Haplosporidium raabei n. sp. (Haplosporidia): a parasite of zebra mussels, Dreissena polymorpha (Pallas, 1771)

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Haplosporidium raabei n. sp. (Haplosporidia): a parasite of zebra mussels, Dreissena polymorpha (Pallas, 1771) D. P. MOLLOY 1 *, L. GIAMBÉRINI 2 , N. A. STOKES 3 , E. M. BURRESON 3 and M. A. OVCHARENKO 4,5 1

Division of Research and Collections, New York State Museum, Albany, NY 12230, USA Université Paul Verlaine – Metz, Laboratoire des Interactions, Ecotoxicologie, Biodiversité, Ecosystèmes (LIEBE), CNRS UMR 7146, Campus Bridoux, Rue du Général Delestraint, F-57070 Metz, France 3 Virginia Institute of Marine Science, College of William & Mary, Gloucester Point, P.O. Box 1346, Virginia 23062, USA 4 Institute of Parasitology, Polish Academy of Sciences, Twarda 51/55, 00-818 Warsaw, Poland 5 Pomeranian Akademy, Arcishewski str. 22b, 76-200, Słupsk, Poland 2

(Received 15 March 2011; revised 28 July, 10 September and 13 October 2011; accepted 27 October 2011; first published online 5 January 2012) SUMMARY

Extensive connective tissue lysis is a common outcome of haplosporidian infection. Although such infections in marine invertebrates are well documented, they are relatively rarely observed in freshwater invertebrates. Herein, we report a field study using a comprehensive series of methodologies (histology, dissection, electron microscopy, gene sequence analysis, and molecular phylogenetics) to investigate the morphology, taxonomy, systematics, geographical distribution, pathogenicity, and seasonal and annual prevalence of a haplosporidian observed in zebra mussels, Dreissena polymorpha. Based on its genetic sequence, morphology, and host, we describe Haplosporidium raabei n. sp. from D. polymorpha – the first haplosporidian species from a freshwater bivalve. Haplosporidium raabei is rare as we observed it in histological sections in only 0·7% of the zebra mussels collected from 43 water bodies across 11 European countries and in none that were collected from 10 water bodies in the United States. In contrast to its low prevalences, disease intensities were quite high with 79·5% of infections advanced to sporogenesis. Key words: Haplosporidium raabei n. sp., Haplosporidia, Dreissena polymorpha, phylogeny, small subunit ribosomal DNA.

INTRODUCTION

Species in the phylum Haplosporidia are coelozoic or histozoic, spore-forming, endoparasitic protists, and although a relatively small group, they are well documented as pathogens of marine invertebrates, especially of commercially important bivalves (Burreson and Ford, 2004). One of the most investigated species, Haplosporidium nelsoni, the causative agent of MSX disease, has contributed to major declines in oyster populations along the eastern coast of the United States for decades (Ford and Tripp, 1996; Burreson and Ford, 2004). Observations of haplosporidian infection in freshwater invertebrates, however, are relatively rare (Burreson and Ford, 2004), and the first report of a haplosporidian in zebra mussels, Dreissena polymorpha (Pallas 1771) (Dreissenidae), was by Bowmer and van der Meer (1991) in The Netherlands. As early as 1988, they had observed a putatively lethal infection in the blood system and all key organs of zebra mussels. The study reported herein is an outgrowth of their discovery * Corresponding author: Present address: Department of Biological Sciences, State University of New York, 1400 Washington Avenue, Albany, NY 12222, USA. Tel: + 1 518 677 8245. Fax: + 1 518 677 5236. E-mail: [email protected]

and represents a 17-year field study using a comprehensive series of methodologies (histology, dissection, electron microscopy, gene sequence analysis, and molecular phylogenetics) to investigate the morphology, taxonomy, systematics, geographical distribution, pathogenicity, and seasonal and annual prevalence of this disease agent. Based on its genetic sequence, morphology, and host, we describe Haplosporidium raabei n. sp. (phylum Haplosporidia) from D. polymorpha – the first haplosporidian species from a freshwater bivalve.

