sporoplasm and spore wall (Figs. 6F and 6G). Examination with SEM showed that spores were slightly ellipsoidal with a truncate pole corresponding to the operculum (Figs. 7A and 7B) and measuring 4.1 – 5.7 µm long (mean = 4.7; SE = 0.31; n = 10) and 3.5 – 4.2 µm wide (mean = 3.8; SE = 0.12; n = 10). The spore surface appeared profusely sculptured with multiple reliefs (Figs 7A-7E). Some spores had long tape-like filaments, either 4 (Fig. 7C) or 2 (Figs. 7D and 7E). The filaments were ca. 700 nm in diameter while the maximal length could not be estimated because the images did not include complete long filaments, but were longer than 20 µm. Favorable view allowed confirmation that two filaments arose from a common knob of the spore wall (Fig. 7D). Knobs on spores lacking obvious filaments probably corresponded to observed truncated filaments (Figs 7A and 7B).

3.4 Molecular analysis Amplification was obtained by PCR in 27 out of 30 analyzed individuals. A sequence with 1,409 base pairs of the SSU rDNA gene of the parasite was obtained from each PCR-positive P. nobilis by aligning the sequences of the different PCR amplicons produced with the 4 primer pairs used in the study (Table 2). Two individuals from Almería and Valencia in which the haplosporidan parasite was not detected by histology were positive by PCR reaction. Amplification was not obtained in the tissue samples from three healthy P. nobilis (from Alfacs Bay, Mar Menor and Calpe) or from crustacean symbionts, confirming the absence of the parasite. Two haplotypes of the

haplosporidan parasite were obtained; the difference between these two haplotypes corresponded to two specific positions of the DNA fragment (nucleotide positions 1206 and 1218), where each one presented and , respectively. The consensus sequence was deposited in GenBank under accession number LC338065. For phylogenetic analysis, the Akaike Information Criterion using JModeltest software, indicated the GTR+G as the best fit model of DNA sequence evolution. Due to the different length of the sequences available in the GenBank database, we built two different phylogenetic trees including the three methods (ML, NJ and BI): in the first we used large sequences of SSU rDNA (about 1400 bp) of Haplosporidia and other protozoa and in the other we used partial sequences of about 650 bp (from the nucleotide position 760 to the 1409 of our sequence) available for a higher number of species of the genus Haplosporidium (Figs. 8A and 8B). The evolutionary pattern was similar using the large sequences with the three described methods. The species in the order Haplosporida were included in a paraphyletic group, showing the most of them clustering in a main clade, clearly separated from the other protozoans, with the exception of Urosporidium and some of Haplosporidium species included in a different branch. The cladogram was supported by significant values of bootstrap. Within the main haplosporidan clade, various sub-clusters were observed. The first one was a monophyletic clade that includes all Minchinia and Bonamia species, which were significantly separated from each other in independent groups (Fig. 8A). A second lineage grouped the majority of Haplosporidium spp. (significant bootstrap value ML equal to 73%). H. nelsoni and Haplosporidium diporeiae as well as Haplosporidium patagon and Haplosporidium sp. from Syllis nipponica remained outside this subclade, forming two separate groups. The parasite infecting P. nobilis was clearly included in the Haplosporida group closely related to Haplosporidan parasite of P. vannamei, supported by significant bootstrap values (Fig. 8A). However, many differences between these more closely related species were identified, with a total of 190 variable sites (13.9%). These two taxa appeared to be a separate group from other Haplosporidium, Minchinia and Bonamia species. In the alternative phylogenetic reconstruction, considering the shorter sequences, the parasite infecting P. nobilis was situated in a single branch, even though most Haplosporidium spp. clustered in different subclades supported by significant bootstrap values (Fig. 8B). The pairwise Nei's genetic distance among species of the Haplosporida order varied from 0.019 to 0.375 (Table Supplementary information), with the highest values reported between Haplosporidium littoralis and H. louisiana, while in the comparisons of all Haplosporidium spp. with other genera excluding outgroup they varied between 0.188 and 0.691. The distance of the H. pinnae with other Haplosporidium species ranged from 0.137 to 0.297 with the lowest values

detected in the relation with Bonamia ex Crassostrea ariakensis and the highest with H. lousiana.

