Observation of a Bonamia sp. infecting the oyster Ostrea stentina in Tunisia, and a consideration of its phylogenetic affinities

Journal of Invertebrate Pathology 103 (2010) 179–185

Contents lists available at ScienceDirect

Journal of Invertebrate Pathology journal homepage: www.elsevier.com/locate/jip

Observation of a Bonamia sp. infecting the oyster Ostrea stentina in Tunisia, and a consideration of its phylogenetic affinities Kristina M. Hill a, Ryan B. Carnegie a,*, Nejla Aloui-Bejaoui b, Refka El Gharsalli b, Delonna M. White a, Nancy A. Stokes a, Eugene M. Burreson a a b

Virginia Institute of Marine Science, College of William and Mary, Gloucester Point, Virginia 23062, USA Institut National Agronomique de Tunisie, 43, Avenue Charles Nicolle, 1082 Tunis, Tunisia

a r t i c l e

i n f o

Article history: Received 9 November 2009 Accepted 23 December 2009 Available online 29 December 2009 Keywords: Bonamia Ostrea stentina Oyster Microcell Haplosporidian Phylogeny

a b s t r a c t The small non-commercial oyster Ostrea stentina co-occurs with commercially important Ostrea edulis in the Mediterranean Sea, yet its disposition with respect to the destructive pathogens Bonamia ostreae and Marteilia refringens is unknown. We began an evaluation of the Bonamia spp. infection status of O. stentina from Hammamet, Tunisia, in June 2007 using polymerase chain reaction diagnostics followed by histology and in situ hybridization. Of 85 O. stentina sampled, nine were PCR-positive for a Bonamia sp. using a Bonamia genus-specific assay; of these nine, one displayed the uninucleate microcells associated with oyster hemocytes characteristic of Bonamia spp. There was no associated pathology. DNA sequencing of the parasite from this one infected individual revealed it to be of a member of the Bonamia exitiosa/Bonamia roughleyi clade, an identification supported by positive in situ hybridization results with probes specific for members of this clade, and by the morphology of the parasite cells: nuclei were central, as in B. exitiosa, not eccentric, as in B. ostreae. There is no basis for identifying the Tunisian parasite as either B. exitiosa or B. roughleyi, however, as these species are genetically indistinguishable. Likewise, there is no basis for identifying any of the other Bonamia spp. with affinities to the B. exitiosa/B. roughleyi clade, from Argentina, Australia, Spain, and the eastern USA, as one or the other of these named species. Though they are clearly distinct from Bonamia perspora and B. ostreae, justification for drawing species boundaries among the primarily austral microcells with affinities to B. exitiosa and B. roughleyi remains elusive. Ó 2010 Elsevier Inc. All rights reserved.

1. Introduction Ostrea stentina Payraudeau, 1826 is an oyster species occurring in the Mediterranean Sea, primarily along the northern African coast, but with a distribution southward possibly to South Africa (Carriker and Gaffney, 1996; Lapègue et al., 2006). It co-occurs with Ostrea edulis L., the European flat oyster, in the northern parts of this range. Unlike O. edulis, however, O. stentina has little economic potential due to its small size (Angell, 1986; Carriker and Gaffney, 1996). Because it is non-commercial, O. stentina has received no parasitological attention, even though it inhabits waters in which economically significant parasites of O. edulis including Bonamia ostreae Pichot et al., 1980 and Marteilia refringens Grizel et al., 1974 are enzootic (Tiscar et al., 1991; Virvilis and Angelidis, 2006; Carrasco et al., 2008). Its role in the epizootiology of these parasites in Mediterranean coastal environments is not known. Ostrea stentina became significant in the context of a larger phylogeographic study of Bonamia spp., the generally non-spore* Corresponding author. Fax: +1 804 684 7796. E-mail address: [email protected] (R.B. Carnegie). 0022-2011/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.jip.2009.12.011

forming ‘‘microcell” haplosporidian parasites of oysters. Two Bonamia spp. were recently observed in the closely related, if not conspecific (Shilts et al., 2007), oyster Ostrea equestris Say, 1834 in North Carolina, USA. The first of these Bonamia spp. is undescribed (Burreson et al., 2004), but has affinities to the ‘‘southern hemispheric clade” (Carnegie and Cochennec-Laureau, 2004) of Bonamia spp. containing Bonamia roughleyi (Farley et al., 1988) Cochennec-Laureau et al., 2003, a parasite of the oyster Saccostrea glomerata in Australia, and Bonamia exitiosa Hine et al., 2001, from the oyster Ostrea chilensis Philippi, 1845 in New Zealand. The second is a novel species, Bonamia perspora Carnegie et al., 2006, that may be sister to B. ostreae. Particularly given the possible synonymy of O. stentina and O. equestris, the possibility that O. stentina would, like O. equestris, harbor one Bonamia sp. or another became an obvious consideration. To address this, we began a preliminary molecular and histological evaluation of O. stentina inhabiting a coastal location in Tunisia in which we generated the first observations of Bonamia sp. infection of O. stentina. Unexpectedly, we found a relative of B. roughleyi and B. exitiosa, rather than B. ostreae, to be associated with the O. stentina we sampled.

