EXAMINATION OF Merceneria mercenaria AS A HOST FOR Perkinsus marinus By AYANA M. MCCOY A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA 2005
Copyright 2005 by Ayana M. McCoy
This document is dedicated to my parents, Larry and Karen McCoy.
ACKNOWLEDGMENTS First, I would like to thank God for allowing me this opportunity and carrying me through. Next, I would like to thank my family, especially my parents, Larry and Karen McCoy, for their love, patience, and support. I also would like to thank my committee, Drs. Shirley Baker, Anita Wright and Ruth Francis-Floyd, for their hard work, patience and dedication. I want to thank Samara Strauber and Dr. Ramon Little of the Department of Statistics for the statistical analysis of my data. I would like to thank Marqus Johnson for his love and support throughout the years since our undergraduate days at Savannah State University. I would especially like to thank Samesha Barnes for the love, laughter, and support that helped me though the tough times of graduate school. The support, love and friendship of my sorors of the Gainesville Alumnae Chapter of Delta Sigma Theta Sorority, Inc, and members of the UF Black Graduate Student Organization have helped me survive the ups and downs of graduate school. I also appreciate the support, advice, and friendship of the past and present members of the Wright and Baker labs: Mark Campbell, Melissa Evans, Melissa Jones, Maria Chatzidaki-Livanis, Ana Quevedo, Masoumeh Rajabi, Carla Beals, Derk Berquist, Jon Fajans, and David Heuberger. I also would like to thank other people in the Fisheries and Aquatic Sciences and Food Science and Human Nutrition Departments: Michael Wood, Jaime Greenawalt, Stephanie Keller, Kristina Garner, and Joel Carlin. Lastly, I would like to thank Alfreda Cole, LaTarsha Smith, Cleveland Miles, Kevin Holloway, and Max for their love and support. iv
TABLE OF CONTENTS page ACKNOWLEDGMENTS .................................................................................................iv LIST OF TABLES ............................................................................................................vii LIST OF FIGURES .........................................................................................................viii ABSTRACT .......................................................................................................................ix CHAPTER 1 INTRODUCTION........................................................................................................1 2 LITERATURE REVIEW.............................................................................................4 Early Studies.................................................................................................................4 Taxonomy.....................................................................................................................4 Lifecycle of Perkinsus..................................................................................................6 Epizootiology of Perkinsus...........................................................................................7 Culture and Diagnostic Methods..................................................................................8 Rayâ€™s Fluid Thioglycollate Media Assay..............................................................8 Molecular Biotechniques.....................................................................................10 Pathogenesis...............................................................................................................14 Crassostrea virginica..................................................................................................15 Mercenaria mercenaria..............................................................................................16 Hypotheses..................................................................................................................17 Specific Aims..............................................................................................................18 3 MATERIALS AND METHODS...............................................................................19 Environmental Study Area and Sampling Plan .........................................................19 Culture of Perkinsus Species......................................................................................20 Infection Studies.........................................................................................................21 Rayâ€™s Fluid Thioglycollate Media Assay...................................................................22 DNA Extractions........................................................................................................23 Polymerase Chain Reaction Diagnostic Assay...........................................................24 DNA Sequencing........................................................................................................26 Statistics......................................................................................................................26 v