3.5 Taxonomic summary Phylum: Cercozoa Cavallier-Smith, 1998 Sub Phylum: Endomyxa Cavalier-Smith, 2002 Class: Ascetosporea Cavalier-Smith, 2002 Order: Haplosporida Caullery & Mesnil, 1899 Family: Haplosporidiidae Sprague, 1979 Genus: Haplosporidium Caullery & Mesnil, 1899 Haplosporidium pinnae sp. nov. Diagnosis: Spherical to elongated uninucleate and binucleate stages, approximately 3.5 µm in maximum length, occur throughout the connective tissue and haemolymph of the host. Sporulation stages occur in the epithelium of the digestive gland tubules of the host, including sporonts, sporocysts and spores, as well as pre-sporulation stages, uninucleate cells, binucleate and multinucleate plasmodia. Sporocysts approximately 30 µm in maximum length, with more than 50 spores. Slightly ellipsoidal spores, 3.6 – 5.7 µm in length and 2.7 – 4.5 µm in width, truncated in one pole with an orifice covered with a hinged operculum, consisting of a wall that encloses the sporoplasm with one nucleus; spores may have an undetermined number of long tape-like filaments (up to 4 were observed by SEM), about 700nm wide and longer than 20 µm, attached to knobs of the wall. Type host: Fan mussel Pinna nobilis Type location: Andratx Harbour, Balearic Islands, Spain (39° 32’ 22.8 N, 2° 22’ 24.11W, ETRS89) Type material: Hapantotype slide with a stained histological section of a fan mussel infected with all the stages deposited in the International Protozoan Type Slide Collection at the Smithsonian Institution, Washington, DC 20560 (accession no. xxxxxx). The paraffin block of the section and additional slides, paraffin blocks and Epon blocks are deposited at the collection of the Laboratori d’Investigacions Marines i Aqüicultura (LIMIA). Ribosomal DNA sequence: Deposited to GenBank under accession number: LC338065: Etymology: The specific epithet refers to the genus of the host: “of Pinna”

4. Discussion The present study includes for the first time the morphological and molecular characterization of the parasite very likely responsible for the mass mortality event (MME) of P. nobilis in SouthWestern Mediterranean Sea. Affected but still living individuals of P. nobilis exhibited a lack of

response to stimuli and slow closing of the valves (Vazquez-Luis et al. 2017). The presence of the parasite was identified by histological and/or by molecular methods in dead or moribund P. nobilis, but not in healthy individuals. On the contrary, Pinna rudis was not affected by abnormal mortality and the parasite was not detected in the collected samples. P. rudis is a of the same genus that cohabits in the same areas as the fan mussel (Vázquez-Luis et al. 2014, Nebot-Colomer et al. 2016), but seems not to be affected by the MME, since abnormal mortality of this species has not been detected

in

any

of

the

sampled

localities

nor

reported

by

citizen

science

(www.observadoresdelmar.es). Morphological (histological and ultrastructural) and molecular analyses of the haplosporidan parasite infecting the fan mussel showed that it is different from previously known haplosporidan species. The occurrence of every known haplosporidan stage (uni- and binucleate cells, multinucleate plasmodia, sporocysts and spores) in the same individual host is not common among haplosporidan species; sporal stages and the sporulation process have never been detected in B. ostreae, B. exitiosa (Engelsma et al., 2014) and Haplosporidium littoralis (Stentiford et al., 2013). Uni- and binucleated cells are not common (or have not been observed) in some species of the genus Haplosporidium, such as H. tapetis (Vilela, 1951; AV pers. observ.), H. nelsoni (Haskin et al. 1966; Couch et al., 1966), H. tumefacientis (Taylor, 1966), H. comatulae (La Haye et al., 1984), H. raabei (Molloy et al. 2012), H. edule (Azevedo et al., 2003) and H. patagon (Di Giorgio et al., 2014; Ituarte et al., 2014). Descriptions of some Haplosporidium spp., such as H. parisi (Ormières, 1980) and H. tuxtlensis (Vea and Siddall, 2011), were focused on sporulation stages, without reference to other stages. The occurrence of abundant uni- and binucleate parasite cells and the scarcity of multinucleate stages throughout the connective tissue and haemolymph sinuses of the hosts suggested that parasite proliferation to reach systemic infection involved karyokinesis in the uninucleate cells producing binucleate cells followed by cytokinesis resulting in two daughter uninucleate cells. Sporulation was exclusively observed in the digestive gland tubules of the fan mussels, thus migration of uni- and binucleate cells from the connective tissue towards digestive gland tubules can be deduced. The heavy accumulation of uni- and binucleate cells in the epithelium of digestive gland tubules near multinucleate plasmodia suggested that karyokinesis in uni- and binucleate cells without cytokinesis gave rise to the multinucleate plasmodia. Fragmentation of plasmodia giving rise to uninucleate cells should not be discounted, as it was observed in a haplosporidan parasite infecting the digestive gland tubules of the shrimp P. vannamei by Dykova et al. (1988). Furthermore, Desportes and Nashed (1983) suggested the formation of multinucleate plasmodia of Minchinia dentali through the fusion of smaller stages with