MATERIALS AND METHODS

Sampling for histological examination During this 17-year field study (1992–2009), 732 zebra mussels from 10 water bodies in the eastern United States and 5514 zebra mussels from 43 water bodies within 11 European countries were histologically examined (mussel length ca. 10–30 mm). For analysis by light microscopy (4 X1000), tissue samples were fixed in either Bouin’s (followed by a water rinse) or 10% neutral buffered formalin, dehydrated in a graded series of alcohols and toluene, embedded in paraffin, cut into 5 μm thick serial sections, and stained with either toluidine blue or haematoxylin and eosin. Water bodies sampled (number of mussels

Parasitology (2012), 139, 463–477. © Cambridge University Press 2012 doi:10.1017/S0031182011002101

D. P. Molloy and others

examined) in the United States were: Hudson River, New York (80), Huron River, Michigan (22), Lake Champlain, New York (25), Lake Erie, Ohio (25), Southeastern Lake Michigan, Indiana (24), Southwestern Lake Michigan, Illinois (25), Mississippi River, Mississippi (25), Mohawk River, New York (456), Niagara River, New York (25), and Raisin River, Michigan (25). Sampling dates are provided in a Supplementary File (in the online version only). Water bodies sampled (no. mussels examined) in Europe were: Belarus, Svisloch River (391), Lake Lepelskoye (25), Lake Chareya (26), Lake Lukomskoe (23), Dnieper-Bug Canal (162); Czech Republic, Lake Zelivka (30); Denmark, Lake Esrum So (26), Lake Jels Nederso at Jutland (30); France, Lake Salagou (112), Moselle River (1158), Meuse River (1496), Mirgenbach Lake (152), Vilaine River (97), Lake Madine (71); Germany, Lake Gartow (49), Unterhavel River at Stoßensee (54), Baggersee Reeserward (55), Bodensee (55), Weser River at Bremen (55); Italy, Garda Lake (30); The Netherlands, Lake Ijsselmeer (167), Lake Maarsseveen (30), Ijssel River near Kampen (38), Noordzeekanaal (89), western distributary of the Rhine and Meuse including lakes Haringvliet (29), Volkerak (149), and Lake Hollands Diep (30); Poland, Lake Insko (30); Russia, Moscow River (40), Moscow Canal Sluice N8 (20), Ivankovskoye Reservoir (48), Uchinskoye Reservoir (20), Klazminskoye Reservoir (31), Chimkinskoye Reservoir (40), Lake Udomlya (19), Volkhof River at Novgorod (43), Gulf of Finland near St. Petersburg (22), Lake Valdaiskoiye (18), Velikaya River near Pskov (20), unnamed pond near Pushkin (158); Switzerland, Lake Geneva (30); and Ukraine, Dniester River at Beliaevka (10) and Dnieper River at Kiev (336). Sampling dates are provided in a Supplementary File (online version only).

Sampling of fresh squashed preparations In addition to the above-mentioned extensive sampling performed exclusively for the purpose of histological examination, other samples were taken solely for dissection of live mussels. To determine if infection could be detected in tissue smears, 1273 live D. polymorpha from The Netherlands were dissected in August through November 2000. Water bodies sampled (number of mussels dissected) were: River Ijssel near Kampen (130), Lake Ijsselmeer (310), and the western distributary of the Rhine and Meuse including lakes Haringvliet (138), Volkerak (541), and Hollands Diep (154). Portions of the gills and visceral mass were dissected out of the mussels, placed on a glass slide, and a coverglass applied with pressure. These fresh squashed preparations were then examined for infection by light microscopy (4X1000).