4 RESULTS...................................................................................................................27 Physical Parameters of Sampling Sites.......................................................................27 RFTM Analysis of Oyster Samples............................................................................31 PCR Analysis of Oyster Samples...............................................................................36 Comparison of Infections from Digestive Tract vs. Gill Tissues...............................40 RFTM and PCR Analysis of Clam Samples...............................................................41 Clam Infection Study..................................................................................................42 5 DISCUSSION AND CONCLUSION........................................................................45 Perkinsus marinus Detection in Oysters.....................................................................46 RFTM and PCR analysis of Clam Samples................................................................49 Comparisons of Detection Assays..............................................................................54 Conclusions.................................................................................................................57 LIST OF REFERENCES...................................................................................................59 BIOGRAPHICAL SKETCH.............................................................................................67 vi
LIST OF TABLES Table page 3-1 Mackin scale used to measure infection intensity of Perkinsus in samples.............22 4-1 Monthly means of physical parameter of all sampling sites....................................28 4-2 Site means of physical parameters of all sampling sites..........................................28 4-3 Effect of months on parameters as determined by ANOVA....................................28 4-4 Effect of site on parameters as determined by ANOVA..........................................29 4-5 Monthly and site percentage of positive oyster digestive tract samples by RFTM assay.........................................................................................................................35 4-6 Mackin scale scores of the monthly and site infection intensity of positive oyster digestive tract samples by RFTM assay...................................................................36 vii
LIST OF FIGURES Figure page 2-1 Lifecycle of Perkinsus marinus..................................................................................6 2-2 A representation of the rRNA locus of P. marinus showing the NTS region..........11 3-1 Oyster and clam sampling sites in the Cedar Key area...........................................20 4-1 Physical parameters for the sampling sites..............................................................30 4-2. Monthly means of the percentage of Perkinsus positive oyster samples as determined by RFTM...............................................................................................32 4-3 Mean infection intensity of Perkinsus positive oyster samples as determined by RFTM using the Mackin scale.................................................................................33 4-4 Monthly mean percentage of positive samples from RFTM assay versus P. marinus PCR assay...................................................................................................38 4-5 Comparisons of monthly mean percentages of positive oyster digestive tract and gill tissue samples by genus-specific and P. marinus PCR assays..........................39 4-6 Comparisons of percentage of positive clam digestive tract samples by RFTM, genus-specific PCR, and P. marinus PCR assays....................................................43 4-7 Percentages of positive June 2004 clam gill samples by RFTM, genus-specific PCR, and P. marinus-specific PCR assays..............................................................44 viii
Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science EXAMINATION OF Mercenaria mercenaria AS HOST FOR Perkinsus marinus By Ayana M. McCoy May 2005 Chair: Shirley Baker Cochair: Anita Wright Major Department: Fisheries and Aquatic Sciences Perkinsus marinus is a parasite of the Eastern oyster, Crassostrea virginica, and has caused mass mortalities of oysters along the Atlantic and Gulf coasts. The commercially important hard clam, Mercenaria mercenaria, is aquacultured in areas that are naturally populated by C. virginica. Susceptibility of clams to P. marinus infections, as well as their possible role as reservoir hosts, has not been well studied. Therefore, the objectives for this study were to 1) compare the incidence of P. marinus in M. mercenaria versus C. virginica from a region of high density clam aquaculture on the Gulf Coast of Florida and 2) experimentally test the susceptibility of clams to P. marinus. Shellfish were collected monthly from February through August 2002, September 2003, and June 2004 from Cedar Key, FL. Salinity, temperature, dissolved oxygen and depth were measured for each month and site. Infections were determined by standard parasite staining in Rayâ€™s Fluid Thioglycollate Media (RFTM), which provided semi-quantitative analysis for estimates of infection levels. Parasitism was also examined by polymerase chain reaction (PCR), using both Perkinsus genus-specific and P. marinus species-specific primers. ix