two or four paired nuclei. Similarly, fusion of uni- or binucleate cells could also occur in the fan mussel parasite. Most ultrastructural characters observed by TEM were consistent with the allocation of the parasite within haplosporidans; however, typical haplosporosomes were not found in the examined stages (multinucleate plasmodia, sporonts, sporocysts and spores) although uni- and binucleate cells were not found in the processed materials. Perkins (1971) used the term haplosporosomes referring to “membrane-bound, osmiophilic structures sub-divided into a cortex and medulla separated by an electron-lucent zone”. Additionally, Perkins (1979) established that both cortex and medulla show high electron-density. Haplosporosomes have been found in the four haplosporidan genera (Minchinia, Haplosporidium, Urosporidium and Bonamia) (Hine, 2009) as well as in parasites of the order Paramyxida (Lester and Hine, 2017), which are included in the class Ascetospora as the haplosporidans. The multinucleate plasmodia of the fan mussel parasite showed abundant vesicles with two concentric membranes (thus allowing distinction of the cortex and medulla), with size within the range stated by Perkins (1971, 1979) for haplosporosomes, although their “medulla” was mostly electron-lucent. Structures typically involved in the formation of haplosporosomes such as multivesicular bodies (Perkins, 1968) or Golgi apparatus (Hine, 1992) were not identified in the multinucleate plasmodia of H. pinnae¸ as observed for the haplosporidan parasite infecting the shrimp P. vannamei (Dykova et al. 1988). Nevertheless, the anastomosing endoplasmic reticulum (ER) cisternae observed in the multinucleate plasmodia of H. pinnae could be involved in the formation of the surrounding vesicles with two concentric membranes, because anastomosing ER cisternae are involved in the formation of haplosporosomes in various haplosporidans (Hine, 2009). Additionally, the image suggesting release of vesicles from a nuclear evagination evoked the involvement of the nuclear surface in the formation of haplosporosomes of B. exitiosa (Hine, 1992). The abundant oval to elongated, membrane bound, electron-dense bodies observed in the spores of the fan mussel parasite resembled the rod-like haplosporosomes reported by Perkins (1971) but the cortex and medulla could not be clearly distinguished in them. Similar membrane-bound, electron dense bodies without typical haplosporosome substructure were described in the sporoplasms of H. lousiana (Perkins 1975), H. comatulae (La Haye et al., 1984) and Minchinia teredinis (Hillman et al., 1990) but the involvement in producing haplosporosomes suggested for those of H. louisiana and M. teredinis, based on the observation of smaller denser spheres within the electron dense bodies, could not be deduced from those of H. pinnae, because the latter did not enclosed smaller denser spheres. Spore ornamentation has been considered relevant in haplosporidan taxonomy (Burreson and Reece, 2006; Azevedo and Hine, 2017); unfortunately, the fixation of the spores examined with