180

K.M. Hill et al. / Journal of Invertebrate Pathology 103 (2010) 179–185

2. Materials and methods Ostrea stentina (n = 85) were collected by a diver in June 2007 from a marina at Hammamet, Tunisia (36°220 N, 10°330 E; Fig. 1). Shell heights were measured, and oysters were shucked, with small pieces of gill and mantle tissue (3–5 mm3) preserved in 95% ethanol for molecular analyses. Sections of remaining gill, mantle, and digestive gland tissues were fixed for standard histopathology in Davidson’s fixative (Shaw and Battle, 1957), and then

transferred to 70% ethanol. Both sets of tissues were shipped by commercial courier to the Shellfish Pathology Laboratory at the Virginia Institute of Marine Science for further processing. 2.1. DNA extraction, Bonamia-generic PCR Tissue from each oyster sample was dried to evaporate ethanol and placed into lysis solution, and DNA was extracted using a QIAamp DNA Kit (QIAGEN, Valencia, CA, USA). DNA was quantified using

Fig. 1. Collection location. (A) Area of collection (box) in the larger context of the Mediterranean Sea. (B) Detail of highlighted area, illustrating location of collection () southwest of Hammamet in northeastern Tunisia. Images adapted from Google Maps.

K.M. Hill et al. / Journal of Invertebrate Pathology 103 (2010) 179–185

a GeneQuant pro spectrophotometer (Amersham Biosciences, Piscataway, NJ, USA). For initial screening, polymerase chain reaction (PCR) amplification of Bonamia spp. small-subunit (SSU) rDNA was attempted using Bonamia-generic primers BON-319F (50 -TTTGACG GGTAACGGGGAATGCG-30 ) and BON-524R (50 -CTTGCCCTCCGCTGG AATTC-30 ) that were newly designed using MacVector 8.0 (Oxford Molecular Ltd., Oxford, UK) and purchased from Sigma–Genosys (Midlands, TX, USA). A 25-ll total reaction volume contained 1 PCR buffer, 1.5 mM MgCl2, 0.2 mM dNTPs, and 0.4 lg/ll bovine serum albumin (all purchased from Invitrogen Corporation, Carlsbad, CA, USA), each primer at 0.25 lM, 0.024 U/ll AmpliTaq DNA polymerase (Applied Biosystems, Foster City, CA, USA), and 200–250 ng template DNA. PCR conditions consisted of a 4-min initial denaturation at 94 °C, followed by 35 cycles of denaturation at 94 °C for 30 s, annealing at 60 °C for 30 s, and extension at 72 °C for 1 min, and then by a final extension at 72 °C for 7 min. Products were electrophoresed on 2.5% agarose gels, stained with ethidium bromide, and visualized using UV light. A 200-base-pair (bp) product was expected to be diagnostic of the presence of a Bonamia sp. Where suspected Bonamia sp. amplicons were generated, a second PCR was run to generate longer amplicons (>700 bp) for sequencing. The Bonamia-generic ‘‘CF/CR” assay of Carnegie et al. (2000) was used to generate this larger product. Conditions were as above except that each cyclic denaturation was at 94 °C for 1 min, each annealing was at 59 °C for 1 min, and the final extension was at 72 °C for 10 min. Products were visualized as previously described. 2.2. Bonamia sp. small-subunit rDNA gene sequencing One Bonamia-positive oyster from the initial PCR screening (using the generic BON-319F/BON-524R PCR assay) was targeted for DNA sequence characterization. Presumptive Bonamia sp. amplification products from triplicate CF/CR PCR reactions were pooled, purified using a QIAquick PCR Purification Kit (QIAGEN, Valencia, CA, USA), and cloned into the plasmid vector pCR4-TOPO using the TOPO TA Cloning Kit (Invitrogen Life Technologies, Carlsbad, CA, USA). Sequencing reactions on inserts from ten clones were performed using a BigDyeÒ Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, CA, USA) with unlabeled M13/ pUC forward and reverse sequencing primers (New England Biolabs, Ipswich, MA, USA). Sequencing reaction products were electrophoresed on a 16-capillary ABI 3130xl Genetic Analyzer (Applied Biosystems). Sequencing Analysis 5.2 software (Applied Biosystems) was used for base-calling. After initial sequence data were produced from CF/CR PCR products, two additional PCR reactions were run to generate amplicons for complete parasite SSU rDNA sequencing. Primer pairs 16S-A (Medlin et al., 1988) + Bon-745R (Carnegie et al., 2006) were used in a reaction to amplify the 50 end of a putative Bonamia sp. SSU rDNA gene, and Bon-1310F (Carnegie et al., 2006) + 16S-B (Medlin et al., 1988) were used to amplify the 30 end of the gene. PCR reagents and reaction conditions were again as above. Initial denaturation at 94 °C for 4 min was followed by 35 cycles of denaturation at 94 °C for 45 s, annealing at 54 °C for 45 s, and extension at 72 °C for 1 min for the 16S-A + Bon-745R reaction, and 1.5 min for the Bon-1310F + 16S-B reaction, with a final extension at 72 °C for 6 min. Products were visualized as described above; cloning and sequencing followed, again as above. Five clones from each of these latter PCR and cloning exercises were sequenced. Products from both sequencing rounds were aligned in MacVector 8.0 (Oxford Molecular Ltd., Oxford, UK) to generate a consensus sequence. 2.3. Bonamia sp. internal transcribed spacer (ITS) region sequencing PCR products from the same BON-319F/BON-524R-positive oyster were used for generation of putative Bonamia sp. ITS sequences.

