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Haplosporidium nelsoni (MSX) rDNA detected in oysters from the Gulf of Mexico and the Caribbean Sea.

ABSTRACT The known range of the oyster pathogen Haplosporidium nelsoni Haskin, Stauber, and Mackin (MSX) extends along the North American Atlantic coast from Nova Scotia to Florida. Our study demonstrates that H. nelsoni is also present throughout the Gulf of Mexico. Thirty of 41 oysters (73%) sampled from sites ranging from Florida to as far south as Venezuela were positive for MSX by PCR amplification of the ribosomal rRNA gene complex. DNA sequences cloned from oysters positive for H. nelsoni were [greater than or equal to] 99% identical to H. nelsoni and clearly divergent (<89% identical) from known haplosporidian congeners, including Haplosporidium costale Wood and Andrews (SSO). The absence of MSX epizootics in the Gulf of Mexico, despite the wide distribution of H. nelsoni infections, raises questions about pathogenicity and the host-parasite relationship in subtropical latitudes.

KEY WORDS: Crassostrea gasar, Crassostrea rhizophorae, Puerto Rico, Venezuela, MSX


In the last century, numerous introductions of nonnative aquaculture species have inadvertently extended the ranges of diseases and exposed native organisms to novel pathogens against which they have little or no defense. Often, these introduced pathogens have become permanently established with little possibility for eradication. Repeated introductions of Crassostrea virginica Gmelin and coincidental warmer weather have extended the range of Perkinsus marinus Mackin, Owen, and Collier, the causative agent of Dermo disease, to previously unexposed oyster populations of the northeastern United States (Ford 1996). Likewise, Bonamia ostreae Pichot, a protozoan that causes heavy mortalities in the flat oyster Ostrea edulis Linne, was likely introduced to Europe and the western Atlantic via imports of infected shellfish from California (Friedman & Perkins 1994).

Haplosporidium nelsoni (MSX: multinucleated sphere X) has spread in a similar manner. Recent studies have shown that H. nelsoni commonly parasitizes the Pacific oyster Crassostrea gigas Thunberg in Japan, Korea, and California (Friedman 1996; Burreson et al. 2000; Kamaishi & Yoshinaga 2002). However, because H. nelsoni does not cause heavy mortalities in C. gigas populations, the parasite remained undetected until it ravaged C. virginica populations of the mid Atlantic coast of the United States in the mid to late-1900s (Haskin et al. 1966). Crassostrea virginica stocks were likely first exposed to H. nelsoni via introductions of C. gigas from the Pacific Ocean (Burreson et al. 2000).

Neither H. nelsoni infections nor MSX disease have been reported among oysters in the Gulf of Mexico. Only one published study to date has included Gulf of Mexico oysters for diagnosis of H. nelsoni. In that study, the parasite was not present in oysters from Grand Isle, LA (Russell et al. 2004). The absence of H. nelsoni is surprising, given that the Gulf of Mexico is relatively close to enzootic regions (Atlantic coast of Florida) and is typified by warm water temperatures and salinities that readily support MSX epizootic events in Atlantic estuaries. In this study we undertook a modest geographical survey for H. nelsoni and demonstrate that H. nelsoni occurs among oyster populations throughout the Gulf of Mexico from Florida to as far south as Venezuela.


Sample Collection

Crassostrea virginica were collected throughout the Gulf of Mexico (Fig. 1, Table 1). In addition, C. rhizophorae tissue samples were obtained from Puerto Rican oysters. Whereas these oysters originated in Puerto Rico, there is some uncertainty as to whether they may have resided for a period at Harbor Branch Oceanographic Insitute (Fort Pierce, FL). Whereas these oysters may have been exposed to water from the Atlantic Ocean, they serve as important indicators as to whether C. rhizophorae can carry H. nelsoni. Oysters (C. rhizophorae and/or C. gasar) were also obtained from Venezuela. Adductor muscle from oysters was dissected and preserved in 95% ethanol. To prevent cross-contamination during dissection, scalpels were washed with 10% bleach and rinsed with deionized water between oysters. Total genomic DNA was extracted from the adductor muscles using DNEasy kits (Qiagen).