SEM in our study was not optimal (spores were isolated from fan mussel tissues preserved in ethanol) and some doubts remain about description of the external spore morphology. The multiple spore wall protrusions observed with TEM likely corresponded to the profuse relief of the spore surface observed with SEM; these protrusions were longer than the “léger replis” (light folds) observed in the spore wall of H. parisi (Ormières, 1980). The protrusions were also different from the folds and the slender longer projections of the spore wall described in H. edule (Azevedo et al., 2003) because the spore ornaments of the latter species are electron-dense, whereas the wall protrusions of H. pinnae had an electron lucent interior, which conferred a hollow appearance. Both TEM and SEM examination of the fan mussel parasite showed that pairs of filaments projected from wall knobs; the number of filaments attached to the spore varied from none to two or four; whether the number of filaments is actually variable or the observed variability was due to loss of filaments during processing remains unresolved. Hypothetically, filaments could have been accidentally detached from knobs observed with SEM in spores without filaments. The occurrence of two long tape-like filaments arising from a knob of the spore wall was also reported from H. lusitanicum (Azevedo, 1984) and a pair of filaments projecting from a knob of the spore wall were observed in spores of H. parisi (Ormières, 1980), H. hinei (Bearham et al., 2008) and H. diporeiae (Winters et al., 2014). The TEM images showing a portion of the sporoplasm not covered by the spore wall could correspond to the orifice area of spores with an open operculum, although a wall flange at the orifice border was not distinguished. This could suggest the possibility of spore excystment within the host, as described for H. lusitanicum (Azevedo et al., 1985) and Minchinia tapetis (Azevedo and Corral, 1989); however, free amoebulae with typical sporoplasm organelles (spherule, electron dense bodies) that would confirm excystment (Azevedo and Corral, 1989) were not observed in our study. Further ultrastructural exploration is required for better characterization of each H. pinnae stage, parasite proliferation and sporulation process, thus answering some of the questions raised in this study. The infection of the epithelium of host digestive gland tubules is rather exceptional in haplosporidans; most haplosporidan species proliferate throughout host hemal spaces and connective tissue, where sporulation usually occurs. Heavy infection of the fan mussel digestive gland tubules should deeply interfere with food absorption, likely leading to host starvation, thus causing severe general dysfunction with fatal outcome. Both H. nelsoni and H. tuxtlensis sporulate in the digestive gland tubules of their hosts, the eastern oyster Crassostrea virginica and the striped false limpet Siphonaria pectinata, respectively (Couch et al., 1966; Vea and Siddall, 2011). Additionally, heavy infection of the epithelium of digestive gland tubules of the shrimp P.

vannamei with Haplosporidium sp., causing disruption of host epithelial cells, has been reported from Caribbean countries and Indonesia (Dykova et al., 1988; Nunan et al., 2007; Utari et al., 2012). Similar to H. pinnae, H. nelsoni and the haplosporidan parasite of P. vanamei have been responsible for mass mortalities of their hosts, eastern oysters (Ford and Tripp, 1996) and penaeid shrimps (Utari et al., 2012). The low prevalence of H. tuxtlensis could explain that no abnormal mortality of false-limpets was mentioned when this parasite was described (Vea and Siddall, 2011). The infection of fan mussel with H. pinnae was associated with heavy hemocytic reaction even without sporulation. However, Darriba (2017) reported light or absent hemocytic reaction in the three infected fan mussels included in her study. Sporulation of H. nelsoni in adult oysters is rare but is frequent in spat (Barber et al., 1991; Burreson 1994) but mortality of infected adult oysters is high despite lack of sporulation; the heavy inflammatory reaction evoked by this parasite in adult oysters (Farley, 1968) may contribute to host death. Likewise, infection with H. pinnae without sporulation could be lethal for P. nobilis. In fact, sporulation was not detected in the fan mussels collected from some of the areas affected by the MME. The spore with hinged operculum and filaments attached to the spore wall would support allocation of the fan mussel parasite into genus Haplosporidium although those morphological characters also apply for B. perspora (Carnegie et al., 2006). Another similarity of the fan mussel parasite with B. perspora was the occasional occurrence of uni- and binucleate cells within hemocytes; whether parasite cells inside fan mussel hemocytes were viable or not could not be assessed. Both B. perspora and the fan mussel parasite produced all the known haplosporidan stages in their respective hosts although the sporulation process took place in different tissues, connective tissue and epithelium of digestive gland tubules, respectively. The occurrence of uni- and binucleate stages of the fan mussel parasite would suggest possible direct transmission from infected to healthy fan mussels, as occurs for B. ostreae and B. exitiosa (Hine, 1996, Culloty and Mulcahy, 2007; Audemard et al., 2014). Additionally, the spores of the fan mussel parasite (a dormant, resistant stage) could allow long persistence in the environment and the hypothetical involvement of an intermediate host as suggested for H. nelsoni and H. costale (Andrews, 1984; Haskin and Andrews, 1988; Powell et al., 1999). The data provided by histological analysis was supported by molecular data; the presence of the parasite was detected by molecular analysis in the analyzed samples of P. nobilis dead or dying. The phylogenetic analyses indicated that this parasite should be included within a large clade consisting of species of the order Haplosporida, in which it appeared to be distinct from the Bonamia/Minchinia clade and from the clade containing most of the Haplosporidium species but grouped with the haplosporidan parasite of P. vannamei.