181

PCR primers HaploITSf (50 -GGGATAGATGATTGCAATTRTTC-30 , newly designed using MacVector 8.0, Oxford Molecular Ltd., Oxford, UK) and ITS-B (Goggin, 1994) were used in this reaction. Reagent concentrations for PCR were as above. Initial denaturation at 95 °C for 7 min was followed by 40 cycles of denaturation at 95 °C for 1 min, annealing at 57 °C for 1 min, and extension at 72 °C for 1.5 min, and then a final extension at 72 °C for 7 min. Products were visualized as described above. Products from triplicate PCR reactions were pooled, purified using a QIAquick PCR Purification Kit (QIAGEN, Valencia, CA, USA), and cloned and sequenced (twenty clones) as above. Sequences were aligned in MacVector 8.0 and a consensus sequence was compared by BLAST search (Altschul et al., 1997) to the National Center for Biotechnology Information GenBank database. 2.4. Oyster mt16S sequencing To confirm the identity of the oyster host from which Bonamia sp. was amplified and sequenced, the mt16S of this oyster was amplified following Kessing et al. (1989), as performed previously by Kirkendale et al. (2004), and cloned and sequenced as above. 2.5. Sequence alignments and molecular phylogenetics The SSU rDNA sequence of the Bonamia sp. from O. stentina was ClustalW-aligned (in MacVector 8.0, using default settings: an open gap penalty of 10.0, an extend gap penalty of 5.0, and with transitions weighted) with SSU rDNA sequences from Bonamia spp. including B. ostreae (GenBank accession numbers AF262995 and AF192759), B. exitiosa (AF337563), B. roughleyi (AF508801), B. perspora (DQ356000), the Bonamia spp. from Crassostrea ariakensis in the USA (AY542903), O. edulis from Spain (EU016528), Ostrea angasi from Australia (DQ312295), and O. chilensis from Chile (AY860060), and outgroup species Minchinia tapetis (AY449710), Minchinia teredinis (U20319) and Minchinia chitonis (AY449711). Bonamia sp. ITS1-5.8S-ITS2 rDNA sequences from O. stentina were similarly ClustalW-aligned with ITS1-5.8S-ITS2 sequences of B. ostreae (EU709108, EU709110), B. perspora (EU709119, EU709122, EU709126, EU709130), undescribed Bonamia sp. microcells with documented affinity to B. exitiosa and B. roughleyi including those from C. ariakensis and O. equestris in the eastern USA (EU709050, EU709052, EU709053, and EU709054), O. angasi from southeastern Australia (DQ312295), and O. chilensis from Chile (AY539840), and one final undescribed Bonamia sp. from O. edulis in Italy (EU672891) that was related to an unpublished observation. Independent parsimony analyses of SSU rDNA and of ITS1-5.8S-ITS2 rDNA sequences were performed in PAUP version 4.0d81 (Swofford, 2002) using default settings. 2.6. Histology and in situ hybridization Davidson’s-fixed tissues from a Bonamia-generic PCR-positive oyster were processed using standard techniques. They were dehydrated, cleared with xylene and embedded in paraffin; sectioned at 6 lm and mounted on glass slides; deparaffinized, rehydrated, and stained with hematoxylin and eosin; and dehydrated, cleared, mounted, coverslipped, and examined on an Olympus BX51 light microscope. Entire sections were evaluated under oil at 1000X magnification. Bonamia sp. cell diameters were measured using an Olympus DP71 camera and imaging system. In situ hybridization (ISH) was performed on tissue sections from the same oyster using a cocktail of three 50 digoxigenin-labeled probes specific for closely related members of the B. roughleyi-B. exitiosa clade (CaBon166: 50 -CGAGC AGGGTTTGTCACGTAT-30 ; CaBon461: 50 -TTCCGAATAGGCAACCGA AG-30 ; and CaBon1704: 50 -CAAAGCTTCTAAGAACGCGCC-30 ), following a protocol published previously (Stokes and Burreson, 2001).

182

K.M. Hill et al. / Journal of Invertebrate Pathology 103 (2010) 179–185

A Bonamia sp.-infected C. ariakensis section from the USA was used as a control.