Salinity averaged 26 3.37 ppt.; temperature averaged 26.9 4.36 C; dissolved oxygen averaged 6.6mg/L 1.08; and depth of sampling sites averaged 0.84m 0.51 at low tide. Parasites were not detected in oyster samples by RFTM during winter months of February and March 2002, and mean levels of infection were <20% in April through June 2002. However, most samples (67 to 100%) were positive during the summer months of July and August 2002, September 2003, and June 2004 by this assay. Conversely, species-specific PCR detected positive samples (mean = 57%) in February and March 2002 when RFTM assays had been negative, while 44% of summer oysters that were positive by RFTM were negative by P. marinus-specific PCR. However, 78% of samples were positive by genus-specific PCR, confirming that many of the samples that were negative for P. marinus-specific PCR were in fact positive for Perkinsus parasites (37%). Possible explanations for these results include 1) decreased sensitivity of P. marinus-specific PCR in summer samples due to background interference, 2) infection of oysters by Perkinsus species other than P. marinus, and/or 3) genetic variability in P. marinus populations. P. marinus infections were not detected in any clam samples by species-specific PCR throughout the entire study, and these results were supported by artificial inoculation of clams with P. marinus, which also indicated that clams were not susceptible to infection. Although evidence of parasitic infection in clams by RFTM and/or genus-specific assays was relatively rare (only 6% of all clam samples), detection of this low incidence of Perkinsus spp. in clams supports continued surveillance for future risk assessment of the threat of these parasites to clam aquaculture. x
CHAPTER 1 INTRODUCTION Perkinsus species are endoparasitic protistans that infect more than 60 molluscan species. Most parasitized species are bivalves that inhabit coastal areas worldwide, including both sides of the Atlantic Ocean, the Mediterranean Sea and the South Pacific Ocean (Perkins, 1996). In the United States, P. marinus is found along the Atlantic and Gulf coasts and has been implicated in mortalities of the Eastern oyster, Crassostrea virginica, causing great concern to shellfish farmers. This parasite was originally classified as a fungus, Dermocystidium marinus (Mackin et al.,1950), but it was reclassified as P. marinus and placed in the phylum Apicomplexa (Levine, 1978). Molecular analyses recently placed the genus into a new phylum (Noren et al., 1999) and suggested that Perkinsus is from an early ancestral group of apicomplexans and dinoflagellates (Saldarriaga et al., 2003). Molecular bio-techniques, along with more detailed morphological studies, have also been used to reclassify species within the genus Perkinsus (Kotob et al., 1999; Noren et al., 1999; Perkins, 1996; Siddall et al., 1997). These techniques, including polymerase chain reaction (PCR), have helped scientists identify and distinguish among several Perkinsus spp. that are associated with various molluscan species. Six species are currently included in the genus Perkinsus: P. marinus, P. olseni, P. qugwadi, P. atlanticus, P. andrewsi and most recently, P. mediterreaneus (Mackin et al., 1950, Lester and Davis, 1981; Blackbourn et al., 1998; Azevedo, 1989; Coss et al., 2001, Casas et al., 2004). 1
2 Mortality caused by P. marinus has been primarily reported in oysters, but it is possible that another bivalve, such as a clam, could serve as a reservoir host. The pathogenicity of Perkinsus spp. is not well understood; however, mortalities seem to be most severe under conditions of elevated salinity and temperature. It is unclear whether Perkinsus is transmitted directly from the environment to the mollusk, or if there are alternative hosts that harbor Perkinsus. P. atlanticus and P. atlanticus-like isolates have been found in several clam species, including Ruditapes decussatus, R. philippinarium, R. semidecussatus and Venerupis aureus (Azevedo, 1989). Coss et al. (2001) detected P. andrewsi in the hard clam, Mercenaria mercenaria, collected from the Atlantic coast. Coss et al. also demonstrated that two different Perkinsus species could coexist in sympatric molluscan species. The commercially important hard clam is often found sympatrically with oysters and could potentially be a vector or an intermediate host, serving as a reservoir of the infectious life stages of P. marinus. Few studies have specifically looked for infection of Perkinsus spp. in M. mercenaria (Ray, 1954; Andrews and Hewatt, 1957; Cheng et al., 1995). The objectives of this project were as follows: 1. To survey natural infection of Perkinsus marinus in M. mercenaria versus C. virginica from a region of high density clam aquaculture on the Gulf Coast of Florida by Rayâ€™s Fluid Thioglycollate Media (RFTM) assay and by polymerase chain reaction (PCR). 2. To test the susceptibility of M. mercenaria to infection by cultured P. marinus cells. 3. To survey clams and oysters for other Perkinsus species, using genus-specific PCR. This study will help determine whether M. mercenaria could serve as a reservoir host for P. marinus. It will survey the presence of Perkinsus spp. on Floridaâ€™s Gulf Coast and examine the potential threat to Floridaâ€™s hard clam industry. Detection of M.
3 mercenaria infection by Perkinsus species may lead to consideration of new management strategies in order to prevent or control infections in both oysters and clams. The possibility of M. mercenaria as a reservoir for Perkinsus may necessitate that farm plots be placed at a greater distance from harvested oyster beds in order to decrease the risk of infection and subsequent mortality of the oysters.