The genetic distance of H. pinnae was variable with regard to other species of the order Haplosporida. Similar or higher values of divergence compared to those detected between Bonamia and Minchinia species and between distinct species of Haplosporidium, indicated the high level of differentiation of the fan mussel parasite. Furthermore, the pairwise genetic distance was similar when comparing this parasite with Bonamia spp. (from 0.137 to 0.148), with the haplosporidan parasite of P. vannamei (0.153), with Minchinia spp. (0.145-0.159) as well as with H. diporeiae (0.159), and with Haplosporidium sp. ex Saccostrea glomerata (0.163). The lower values of genetic distance detected between the fan mussel parasite and Bonamia spp. was not entirely consistent with reconstruction trees, probably due to the presence of indels that were not completely considered in the Neighbor-Joining distance method. Previous phylogenetic analysis of haplosporidans have shown the paraphyly of the genus Haplosporidium as well as the monophyly of the genera Bonamia and Minchinia (Reece et al. 2004; Burreson & Reece 2006). The relationships among some lineages of Haplosporidium are difficult to resolve and a taxonomic review involving differentiation of new genera would be required (Hine, 2009; Arzul & Carnegie, 2015; Azevedo and Hine, 2017). The haplosporidan parasite of P. vannamei that was grouped with the fan mussel parasite in the phylogenetic analysis has not been described and named as a new species because of the lack of detection of a sporal stage (Dykova et al., 1988; Nunan et al., 2007; Utari et al., 2012). The SSU rDNA sequences of the P. vannamei parasites from Belize and Indonesia allowed assignment to the order Haplosporida, in same cluster as Minchinia spp. and Bonamia spp., but in a different branch, and in a different cluster from Haplosporidium spp. (Nunan et al., 2007; Arzul and Carnegie, 2015, Azevedo and Hine, 2017). Although we detected substantial nucleotide variability between the fan mussel parasite and that of P. vannamei, which strongly support designation as different species, our results suggest that these parasites could represent a new distinct genus within the order Haplosporida. However, considering that molecular analysis did not place the fan mussel parasite within the genera Bonamia or Minchinia, and assuming the current paraphyletic condition of the genus Haplosporidium, we opt for placing the fan mussel parasite in the genus Haplosporidium, as the new species H. pinnae. This species could be assigned to another genus in the future if taxonomic and phylogenetic thorough review are accomplished. The difference in clustering power between the two phylogenetic reconstructions built in this study was due to less genetic information when using the shorter sequences. Nevertheless, with this second phylogenetic approach, H. pinnae did not group with any other haplosporidan species. The scarce variability detected in the SSU rDNA sequence of H. pinnae could suggest a recent arrival of this parasite in the Spanish Mediterranean coast, which would be consistent with the

unprecedented recent MME. This parasite seems to be host specific because it was not detected in P. rudis, which occurs in the same areas and habitat as the co-generic species P. nobilis affected by mass mortality. It is unknown how the parasite is transported, but, according to the spatial and temporal mortality data of P. nobilis observed along the Southern coasts of Spain, the outbreak spread following the direction of the summer marine currents (Fernandez et al, 2005). Now identified, a thorough study of epizootic dynamics and the complete life cycle of H. pinnae is needed to find ways to mitigate disease and limit spread of the pathogen, and to reverse the “critically endangered” status of P. nobilis in the Spanish Mediterranean coast.