Polymerase Chain Reaction and DNA Sequencing

A 564-bp fragment of the MSX small subunit rRNA gene was amplified from the DNA extracts using the MSXA (5'GCATTAGGTTTCAGACC-3') and MSXB (5'ATGTGTTGGTGACGCTAACCG-3') primers (Stokes et al. 1995). Polymerase chain reactions (PCR) were carried out in 20 [micro]L containing 1x Optiprime buffer #4 (Stratagene: 10 mM Tris-HCl, 3.5 mM Mg[Cl.sub.2], 75 mM KCl), 200 [micro]M of each dNTP, 40 nM each of MSXA and MSXB primers, 0.5 units of Taq polymerase (Fisher), and 1 [micro]L of DNA extract. Cycling conditions for PCR began with an initial denaturation of the sample at 94[degrees]C for 2 min followed by 34 cycles of 94[degrees]C for 30 s, 55[degrees]C for 1 min, and elongation at 72[degrees]C for 1 min +4 s [cycle.sup.-1]. After cycling, a final elongation period at 72[degrees]C was performed for 15 min. Positive and negative control reactions, loaded with 1 [micro]L of 10 ng [micro][L.sup.-1] pMSX414 plasmid (provided by N. Stokes) or 1 [micro]L of water, respectively, were performed with each set of amplifications.

Amplification products were separated by electrophoresis on 2% agarose (IBI) gels and stained with SYBR Gold (Invitrogen). Putative MSX amplicons were ligated into pCR-TOPO2.1 cloning vector and used to transform chemically-competent Machl E. coli cells (Invitrogen). Transformants were plated on LB agar containing 50 [micro]g [mL.sup.-1] kanamycin (MP Biomedicals) and 100 [micro]g X-gal and grown overnight at 37[degrees]C. Colonies selected for sequencing were subcultured overnight at 37[degrees]C in 2 mL LB broth containing 50 [micro]g [mL.sup.-1] kanamycin. Plasmid DNA was isolated with the UltraClean Standard Mini Plasmid Prep Kit (MO BIO Laboratories). Sequencing reactions (10 [micro]L) contained 40 mM Tris-HCl (pH 9.0), 1 mM Mg[Cl.sub.2], 1.6 pmole of M13 reverse primer, 2 [micro]L of BigDye terminator v 1.1 mix (Applied Biosystems), and 500 ng of plasmid DNA. Sequencing reactions were amplified for 25 cycles consisting of denaturation at 96[degrees]C for 30 s, annealing at 50[degrees]C for 15 s, and elongation at 60[degrees]C for 4 min on a PTC-200 thermal cycler (MJ Research). Capillary sequencing was performed with an ABI 310 sequencer (Applied Biosystems).

Sequence analysis was performed with Vector NTI 10 software (Invitrogen). All alignments were made with exclusion of gaps, and phylogenetic trees were generated using Kimura's correction (Kimura 1983).


Thirty of the 41 oysters that we sampled were positive for MSX. At all of the sites that we sampled from, DNA sequences of the PCR products (Range: 288-543 bp, mean = 464 bp) confirmed that the parasite detected was H. nelsoni. To assess the degree of variation in rRNA sequence in multiple clones from the same oyster DNA sample, we aligned a 215-bp region represented in clones from all oysters (Fig. 2). With the DNA extracts for which we had paired clone sequences (18 clones from 9 oyster DNA extracts), sequences were 99.4 [+ or -] 0.5% identical between pairs. 94.4% of the bases were identical when the same region among all of these clones was aligned. Further inspection revealed that all of the differences were attributable to single nucleotide differences that were not represented in a given extract's paired clone sequence. Thus, most of the variation among clones was likely a function of sequencing error. Assembled sequences for each DNA extract yielded 16 sequences with a shared region of 288 bases. All sixteen sequences were aligned with the same region in H. nelsoni (GenBank U19538), H. costale (GenBank U20858), Haplosporidium louisiana Sprague (GenBank U47851), Haplosporidium lusitanicum Azevedo (GenBank AY449713), Haplosporidium montforti n. sp. Azevedo (GenBank DQ219484), and Haplosporidium pickfordi Barrow (GenBank AY452724). Sites that exhibited degeneracies introduced by sequencing error were excluded from the alignment. All of the clone sequences showed [greater than or equal to] 99% identity to H. nelsoni (Fig. 3). Haplosporidium costale and H. lusitanicum, the most similar congeners, were <89% identical to the clone sequences.