CHAPTER 2 LITERATURE REVIEW Early Studies The study of Perkinsus spp. began with a Texas A & M University project, which was funded by several oil companies. The projectâ€™s purpose was to determine the cause of oyster mortalities in Louisiana oil fields. Extensive studies were conducted on the effects of crude oil, bleed water, natural gas, drilling mud, and seismographic surveys on oysters (Mackin and Hopkins, 1961). The conclusion of the study was that none of those pollutants or activities was causing death of oysters in affected areas. A parasite associated with elevated temperatures and salinities was implicated as the cause of widespread oyster mortality (Mackin and Hopkins, 1961). Mackin et al. (1950) had previously described this parasite as Dermocystidium marinum (Dermo) because it looked similar to a freshwater parasitic fungus, Dermocystidium salmonis. However, current taxonomy places the genus in it own phylum, Protalveolata (Cavalier-Smith, 1998). Taxonomy Determining the taxonomy of Perkinsus has proven to be difficult. First, it was classified as Dermocystidium marinum (Mackin et al., 1950). Since its first classification as a fungus, the taxon has been placed in several groups, including the Saprolegniales (Perkins and Menzel, 1967). Mackin and Ray (1966) renamed it Labyrinthomyxa marina and placed the parasite in the fungal group, Labyrinthomyxa, because of the similarity in developmental stages and cytology. The members of this group are parasites of aquatic 4
5 plants (Mackin and Ray, 1966). Perkins and Menzel (1967) were the first to affiliate Perkinsus with the Apicomplexans, basing their conclusions on the presence of more than one limiting membrane, a characteristic of malaria parasites. In 1976, Perkins discovered a pinocytic structure in some developmental stages and compared this to the microspores of Apicomplexans. He then argued for the placement of Perkinsus into the phylum Apicomplexa because of the presence of the subpellicular membrane, the microspore and the conoid. Based on Perkinsâ€™s findings, Levine (1978) established the genus, Perkinsus, and formally placed it into the phylum Apicomplexa. Until recently, there was widespread acceptance of the classification of Perkinsus as an apicomplexan. However, with more advanced molecular and morphological methods, Perkinsus is currently thought to be more closely associated with dinoflagellates than with apicomplexans. Perkins acknowledged the similarities between dinoflagellates and Perkinsus and stated that Perkinsus could be the intermediate phylogenetic group between the Dinoflagellates and the Apicomplexans (Perkins, 1996). He based the terminology used to describe the lifecycle of Perkinsus on this idea. However, more recent phylogenetic data, presented by Siddall et al., (1997), concluded that Perkinsus did not belong to the apicomplexans at all and is more closely related to the dinoflagellates. Since Perkinsus is closely related to both apicomplexans and dinoflagellates, Noren et al. (1999) created a new phylum, Perkinsozoa. In 2003, Saldarriaga et al. indicated that Perkinsus should be included with organisms called Protalveolata (Cavalier-Smith, 1998), which are considered an intermediate group between apicomplexans and dinoflagellates, two of the three major groups of alveolates. They compared protein phylogenies and morphological features that suggested that
6 Perkinsus should be placed in an early ancestral group of the dinoflagellate lineage that appeared before the dinoflagellates and apicomplexans diverged since it has features of both, including a modified apical complex (apicomplexans), as well as flagellum, and cell division (dinoflagellates). Lifecycle of Perkinsus The lifecycle of Perkinsus spp has been studied intensively (Figure 2-1). There are two distinct phases of the lifecycle: 1) the developmental stage within the host as a trophozoite in which the flagellum is lost to become a rounded immature form. This matures to form a large vacuole with a vacuoplast (Figure 2-1A). The cell engages in palintomy, which is defined as rapid asexual reproduction in dinoflagellates (Paynter, 1996), and immature trophozoites free themselves through a tear in the cell wall and ultimately spread throughout the hostâ€™s body. In seawater, 2) zoosporulation occurs in which a mature trophozoite enlarges, losing its vacuoplast (Figure 2-1B). A discharge tube and pore develops in the cell wall. Palintomy occurs again, which results in numerous biflagellate zoospores moving through the discharge tube into the seawater. A B A A B B Figure 2-1. Lifecycle of Perkinsus marinus A) Developmental cycle of P. marinus in C. virginica. B) Zoosporulation of P. marinus in seawater, free of the host. (Perkins, 1996)