Acknowledgements GC would like also to thank INIA for the INIA/CCAA research contract program. MV is supported by a postdoctoral contract co-funded by the Regional Government of the Balearic Islands and the European Social Fund 2014– 2020. EA was also supported by a Personal Técnico de Apoyo contract (PTA2015-10829-I) funded by the Spanish Ministry of Economy and Competitiveness. The authors thank Elisabet Nebot, Francisca Giménez, Andrés Izquierdo, Juan Barja, Fernando Garfella, Guillermo Follana, Quique Massutí, Agustín Barrajón, Diego Moreno, Santiago Jiménez, José Luis Crespo, Manu Guimerans for providing samples. A special thank also to Subdirección General para la Protección del Mar (MAPAMA) for financial support to sample delivery. Sample processing and examination with transmission electron microscopy were performed in the Servicio de Microscopía Electrónica of CACTI (University of Vigo) with technical assistance by Jesús Méndez and Inés Pazos.

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FIGURE LEGEND Figure 1. Sampling sites of Pinnae nobilis specimens Figure 2. Histological sections through the digestive gland of fan mussels Pinna nobilis infected with Haplsoporidium pinnae. Bars: 5 µm. A: Detail of the connective tissue showing an uninucleate cell of the parasite (arrow). MHE staining. B: Binucleate cell of the parasite (arrow) in the connective tissue. HHE staining. C: Parasite cells (arrows) within host hemocytes in the connective tissue. HHE staining. D: Plasmodial stages (arrows) in the host stomach epithelium. HHE staining. Figure 3. Histological sections through the digestive gland of fan mussels Pinna nobilis infected with Haplsoporidium pinnae showing sporulation stages of the parasite in the epithelium of digestive gland tubules of the host. A: Accumulation (star) of uni- and binucleate cells of the parasite in the epithelium of a digestive gland tubule; the arrow points a plasmodium with 5 nuclei; arrowheads point large vacuoles with amorphous eosinophilic material. Bar: 10 µm; HHE staining B: Detail of the epithelium of a digestive gland tubule showing a deeply stained multinucleate plasmodium (double arrow) in which some cytoplasmic compartmentalisation is distinguished; the arrow points a smaller, less deeply stained plasmodium with 6 nuclei; a large vacuole with amorphous eosinophilic material (arrowhead) is shown. Bar: 5 µm; HHE staining. C: Detail of the epithelium of a digestive gland tubule showing a sporocyst enclosing numerous uninucleate sporoblasts; tightly packed uni- and binucleate parasite cells (star) also occur. Bar: 5 µm; HHE staining. D. Digestive gland tubules in which the epithelium is occupied by parasite sporocysts (arrow) and deeply stained plasmodia with cytoplasm compartmentalization is distinguished (double arrow); note brown cells in the connective tissue (arrowheads). Bar: 20 µm; MHE staining. E. Digestive gland tubules showing the epithelium occupied by sporocyst enclosing sporoblasts (blue colour, arrow) and sporocysts enclosing more or less mature spores (yellow-orange colour, double arrow). Bar: 20 µm; MH-VOF staining. Figure 4. Histological sections through the digestive gland of fan mussels Pinna nobilis, one individual infected (A) with Haplsoporidium pinnae and another non-infected (B and C). HHE staining. A: Low magnification micrograph of the digestive gland area of an infected individual showing heavy hemocytic infiltration of the connective tissue in the digestive gland. Bar: 200 µm. B: Low magnification of the digestive gland of a non-infected individual in which the connective tissue is not abnormally infiltrated by hemocytes. Bar: 100 µm. C: Higher magnification of the digestive gland of a non-infected individual showing a digestive gland tubule in absorptive phase, with reduced lumen and high epithelium with numerous cytoplasmic brownish granules. Bar: 20 µm.

Figure 5. Transmission electron micrographs of Haplosporidium pinnae. A: Multinucleate plasmodium showing two paired nuclei (N), areas of accumulation of vesicles with two concentric membranes (double arrows), lipid droplets (L) and mitochondria (m). Bar: 2 µm. The inset in the lower right corner corresponds to enlargement of the framed area in the mid zone of the plasmodium, showing an evagination of the envelope of one of the paired nuclei from which the