We are confident that these results are not false positives for three reasons. (1) The primers that we used to amplify MSX are specific to H. nelsoni and do not amplify H. costale, H. louisiana, or Minchinia teredinis Hillman, Ford, and Haskin (Stokes et al. 1995). (2) All dissection equipment was washed with bleach between samples to prevent cross-contamination. Contamination of DNA samples is contraindicated by the presence of both positive and negative amplifications within our data sets. For instance, of nine oysters that we sampled from Port Aransas, TX, only two were positive for MSX. Had contamination in the laboratory environment or from another oyster sample occurred during dissection, DNA extraction, or during setup of the PCR reactions, positive amplifications would have been more frequent. (3) We performed a second set of PCR amplifications for detection of H. nelsoni and P. marinus (using primers described in Marsh et al. 1995) with additional samples from Apalachicola Bay, FL; Cedar Key, FL; and Port Aransas, TX. Had cross contamination among samples occurred at the time of dissection or DNA extraction, false positives would have been evident as positive reactions for both pathogens in all of these samples. On the contrary, we found that some animals carried only one or the other pathogen but not both. Among the 20 samples, 9 animals tested positive exclusively for MSX and 4 tested positive only for P. marinus. In addition, we are certain that these results derive from H. nelsoni infections in oyster tissue rather than from contamination via environmental source waters. The detection limit of the PCR protocol defined by Stokes et al. (1995) is 10 fg, or ~16,000 template copies. The modified PCR protocol we used in this study used even more stringent conditions (annealing temperature of 55[degrees]C rather than 50[degrees]C) and is likely less sensitive. In a recent study by Audemard et al. (2006), DNA was extracted from 250-mL water samples to monitor Chesapeake Bay for P. marinus. It is unlikely that volumes of environmental water equivalent to that of the tissue mass we sampled (1-3 mg) would yield the H. nelsoni DNA necessary for detection, much less the strong amplifications we observed.


Range extension of bivalve pathogens through intentional or unintentional introductions of shellfish presents a real hazard to native species and commercial aquaculture. Haplosporidium nelsoni, a parasite of C. gigas in the western Pacific Ocean, was likely introduced to North America from Asia (Burreson et al. 2000, Friedman 1996). Haplosporidium nelsoni initially established itself in C. virginica stocks of Delaware Bay and spread throughout populations along the eastern seaboard of the United States (Renault et al. 2000). Though temperatures and salinities in the Gulf of Mexico are amenable to H. nelsoni, MSX has not yet been reported in the Gulf of Mexico (Russell et al. 2004). All major epizootic events in Gulf of Mexico oysters have been caused by P. marinus. In the absence of known H. nelsoni epizootics, research has focused on the epizootiology of Dermo disease, and no geographic survey of H. nelsoni range has yet been published.

Our study demonstrates the presence of H. nelsoni in oysters of the Gulf of Mexico and Caribbean Sea, extending the known range of the parasite to as far south as Venezuela. We found H. nelsoni rDNA in over 70% of the oysters that we analyzed. DNA sequences of PCR products were highly similar ([greater than or equal to] 99%) to a published rDNA sequence for H. nelsoni (Stokes & Burreson 1995) and were clearly divergent from all other haplosporidian rDNA sequences, including that of H. costale (SSO: seaside organism), a congener of H. nelsoni that commonly infects C. virginica. Whereas the census of haplosporidia worldwide (Burreson & Ford 2004, Reece et al. 2004) is likely incomplete, we are confident that these infections represent H. nelsoni. However, the rDNA we sampled may be from strains that are divergent from those in the Atlantic Ocean. The origin of H. nelsoni in the Gulf of Mexico remains unresolved. It is possible that the pathogen was introduced via imports of C. gigas to the Gulf of Mexico during early aquaculture efforts or transplantation of infected C. virginica from the Atlantic Ocean. We are aware of only one published example of intentional introduction of C. gigas to the Gulf of Mexico (Kavanagh 1940). Carlton (1992), however, states that occasional introductions of C. gigas to the Gulf of Mexico have occurred since the 1930s. In addition, C. virginica were occasionally imported during the last 25 y from Chesapeake Bay to the Galveston Bay, TX, area to support the local oyster shucking industry (S. Ray, pers. comm.). Effluent from shucking operations could potentially have introduced H. nelsoni by way of infected tissue or small live oysters. Alternatively, H. nelsoni spores could have been transported via ballast water with the increased shipping traffic between the Atlantic and Pacific oceans after World War II (Burreson & Ford 2004).