3. Results Evaluation of 85 O. stentina from Hammamet, Tunisia, sampled in June 2007 revealed nine oysters (10.6%) PCR-positive for a Bonamia sp. using the more sensitive BON-319F/BON-524R PCR assay. One oyster of these nine was subsequently PCR-positive for Bonamia sp. using the less sensitive CF/CR protocol. Subsequent histopathological examination of the nine BON-319F/BON-524R PCRpositive oysters confirmed the presence of small numbers of primarily uninucleate Bonamia sp. cells but also binucleate stages and at least one plasmodium (Fig. 2) in the hemolymph of one oyster’s gill and mantle. These were the only organs represented in the histological sample for this particular oyster. Bonamia sp. cells were usually intrahemocytic, with as many as five parasite cells per hemocyte observed. Uninucleate Bonamia sp. cells were typically spherical and 2.4 ± 0.1 lm in diameter (mean ± SEM, n = 40; range 1.7–2.9 lm). Bonamia sp. cell nuclei were more or less central in position. Hemocytosis in response to this infection was not observed in the organs available for evaluation. PCR-cloning and DNA sequencing of the SSU rDNA from the Bonamia sp. infecting O. stentina produced a 1749-bp sequence that was nearly identical to sequences from southern hemispheric B. exitiosa and B. roughleyi and their relatives. This sequence was submitted to GenBank and given accession number GQ385242. Sequence similarity was 99.1–99.7% to B. exitiosa (depending on the treatment of a 10-bp sequence unique to B. exitiosa sequence AF337563 as ten sequence changes, as one sequence change, or as a sequencing error and thus not to be considered); 99.4% to B. roughleyi; 99.5–100% to the Bonamia sp. from O. edulis in Galicia, Spain (depending on the identity of eight unresolved bases); 99.7% to the Bonamia sp. from O. chilensis in Chile; and 100% to the Bonamia spp. from O. angasi in Australia and C. ariakensis and O. equestris from the United States. Parsimony analysis united these SSU rDNA sequences, with strong support, on a clade sister to a more weakly supported B. perspora–B. ostreae clade (Fig. 3).

PCR-cloning and DNA sequencing of the parasite’s ITS1-5.8SITS2 region produced four 462–465-bp sequences from among twenty clones that differed only slightly. Sequences of two clones (GenBank accession number GU356033) differed from the most common sequence (GU356032, which occurred in fifteen of twenty clones) in possessing TC substitutions at two positions. Another two clones were characterized by an AA deletion (GU356034). One clone possessed a unique TA insertion, followed sixteen positions later by a T deletion (GU356035). Parsimony analysis of these sequences placed them on an unresolved Bonamia clade that included B. exitiosa, the Bonamia sp. from C. ariakensis and O. equestris in the USA, and the Bonamia sp. from O. angasi in Australia (Fig. 4). A Bonamia sp. sequenced from O. edulis from Italy (GenBank accession number EU672891, unpublished) appears from a short, partial ITS1 sequence to have an affinity to this group, but the Bonamia sp. sequenced from O. chilensis in Chile (AY539840) does not. In situ hybridization on the Bonamia sp.-infected O. stentina section was positive, indicating the presence of SSU rRNA belonging to a member of the ‘‘southern hemispheric clade” of Bonamia spp. (Fig. 5). DNA sequencing of the mt16S region of the oyster host generated a 447-bp sequence that was a perfect match to O. stentina (GenBank accession number DQ180744; Shilts et al., 2007), confirming the host’s identity.

4. Discussion Together, microscopic evidence and DNA sequencing confirmed the presence of a relative of B. exitiosa and B. roughleyi in the Mediterranean Sea, in O. stentina in Tunisia. Tissue tropism clearly suggests a member of the genus Bonamia, rather than Mikrocytos, in that hemocytes, rather than vesicular connective tissue cells, are the infected host cells (Balouet et al., 1983; Carnegie and Cochennec-Laureau, 2004). Cell appearance suggests a relative of B. exitiosa, rather than B. ostreae, in that the uninucleate cells observed were larger than typical B. ostreae (see Abollo et al., 2008, for comparison), and did not display the typically eccentric nucleus of B. ostreae that may be recognized in images from numerous publica-

Fig. 2. Bonamia sp. in the mantle of Ostrea stentina observed histologically. A single uninucleate Bonamia sp. cell (arrow) is identified, with a second uninucleate cell just out of focus to its right. A trinucleate Bonamia sp. plasmodium (arrowhead) is also present.