outer membrane appears inflated, suggesting the release of vesicles from the nucleus. Bar: 200 nm. B: Three vesicles with two concentric membranes occurring in the cytoplasm of a plasmodium; the marginal area comprised between the outer and the inner membrane is electron-dense while the inner core is lighter with sparse granulation. Bar: 500 nm. C: Detail of a plasmodial area with accumulation of vesicles with two concentric membranes and tubular cisternae of smooth endoplasmic reticulum, with dense content. Bar: 200 nm. Figure 6. Transmission electron micrographs of the sporulation process of Haplosporidium pinnae. A: Area of a sporont with a sporoblast showing two compartments, the episporplasm (asterisk) and the sporoplasm (double asterisk), separated by two membranes; the developing spore wall appears as nodes of dense material deposited on the outer of these two membranes, in the episporoplasm side. Arrows point out the operculum border confronting a flange of the spore wall in two sporoblasts. Mitochondria (m), one spherule (sp) and one nucleus (N) are shown in the sporoplasm. Bar: 1 µm. B: Sporoblast in which the nodes with dense material have fused producing an almost continuous envelope; wall flanges (arrow) confront the borders of the operculum (double arrow). Lipid droplets (L) appear in the sporoplasm. Bar: 1 µm. C: Maturing spore with the sporoplasm partially surrounded by a wall; multiple wall protrusions (black arrows) are shown and a fragment of a filament (F) appears close to the maturing spore; the sporoplasm shows abundant oval to elongated electron-dense bodies (white arrows) and one nucleus (N). Bar: 1 µm. D: Maturing spore with the wall partially surrounding the sporoplasm showing various wall protrusions (black arrows) as well as a filament (F), composed of wall material, tangential to the wall; oval to elongated electron-dense bodies (white arrows) are abundant in the sporoplasm. Bar: 1 µm. E: Detail of a maturing spore showing its wall with protrusions (black arrows) and electron-dense bodies (white arrows) in the sporoplasm. Bar: 500 nm. F: Mature spore showing operculum, sporoplasm retraction and various wall protrusions (arrows). Bar: 1 µm. G: Detail of an area of a mature wall showing a knob of its wall from which two filaments (F) arise. Bar: 500 nm Figure 7. Scanning electron micrographs of spores of Haplosporidium pinnae. A: Spore showing profuse relieves on the surface, the operculum (white arrow) and knobs (arrowhead) that could correspond to truncated filaments; the black arrow points out a ridge. Bar 1 µm. B. Another spore showing a ridge (black arrow), the operculum (white arrow) and knobs (arrowheads) that could correspond to truncate filaments. Bar 1 µm. C. Spore with four tape-like filaments (arrows). Bar 2 µm. D. Spore with two tape-like filaments (arrows) projecting from a common knob of the spore wall. Bar 2 µm. E: Spore with two long tape-like filaments (arrows). Bar 5 µm.

Figure 8. Phylogenetic relationships based on the SSUrDNA sequence. A: Among species of the order Haplosporida (highlighted in grey) and other species of protozoans using the sequence length of about 1400 bp. B: Among species of the order Haplosporida using the sequence length of about 650 bp. Maximum likelihood, Neighbor-Joining and Bayesian Inference bootstrap values higher than 50% are indicated below nodes, respectively.

Table 1. Sampling data of the Pinna nobilis specimens analysed in the study and results of detection of Haplosporidium pinnae by histologic and molecular analysis. NP: not processed; ND: not determined. PO: Posidonia oceanica, PO and B: Posidonia oceanica and boulders, M and B: mud and boulders, CN and M: Cymodocea nodosa and mud; and D: detritic. Date

Pinna nobilis 03/11/2016 03/11/2016 03/11/2016 03/11/2016 17/10/2016 17/10/2016 17/10/2016 24/11/2016 24/11/2016 24/11/2016 24/11/2016 24/11/2016 24/11/2016 24/11/2016 24/11/2016 24/11/2016 24/11/2016 24/11/2016 24/11/2016 24/11/2016 24/11/2016 27/12/2016 27/12/2016 27/12/2016 03/03/2017 03/03/2017 03/03/2017 03/03/2017 03/03/2017 03/03/2017 10/03/2017 18/05/2017 03/05/2017 16/05/2017 03/05/2017 03/05/2017 05/04/2017 04/07/2017 04/07/2017 26/07/2017 19/07/2017 26/06/2017 27/06/2017 02/11/2017 02/11/2017 02/11/2017 02/11/2017 Pinna rudis 20/05/2017 31/01/2018