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Transmission electron microscopy For TEM, tissues from 6 heavily infected mussels were fixed in 2% glutaraldehyde, rinsed in buffer solution (0·05 M), and were post-fixed with 1% osmium tetroxide (Sigma) buffered with sodium cacodylate. After dehydration through graded alcohols, tissues were embedded in Epon-Araldite (Sigma). Ultrathin sections (60–80 nm), cut with a diamond knife on a LKB Ultratom V ultramicrotome, were placed on copper grids and stained with uranyl acetate and lead citrate. Sections were examined with a Jeol CX100 (80 kV) transmission electron microscope. Scanning electron microscopy For SEM performed at the University of Metz (Figs 4C, 6A), tissues from 2 heavily infected mussels were fixed as for TEM, followed by dehydration through graded alcohols and propylene oxide. Then they were placed onto a carbon stub, air dried, and coated with gold-palladium film. Tissues were examined for signs of haplosporidian infection with a Leica S440 scanning electron microscope at 15 kV. SEM performed at the Virginia Institute of Marine Science (Fig. 6C) followed the protocol of Ford et al. (2009). DNA extraction and PCR Tissue samples of 8 infected mussels collected from the Meuse River, Commercy, France in June 2001 (n = 6) and September 2002 (n = 2) and of 1 infected mussel from the Moselle River, Cattenom, France in April 2002 were lysed and DNA extracted using the QIAamp DNA mini kit (Qiagen). DNA was quantified using a GeneQuant pro spectrophotometer (Amersham Biosciences). Initially, genomic DNA was PCR amplified using 16S-A + HAP-R1 and HAP-F1 + 16S-B (Table 1). Primers 16S-A and 16S-B are general eukaryotic small subunit ribosomal DNA (SSU rDNA) primers with polylinker bases removed (Medlin et al. 1988). The HAP primers were designed to specifically amplify most haplosporidians (Renault et al. 2000). Reactions (25 μl) contained buffer (10 mM Tris-HCl, pH 8·3, 50 mM KCl, 1·5 mM MgCl2, 0·001% gelatin), 0·2 mM each deoxyribonucleotide, 0·4 μg bovine serum albumin, 12·5 pmol each primer, 0·6 U AmpliTaq polymerase (Applied Biosystems), and 200–250 ng DNA. All PCR were done in a TGradient (Biometra) or DNA Engine (Bio-Rad) thermal cycler. All PCR had 4 min initial denaturation at 94 °C, cycling 35 times at temperatures and times indicated for each primer pair in Table 2, and final extension for 5 min. Expected, and in one case actual, size PCR product from each primer pair is shown in Table 2. 16S-A + HAP-R1 PCR generated a product of approximately 400 bp rather than the expected approximately 1400 bp.

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Table 1. PCR primers and their binding sites within the Haplosporidium raabei SSU rDNA (Primer specificity notes the organisms to which these SSU rDNA primers bind. The expected binding site for HAP-R1 is shown in brackets; when paired with 16S-A its actual binding site was as noted. HMB refers to Haplosporidium spp., Minchinia spp., and Bonamia spp. of the Haplosporidia.) Primer name

Sequence (5′ – 3′)

Binding site

Primer specificity

Reference

16S-A

AACCTGGTTGATCCTGCCAGT

5′ end

eukaryotes

H135f ZMH5 seqZMH480 HAP750f ssu980 HAP-F1

ACTCTAGGGCTAATACGTGA CGTGCATATTAGACTAAAACCAATGTC AGTACAACGCAAAAGCCTTAAC CGCCTGAATGCATTAGCA CGAAGACGATCAGATACCGTCGTA GTTCTTTCWTGATTCTATGMA

136–155 172–198 469–490 756–773 978–1001 1224–1244

Haplosporidium spp. H. raabei internal seq primer most HMB eukaryotes most HMB

HAP-R1

CTCAWKCTTCCATCTGCTG

most HMB

BON-1110r seqZMH1400 HAP1590r ZMH3 16S-B

CCTTTAAGTTTCACTCTTGCGAG GGCCCCAAACTTCCCACTGCTG GACGTAATCAATGCAAGTTG CCACTCCATTCACCTGATTATTCG GATCCTTCCGCAGGTTCACCTAC

400–382 [1407–1389] 1121–1099 1410–1389 1605–1586 1667–1644 3′ end

Medlin et al. 1988 This study This study This study This study This study Renault et al. 2000 Renault et al. 2000 White, 2008 This study This study This study Medlin et al. 1988

several HMB internal seq primer most HMB H. raabei, H. edule eukaryotes

Table 2. Primer pairs and PCR cycling parameters used for amplification of Haplosporidium raabei SSU rDNA (All reactions had initial denaturation of 4 min at 94 °C, final extension of 5 min at cycling extension temperature, and cycling 35 times. The expected product for 16S-A + HAP-R1 was much larger than the actual size product obtained, shown in brackets. Cycling parameters marked with an asterisk were preceded with 10 cycles of the same conditions but annealing starting at 63 °C and decreasing 1 °C/cycle for touchdown PCR. Primer pairings are listed in the order they were tested on H. raabei.)