K.M. Hill et al. / Journal of Invertebrate Pathology 103 (2010) 179–185

183

Fig. 4. Parsimony analysis of ITS1-5.8S-ITS2 sequences. Numbers at nodes represent means of 1000 bootstrap replicates. Fig. 3. Parsimony analysis of SSU rDNA genes. Representatives of the sister genus Minchinia were included as an outgroup. Numbers at nodes represent means of 1000 bootstrap replicates. The value 55 on the upper part of the tree marks the node at which B. exitiosa (AF337563) diverges from B. roughleyi (AF508801) and the other parasites in that clade.

tions (e.g., Pichot et al., 1980; Elston et al., 1986; Mialhe et al., 1988; Cochennec et al., 1998; Tiscar et al., 2002; Marty et al., 2006; Abollo et al., 2008). Phylogenetic analyses of ribosomal gene sequences and in situ hybridization provide compelling support for the presence of a relative of B. exitiosa and B. roughleyi, and not B. ostreae, in these oysters sampled from Tunisia. While this represents the first record of a Bonamia sp. in O. stentina, the specific identity of the parasite is unresolved. Numerous Bonamia sp. parasites have been observed in various hosts and locations in recent years, and all have been casually identified as B. exitiosa. Corbeil et al. (2006) identified a Bonamia sp. observed in Ostrea angasi Sowerby, 1871 in southeastern Australia as B. exitiosa, and implied that parasites observed by others in O. chilensis in Chile (Campalans et al., 2000), in Crassostrea ariakensis Fujita, 1913 in the southeastern USA (Burreson et al., 2004), and in Ostrea puelchana d’Orbigny, 1841 in Argentina (Kroeck and Montes, 2005) were conspecific. Abollo et al. (2008) likewise identified the Bonamia sp. recently detected in O. edulis in Spain as B. exitiosa, basing this identification on the assertion by the World Organization for Animal Health Reference Laboratory for infection by B. exitiosa that the parasites observed in O. angasi in Australia, C. ariakensis in the USA, O. chilensis from Chile as well as New Zealand, and O. puelchana from Argentina were all indeed B. exitiosa (López-Flores et al., 2007). Yet identification of all these Bonamia sp. parasites as B. exitiosa is complicated by one important consideration: the close phylogenetic affinity of a second Bonamia species, B. roughleyi, to

this same group. Potentially more informative ITS1-5.8S-ITS2 rDNA sequences for B. roughleyi do not exist, but the SSU rDNA sequence of B. roughleyi is nearly identical to that of all those parasites identified as B. exitiosa. SSU rDNA-based parsimony analysis performed by Abollo et al. (2008) placed B. roughleyi, B. exitiosa, and the other undescribed Bonamia sp. parasites in a single, strongly supported clade distinct from a sister clade occupied by B. ostreae and B. perspora; the Bonamia sp. parasites identified as B. exitiosa were paraphyletic in their analysis. Our SSU rDNA-based analysis has reproduced this result. Parsimony analysis of ITS1-5.8S-ITS2 sequences, which unfortunately could not incorporate sequence data from B. roughleyi, provided no better resolution of this clade, with one possible exception: the ITS1 sequence of the Bonamia sp. from Chilean O. chilensis diverges from the other sequences. While parsimony analyses of ribosomal RNA genes provide no compelling argument for identifying the Tunisian Bonamia sp. and other unidentified parasites as B. exitiosa, rather than B. roughleyi, it is the case that none of these parasites besides B. roughleyi possess the ‘‘A” that Cochennec-Laureau et al. (2003) identified as diagnostic for B. roughleyi (at position 762 in B. roughleyi sequence AF508801, corresponding with a ‘‘G” at position 773 in B. exitiosa sequence AF337563, and a ‘‘G” at position 807 in B. ostreae sequence AF262995). Based on the diagnostic PCR-RFLP assays presented by these authors, one component of which would involve the attempted restriction digestion of a PCR amplicon at this site with HaeII (digesting a product from B. exitiosa or B. ostreae, but not B. roughleyi), all of these Bonamia spp. would be identified as B. exitiosa (B. ostreae having been ruled out using BglI digestion at a second SSU rDNA position). On the other hand, only the type SSU rDNA sequence for B. exitiosa (AF337563; Hine et al., 2001) possesses a 10-bp tandem repeat sequence near its 50 end (posi-

184

K.M. Hill et al. / Journal of Invertebrate Pathology 103 (2010) 179–185

Fig. 5. In situ hybridization of Bonamia sp.-specific probes to parasite cells in two clusters (arrows) in Ostrea stentina gills.