Given the warm water temperature and prevalence of infections in the Gulf of Mexico, it is surprising that H. nelsoni has not manifested itself in epizootic events. Oysters from the Gulf of Mexico develop MSX infections and experience heavy mortalities when transplanted to Chesapeake Bay (Ragone-Calvo et al. 2003, Encomio et al. 2005). Why don't oysters succumb to H. nelsoni infections in the Gulf of Mexico? Strain-dependent virulence of MSX has not yet been demonstrated but may explain the resistance of C. virginiCa to Gulf of Mexico H. nelsoni. The Dermo pathogen P. marinus exhibits such a pattern of virulence. Bushek and Allen (1996) demonstrated that virulence of P. marinus from the Atlantic Ocean is higher than that of pathogen strains from the Gulf of Mexico. And Reece et al. (2001) further documented the existence of different Dermo strains along the Gulf and Atlantic coasts of the United States. In addition, resistance of particular oyster strains to H. nelsoni could contribute to the tolerance that Gulf of Mexico oysters demonstrate toward MSX strains endemic to the area. Both artificial and natural selection have resulted in MSX-resistant oyster lines in the mid-Atlantic (Haskin & Ford 1979, Ragone-Calvo et al. 2003). Whereas we detected H. nelsoni rDNA in the oysters, we did not qualitatively confirm the infection intensity in the sampled oysters by histology. Clearly, research is necessary to further resolve the nature of the host-parasite interaction and the degree of resistance that Gulf of Mexico oysters may have against H. nelsoni.

We also identified H. nelsoni infections in oysters originating from Puerto Rico (Crassostrea rhizophorae Guilding) and Venezuela (Crassostrea gasar Adanson and/or C. rhizophorae). We are unable to conclude that H. nelsoni is present in Puerto Rico because the oysters collected there may have been exposed to water from the Atlantic Ocean after their transportation to Harbor Branch Oceanographic Institute (Ft. Pierce, FL). However, we are confident that H. nelsoni is present in Venezuela. Interestingly, these observations suggest that H. nelsoni is a general parasite of oysters but triggers pathogenic states only in C. virginica, a pattern that has been established also for P. marinus (Barber & Mann 1994, Chu 1996, Chu et al. 1996, Calvo et al. 1999; 2001).

The presence of H. nelsoni in the Gulf of Mexico raises several interesting questions: (1) When and where did H. nelsoni establish itself in the Gulf of Mexico? (2) How does MSX, an introduced species, propagate throughout oyster populations of the Gulf of Mexico? and (3) Why are epizootic events restricted to C. virginica populations on the Atlantic coast of North America? The answers to each of these questions await further analysis of host-parasite interactions and population structure in both MSX and oysters.


The authors thank N. Stokes for graciously providing the pMSX414 plasmid as a positive control for H. nelsoni. We also thank W. Fisher, K. Bayha, G. Burreson, J. Perez, and J. Scarpa for providing oyster samples for this study.


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Collection dates and sites for oysters. An asterisk indicates that a
given date is that of provision of a tissue sample from a collector
rather than actual collection date.

 Figure 1
 Collection Reference
 Origin Date Species Number

Apalachicola Bay, FL Jul-94 C. virginica 1
Cedar Key, FL Jul-94 C. virginica 2
Port Aransas, TX Jan-00 C. virginica 3
Tabasco, Mexico Aug-94 * C. virginica 4
Puerto Rico Apr-95 * C. rhizophorae 5
Venezuela Jul-02 C. rhizophorae or gasar 6

P. N. ULRICH, (1,2)* C. M. COLTON, (2) C. A. HOOVER, (2) P. M. GAFFNEY (2) AND A. G. MARSH (2)

(1) University of Georgia, 360 Paul D. Coverdell Center, 500 D.W. Brooks Dr., Athens, GA 30602; (2) College of Marine and Earth Studies, University of Delaware, Lewes, Delaware 19958 USA

* Corresponding author. E-mail:
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Author:Marsh, A.G.
Publication:Journal of Shellfish Research
Date:Apr 1, 2007
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