Primer pair

PCR cycling

Expected product (approx bp)

16S-A + HAP-R1 HAP-F1 + 16S-B ssu980 + HAP-R1 ZMH5 + ZMH3 16S-A + BON-1110r MBH1088f + 16S-B H135f + BON-1110r HAP750f + HAP1590r

94 °C(30 sec) – 54 °C(30 sec) – 72 °C(1·75 min) 94 °C(30 sec) – 46 °C(30 sec) – 70 °C(1·5 min) 94 °C(30 sec) – 54 °C(30 sec) – 72 °C(1·5 min) 94 °C(30 sec) – 65 °C(30 sec) – 72 °C(2·0 min) 94 °C(30 sec) – 53 °C(30 sec) – 72 °C(2·0 min) 94 °C(30 sec) – 53 °C(30 sec) – 72 °C(1·5 min) 94 °C(30 sec) – 53 °C(30 sec) – 72 °C(2·0 min)* 94 °C(30 sec) – 53 °C(30 sec) – 72 °C(2·0 min)*

1400 [400] 550 430 1300 1130 680 983 849

PCR of a portion of the 5′ end was then successful after pairing HAP-R1 with a different general eukaryotic SSU rDNA primer, ssu980. Sequence information from these amplification products confirmed that we had novel haplosporidian DNA and allowed design of specific primers, ZMH5 and ZMH3 (Table 1). PCR of samples extracted later used the other 4 primer pairs listed in Table 2. All PCR reactions were electrophoresed on agarose gels, stained with ethidium bromide, and visualized under UV light.

Sequencing For cloning, triplicate PCR products were pooled, purified with the QIAquick kit (Qiagen), and ligated

into plasmid vector pCR2.1 or pCR4-TOPO using the TA Cloning kit or TOPO TA Cloning kit (Invitrogen) following the manufacturer’s protocol. Initially, clones were sequenced by simultaneous bidirectional cycle-sequencing using M13 forward and reverse infrared-labelled primers (LI-COR) and the ThermoSequenase sequencing kit (Amersham) according to the manufacturer’s directions. After partial sequences were obtained, primers seqZMH480 and seqZMH1400 were designed (Table 1), synthesized with infrared-labels (LI-COR), and used to complete sequencing of the larger-sized clones. Sequencing reactions were electrophoresed and detected on a LI-COR model 4200 automated sequencer. Later in the project, clones were sequenced using unlabelled M13 forward and reverse primers (New England Biolabs) and the BigDye Terminator

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v3.1 Cycle Sequencing kit (Applied Biosystems). Sequencing reactions were electrophoresed on a 16capillary ABI 3130 Genetic Analyzer using Sequencing Analysis 5.2 software for base-calling (Applied Biosystems).

Sequence analyses and molecular phylogenetics Sequences generated from the cloned PCR products were aligned using ClustalW of the software package MacVector to generate consensus sequences for each mussel sample. When the consensus sequences from the 9 mussels were aligned together, there were 2 nearly identical sequences, designated clones A and B. These were checked by BLAST search (Altschul et al. 1997) against the GenBank sequence database and both were most similar to SSU rDNA of haplosporidians. Sequences of clones A and B were ClustalW-aligned with SSU rDNA sequences of the haplosporidians Haplosporidium edule (GenBank Accession DQ458793), H. pickfordi (AY452724), H. lusitanicum (AY449713), H. montforti (DQ219484), H. armoricanum (AY781176), H. costale (AF387122), H. nelsoni (U19538), H. louisiana (U47851), Minchinia teredinis (U20319, U20320), M. chitonis (AY449711), M. occulta (EF165631), M. mercenariae (FJ518816), M. tapetis (AY449710), Bonamia exitiosa (EU016528), Bonamia sp. ex Crassostrea ariakensis (AY542903), B. perspora (DQ356000), B. ostreae (AF262995), Urosporidium crescens (U47852), and unnamed haplosporidians from Cyrenoida floridana (AY449712), Penaeus vannamei (DQ653412), Syllis nipponica (DQ444238), Stictodora lari (AY449714), Haliotis iris (AF492442), and Pandalus platyceros (AY449715, AY449716), and cercozoans Massisteria marina (AF174374), Heteromita globosa (U42447), Euglypha rotunda (X77692), Cercomonas sp. (U42449, U42450), Cercomonas longicauda (AF101052), and Chlorarachnion reptans (U03477). Cercozoans were used as the outgroup for phylum Haplosporidia based on the phylogenetic results obtained by Reece et al. (2004). Parsimony analysis was done in PAUP version 4.0b10 (Swofford, 2002) using default settings.

RESULTS

Geographical distribution and prevalence Although zebra mussel samples from 11 countries across Europe were examined by histological analysis, haplosporidian infection was detected only from the Rhine and Meuse river basins in France, Germany, and The Netherlands (Fig. 1). Moreover, infection was only intermittently observed in this region during the 17-year sampling period, and when the parasite was detected, prevalence was typically