tions 593–602 in B. exitiosa sequence AF337563, which would be just 30 of position 591 in B. roughleyi sequence AF508801). All the undescribed Bonamia spp. resemble B. roughleyi in lacking this repeat. To our knowledge, the B. roughleyi-diagnostic ‘‘A” has never been recovered in any subsequent sequencing of B. roughleyi; there is no published evidence of more than a single presumptively B. roughleyi-infected oyster ever having been sequenced at all. There is also no documentation of the diagnostic PCR-RFLP assays ever having been performed on presumptively B. roughleyi-infected oysters beyond the single oyster from which the SSU rDNA of B. roughleyi was originally sequenced. Likewise, there is no evidence that the unusual 10-bp tandem repeat in the type SSU rDNA sequence of B. exitiosa has ever been recovered in subsequent sequencing efforts. Given all this uncertainty concerning basic DNA sequence profiles of these parasites in their type localities, the use of any particular SSU rDNA sequence signature, like the ‘‘A” in B. roughleyi or the 10-bp tandem repeat in B. exitiosa, to identify species in this clade is not justified. Unfortunately, neither parasite cell morphology nor ultrastructure yields any insight concerning species distinctions among this group either (e.g., Lohrmann et al., 2009). There is at present, therefore, no defensible basis – morphological, ultrastructural, or molecular – for drawing species boundaries among the closely related, primarily austral microcells. We can only conclude that as a group they are distinct from B. ostreae and B. perspora. While they may belong to a single species, it is also possible that additional loci will reveal phylogenetic divergence among these parasites, or that phenotypic differences exist in key traits such as physical tolerances, host range, or pathogenicity. While only limited histological material was evaluated, there was no indication of any pathology in the O. stentina that could have been attributed to Bonamia sp. infection. This is similar to our observations of Bonamia sp. in O. equestris in North Carolina, USA. Infections in O. equestris are typically very light in intensity, and low in prevalence (<10%; Carnegie and Burreson, unpublished). It is also similar to the observation of Haplosporidium nelsoni Haskin et al., 1966, the devastating parasite of Crassostrea virginica Gmelin, 1791, in Crassostrea gigas Thunberg, 1793, which is believed to be its natural host (Burreson et al., 2000). Light infection and low prevalence in O. stentina and O. equestris contrast sharply, however, with the observations of Bonamia sp. in O. chilensis (Dinamani et al., 1987), O. angasi

(Hine and Jones, 1994), C. ariakensis (Burreson et al., 2004), and O. edulis (Abollo et al., 2008). We might ask whether epizootic parasitic infection and host mortality in O. chilensis, C. ariakensis, and O. edulis reflect new and non-equilibrium host–parasite associations. The generally innocuous infections in O. stentina and O. equestris, on the other hand, may reflect more established relationships. Recall once again the possible synonymy, supported by the latest phylogenies, not only of the various undescribed Bonamia spp. with B. exitiosa and B. roughleyi (Abollo et al., 2008, and this study), but also of the hosts O. stentina from the Mediterranean and Atlantic coasts of Africa, O. equestris from the western Atlantic, and Ostrea auporia Dinamani and Beu, 1981 from northern New Zealand (Carriker and Gaffney, 1996; Shilts et al., 2007). It is possible that the lack of observed, significant pathology in O. stentina and O. equestris reflects new host–parasite associations, but interactions in which the host is relatively resistant. It is also possible, however, that the association is long-established and stable, and predates the dispersal of O. stentina/O. equestris/O. auporia in the years prior to the initial descriptions of these species in the early 19th century. The distribution of the B. exitiosa/B. roughleyi-like Bonamia sp. does correspond roughly to the distribution of O. stentina/O. equestris/O. auporia, though infection in O. auporia itself has yet to be confirmed. The range expansion of this parasite, if indeed it is a single species, was not necessarily a recent one. Acknowledgments The Société d’études et d’aménagement de la Marina Hammamet provided assistance with oyster collecting. Rita Crockett and Susan Denny performed the histopathology. We thank our colleagues from the New South Wales (Australia) Department of Primary Industries, Jeffrey Go, Cheryl Jenkins, Ian Marsh, Zoe Spiers, and Wayne O’Connor, for their improvements to the manuscript and helpful discussions on bonamiasis in Australia. This is VIMS Contribution Number 3057. References Abollo, E., Ramilo, A., Casas, S.M., Comesaña, P., Cao, A., Carballal, M.J., Villalba, A., 2008. First detection of the protozoan parasite Bonamia exitiosa (Haplosporidia)