Location

Histological Molecular Length (cm) evaluation evaluation

Total weight (g)

Total soft tissue weight (g)

Depth (m)

Habitat

Calpe (Alicante)* Calpe (Alicante)* Calpe (Alicante)* Calpe (Alicante)* El Calón (Almeria) El Calón (Almeria) El Calón (Almeria) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Andratx (Mallorca) Cabrera (Mallorca) Cabrera (Mallorca) Marina Real (Valencia) Marina Real (Valencia) Marina Real (Valencia) Marina Real (Valencia) Marina Real (Valencia) Marina Real (Valencia) Marina Real (Valencia) Mar Menor (Murcia) Alfacs (Tarragona) Sa Farola (Menorca) Son Saura (Menorca) Tossa de Mar (Girona) Tossa de Mar (Girona) Tossa de Mar (Girona) Tossa de Mar (Girona)

+ + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + +

+ + + + + + NP NP NP NP NP NP NP NP NP NP NP NP NP NP + + + + + + + + + + + + + + + + + + + + + + + +

50.2 41.2 41.5 40.1 69.3 70.2 71.3 44.6 39.8 46.5 46.5 43 44.3 43.5 39.5 35.5 36.6 48.0 28.1 33.5 46.1 ND ND 26.3 51.3 38.5 44.0 40.4 30.5 41.3 36.5 65 ND ND 48.2 50.8 56 ND ND 22.2 46.7 34.5 35.7 46.3 41.5 38.5 49.5

598.5 414.9 438.5 388.5 ND ND ND 541.1 404.6 745.7 780.2 521.5 594.7 687.1 495.7 325.9 396.5 681.7 376.2 275.4 616.0 ND ND 212.2 1280.4 610.1 580.1 450.0 210.2 490.1 301.3 ND ND ND 1290.1 880.4 980.3 ND ND 82.1 1159.5 ND 529.6 716.9 600.5 325.5 1036.1

84.5 56.5 52.9 41.5 ND ND ND 88.8 53.4 122.8 136.4 72.4 101.8 152.8 70.7 41.5 56.5 88.4 69.6 48.9 131.2 ND ND ND 246.5 74.8 101.5 54.5 38.7 79.2 63.5 111 ND ND ND ND ND ND ND 12.7 206.95 ND 63.5 108.0 87.2 48.2 124.9

6-8 6-8 6-8 6-8 20 20 20 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 5-8 8 30 10-15 10-15 10-15 10-15 10-15 10-15 10-15 1 0.3 5 8 6-12 6-12 6-12 6-12

PO PO PO PO PO PO PO PO and B PO and B PO and B PO and B PO and B PO and B PO and B PO and B PO and B PO and B PO and B PO and B PO and B PO and B PO PO PO PO PO PO PO PO PO PO PO and B M and B M and B M and B M and B M and B M and B M and B CN and M CN and M PO PO PO PO PO PO

Cabrera (Mallorca) Cabrera (Mallorca)

-

-

43.4 34.5

1237 520

188 130

36 28

D D

* These specimens had been used in the study by Darriba et al. (2017) and were involved in this study for the molecular characterisation of Haplosporidium pinnae.

Table 2 Primers used in the study for PCR amplification of the SSU rDNA sequence of Haplosporidium pinnae Primer name Hap-F1 Hap-R2 HPN-F1 HPN-F3 HPN-R3 16SB

Primer pair Hap-F1-Hap-R2 HPNF1-HPNR3 HPNF3-HPNR3 HPNF3-16Sb

Sequence 5’-3’ GTTCTTTCWTGATTCTATGMA GATGAAYAATTGCAATCAYCT AGCTTGACGGTAGGATATGGG CATTAGCATGGAATAATAAAACACGAC GCGACGGCTATTTAGATGGCTGA GATCCTTCCGCAGGTTCACCTAC

Tm 49ºC 52ºC 55ºC 57ºC

Reference Renault et al., 2000

This work Medlin et al. 1988

Amplicon size (bp) 330 1100 600 1000

Highlights - A new species, Haplosporidium pinnae, in the order Haplosporida, is described - Histology, TEM, SEM and rDNA sequence confirmed distinction as a new species - H. pinnae is the most plausible cause of mass mortalities of the bivalve Pinna nobilis

Graphical abstract