K.M. Hill et al. / Journal of Invertebrate Pathology 103 (2010) 179–185 infecting flat oyster Ostrea edulis grown in European waters. Aquaculture 274, 201–207. Altschul, S.F., Madden, T.L., Schäffer, A.A., Zhang, J., Miller, W., Lipman, D.J., 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25, 3389–3402. Angell, C.L., 1986. The biology and culture of tropical oysters. International Center for Living Aquatic Resources Management, Studies and Reviews 13. Manila, Philippines. Balouet, G., Poder, M., Cahour, A., 1983. Haemocytic parasitosis: morphology and pathology of lesions in the French flat oyster, Ostrea edulis L. Aquaculture 34, 1–14. Burreson, E.M., Stokes, N.A., Friedman, C.S., 2000. Increased virulence in an introduced pathogen: Haplosporidium nelsoni (MSX) in the eastern oyster Crassostrea virginica. J. Aquat. Anim. Health 12, 1–8. Burreson, E.M., Stokes, N.A., Carnegie, R.B., Bishop, M.J., 2004. Bonamia sp. (Haplosporidia) found in non-native oysters Crassostrea ariakensis in Bogue Sound, North Carolina. J. Aquat. Anim. Health 16, 1–9. Carnegie, R.B., Cochennec-Laureau, N., 2004. Microcell parasites of oysters: recent insights and future trends. Aquat. Living Resour. 17, 519–528. Carnegie, R.B., Barber, B.J., Culloty, S.C., Figueras, A.J., Distel, D.L., 2000. Development of a PCR assay for detection of the oyster pathogen Bonamia ostreae and support for its inclusion in the Haplosporidia. Dis. Aquat. Org. 42, 199–206. Carnegie, R.B., Burreson, E.M., Hine, P.M., Stokes, N.A., Audemard, C., Bishop, M.J., Peterson, C.H., 2006. Bonamia perspora n. Sp. (Haplosporidia), a parasite of the oyster Ostreola equestris, is the first Bonamia species known to produce spores. J. Euk. Microbiol. 53, 232–245. Carriker, M.R., Gaffney, P.M., 1996. A catalogue of selected species of living oysters (Ostreacea) of the world. In: Kennedy, V.S., Newell, R.I.E., Eble, A.F. (Eds.), The Eastern Oyster Crassostrea virginica. Maryland Sea Grant, College Park, Maryland, pp. 1–18. Carrasco, N., Arzul, I., Berthe, F.C.J., Fernández-Tejedor, M., Durfort, M., Furones, M.D., 2008. Delta de l’Ebre is a natural bay model for Marteilia spp. (Paramyxea) dynamics and life-cycle studies. Dis. Aquat. Org. 79, 65–73. Campalans, M., Rojas, P., Gonzalez, M., 2000. Haemocytic parasitosis in the farmed oyster Tiostrea chilensis. Bull. Eur. Assoc. Fish Pathol. 20, 31–33. Cochennec-Laureau, N., Reece, K.S., Berthe, F.C.J., Hine, P.M., 2003. Mikrocytos roughleyi taxonomic affiliation leads to the genus Bonamia (Haplosporidia). Dis. Aquat. Org. 54, 209–217. Cochennec, N., Renault, T., Boudry, P., Chollet, B., Gerard, A., 1998. Bonamia-like parasite found in the Suminoe oyster Crassostrea rivularis reared in France. Dis. Aquat. Org. 34, 193–197. Corbeil, S., Arzul, I., Robert, M., Berthe, F.C.J., Besnard-Cochennec, N., Crane, M.St.J., 2006. Molecular characterization of an Australia isolate of Bonamia exitiosa. Dis. Aquat. Org. 71, 81–85. Dinamani, P., Beu, A.G., 1981. Description of a new species of incubatory oyster from northern New Zealand, with notes on its ecology and reproduction. N.Z. J. Mar. Freshwat. Res. 15, 109–119. Dinamani, P., Hine, P.M., Jones, J.B., 1987. Occurrence and characteristics of the haemocyte parasite Bonamia sp. in the New Zealand dredge oyster Tiostrea lutaria. Dis. Aquat. Org. 3, 37–44. d’Orbigny, A., 1841. Voyage dans l’Amérique Méridionale executé pendant les années 1826-1833. Paris. Elston, R.A., Farley, C.A., Kent, M.L., 1986. Occurrence and significance of bonamiasis in European flat oysters Ostrea edulis in North America. Dis. Aquat. Org. 2, 49–54. Farley, C.A., Wolf, P.H., Elston, R.A., 1988. A long-term study of ‘‘microcell” disease in oysters with a description of a new genus, Mikrocytos (g. n.), and two new species, Mikrocytos mackini (sp. n.) and Mikrocytos roughleyi (sp. n.). Fish. Bull. 86, 581–593. Fujita, 1913. Nihon suisan dobutsugaku, vol. 2. Shokabu, Tokyo. Gmelin, J.F., 1791. Systema naturae per regna tria naturae. In: G. E. Beer (Ed.), Editio Decimo Tertia, Aucta, Reformata, Leipzig. vol. 1, pp. 3021–3909. Goggin, C.L., 1994. Variation in the two internal transcribed spacers and 5.8S ribosomal-RNA from five isolates of the marine parasite Perkinsus (Protista, Apicomplexa). Mol. Biochem. Parasitol 65, 179–182. Grizel, H., Comps, M., Bonami, J.R., Cousserans, F., Duthoit, J.L., Le Pennec, M.A., 1974. Recherche sur l’agent de la maladie de la glande digestive de Ostrea edulis Linné. Bull. Inst. Pêches Marit. Maroc. 240, 7–30. Haskin, H.H., Stauber, L.A., Mackin, J.A., 1966. Minchinia nelsoni n. sp. (Haplosporida, Haplosporidiidae): causative agent of the Delaware Bay oyster epizootic. Science 153, 1414–1416.

185

Hine, P.M., Jones, J.B., 1994. Bonamia and other aquatic parasites of importance to New Zealand. New Zealand J. Zool. 21, 49–56. Hine, P.M., Cochennec-Laureau, N., Berthe, F.C.J., 2001. Bonamia exitiosus n. sp. (Haplosporidia) infecting flat oysters Ostrea chilensis in New Zealand. Dis. Aquat. Org. 47, 63–72. Kessing, B., Croom, H., Martin, A., McIntosh, C., McMillan, W.O., Palumbi, S., 1989. The simple fool’s guide to PCR. Department of Zoology, University of Hawaii, Honolulu, Hawaii. Kirkendale, L., Lee, T., Baker, P., Ó Foighil, D., 2004. Oysters of the Conch Republic (Florida Keys): a molecular phylogenetic study of Parahyotissa mcgintyi, Teskeyostrea weberi, and Ostreola equestris. Malacologia 46, 309–326. Kroeck, M.A., Montes, J., 2005. Occurrence of the haemocyte parasite Bonamia sp. in flat oysters Ostrea puelchana farmed in San Antonio Bay (Argentina). Dis. Aquat. Org. 63, 231–235. Lapègue, S., Salah, I.B., Batista, F.M., Heurtebise, S., Neifar, L., Boudry, P., 2006. Phylogeographic study of the dwarf oyster, Ostreola stentina, from Morocco, Portugal and Tunisia: evidence of a geographic disjunction with the closely related taxa, Ostrea aupouria and Ostreola equestris. Mar. Biol. 150, 103– 110. Lohrmann, K.B., Hine, P.M., Campalans, M., 2009. Ultrastructure of Bonamia sp. in Ostrea chilensis in Chile. Dis. Aquat. Org. 85, 199–208. Lopez-Flores, I., Suarez-Santiago, V.N., Longet, D., Saulnier, D., Chollet, B., Arzul, I., 2007. Characterization of actin genes in Bonamia ostreae and their application to phylogeny of the Haplosporidia. Parasitology 134, 1941– 1948. Marty, G.D., Bower, S.M., Clarke, K.R., Meyer, G., Lowe, G., Osborn, A.L., Chow, E.P., Hannah, H., Byrne, S., Sojonky, K., Robinson, J.H., 2006. Histopathology and a real-time PCR assay for detection of Bonamia ostreae in Ostrea edulis cultured in western Canada. Aquaculture 261, 33–42. Medlin, L., Elwood, H.J., Stickel, S., Sogin, M.L., 1988. The characterization of enzymatically amplified eukaryotic 16S-like rRNA-coding regions. Gene 71, 491–499. Mialhe, E., Bachère, E., Chagot, D., Grizel, H., 1988. Isolation and purification of the protozoan Bonamia ostreae (Pichot et al., 1980), a parasite affecting the flat oyster Ostrea edulis L. Aquaculture 71, 293–299. Payraudeau, B.C., 1826. Catalogue déscriptif et methodique des annelides et de mollusques de l’Isle de Corse. Paris. Philippi, R.A., 1845. Abbildungen und Beschreibung neuer oder wenig bekannter Conchylien, vol. 2. Theodore Fischer, Cassel, Germany. Pichot, Y., Comps, M., Tigé, G., Grizel, H., Rabouin, M.-A., 1980. Recherches sur Bonamia ostreae gen. n., sp. n., parasite nouveau de l’huître plate Ostrea edulis. Rev. Trav. Inst. Pêches Marit. 43, 131–140. Say, T., 1834. American conchology, or descriptions of the shells of North America illustrated by coloured figures from original drawings executed from nature, vol. 1, School Press, New Harmony, Indiana. Shaw, B.L., Battle, H.I., 1957. The gross and microscopic anatomy of the digestive tract of the oyster Crassostrea virginica (Gmelin). Can. J. Zool. 35, 325–346. Shilts, M.H., Pascual, M.S., Ó Foighil, D., 2007. Systematic, taxonomic and biogeographic relationships of Argentine flat oysters. Mol. Phylogenet. Evol. 44, 467–473. Sowerby Jr., G., 1871. Monograph of the genus Ostraea. In: Reeve, L. (Ed.), Conchologia Iconica; or Illustrations of the Shells of Molluscous Animals, vol. 18. L. Reeve & Co., London. Stokes, N.A., Burreson, E.M., 2001. Differential diagnosis of mixed Haplosporidium costale and Haplosporidium nelsoni infections in the eastern oyster, Crassostrea virginica, using DNA probes. J. Shellfish Res. 20, 207–213. Swofford, D.L., 2002. PAUP: Phylogenetic Analysis Using Parsimony ( and other methods), Version 10. Sinauer Associates, Sunderland, Massachusetts. Thunberg, C.P., 1793. Techning och beskrifning pa en stor Ostronsort ifran Japan. K. Svenska Vetensk. Akad., Handlingar, 14, 140–142. Tiscar, P.G., Zizzo, N., Tempesta, M., 1991. Su alcune patologie riscontrate in ostriche piatte (Ostrea edulis) provenienti da banco naturale. Boll. Soc. Itl. Patol. Ittica 7, 13–18. Tiscar, P.G., Quaglio, F., Della Salda, L., Ceschia, G., Delgado, M.L., Restani, R., 2002. Presenza di Bonamia ostreae in ostriche piatte (Ostrea edulis) del Nord Adriatico. Boll. Soc. It. Patol. Ittica 35, 2–10. Virvilis, C., Angelidis, P., 2006. Presence of the parasite Marteilia sp. in the flat oyster (Ostrea edulis L.) in Greece. Aquaculture 259, 1–5.