7 Volety and Chu (1994) determined that two stages of the lifecycle, the trophozoites and the prezoosporangia (enlarged trophozoites also called hypnospores) are both infective; however, the trophozoites are more infective and are possibly the infectious agents of disease transmission in the field. Based on low condition index and protein levels in infected oysters, Volety and Chu (1994) suggested that prezoosporangia may assert a higher energetic demand on the host than do trophozoites. Epizootiology of Perkinsus Perkinsus marinus has been implicated as a cause of mass mortalities of oysters along the Gulf of Mexico since its discovery in 1950. In 1989, Craig et al. studied the Gulfâ€™s regional variations in P. marinus prevalence and intensity of infection between January and March of 1986. In this study, 39 out of 49 Gulf sites, including 15 Florida sites mostly south and east of Apalachicola Bay, had prevalence above 75%. Most of the 15 Florida sites had prevalence above 95%. The median infection intensity (the level of infection), in the Cedar Key, Florida sites was low ( 1 on a 0-5 scale). Prevalence and infection intensity did not correlate with temperature; however, both significantly correlated with salinity. The study concluded that no site was parasite-free and that the low infection intensity represented the near minimum values since sampling was done in the winter time when infection intensities are typically low. Soniat (1996) described the 45â€“year progress regarding our knowledge of the epizootiology of P. marinus in the Gulf of Mexico. This paper reviewed the distribution as well as the environmental and biological factors that affected the transmission and prevalence of this parasite. He concluded that although Perkinsus was a well-studied parasite, there is still much to learn. Suggestions included a model of the disease process, field and laboratory verification of model responses, and ongoing iterative interactions
8 among model, field and laboratory studies. Subsequent papers have discussed models that were designed to investigate the population dynamics of P. marinus (Brousseau and Baglivo, 2000; Ragone-Calvo et al., 2000), although the areas studied were not in the Gulf of Mexico. Field and laboratory studies have clearly shown that salinity and temperature are two major environmental factors that affect the susceptibility, infectivity and transmission of P. marinus (Soniat, 1985; Soniat and Gauthier, 1989; Fisher et al., 1992; Ragone and Burreson, 1993; Ragone-Calvo and Burreson, 1994; Volety and Chu, 1994; Chu, 1996; Ford et al., 1999). High salinities and high temperature enable P. marinus cells to proliferate, causing increased mortalities of oysters (Fisher et al., 1992). Lower salinities (6-9 ppt) delay but do not always eliminate the risk of infection (Ragone and Burreson, 1993). P. marinus can survive low temperatures characteristic of winter, even temperatures as low as 4C (Ragone-Calvo and Burreson, 1994). Ford et al. (1999) found that after 11 weeks at 15C, oysters were not able to eliminate infections. In contrast to P. marinus, oysters are able to rapidly and completely discharge the parasitic pathogen, Haplosporidium nelsoni, when exposed to salinities of 10 ppt or less (Ford et al., 1999). The fact that oysters cannot completely eliminate P. marinus is important because it explains the persistence of this organism in endemic regions. Culture and Diagnostic Methods Rayâ€™s Fluid Thioglycollate Media Assay Several methods have been developed to study the prevalence and intensity of Perkinsus infections. Ray (1952) devised a technique to culture P. marinus within oyster tissue. This technique consists of incubating oyster cells in Rayâ€™s fluid thioglycollate medium (RFTM) with antibiotics for 5-7 days. The trophozoites, which are the feeding
9 and growing stage of Perkinsus, enlarge and become hypnospores, which are stained blue or black with Lugolâ€™s staining solution. The level of infection, or infection intensity, is based on a scale devised by Mackin (Ray et al, 1953). The scale ranges from 0, meaning no infection to 5, meaning high level of infection. RFTM assay provides an accurate estimate of the infection intensity of P. marinus cells because multiplication of cells does not occur (Fisher and Oliver, 1996). Also, only live cells are able to take up the media and enlarge; therefore, accidental diagnosis of dead parasites is eliminated (Fisher and Oliver, 1996). This method has been used to examine the organelles of Perkinsus as well as the lifecycle. Improvements on the RFTM method have been developed over the years. Choi et al. (1989) described a technique for extracting P. marinus from infected oyster tissues cultured in RFTM. This technique frees hypnospores from oyster tissue and other parasites. With this technique, the number of hypnospores per gram of oyster tissue and provided a semiquantitative RFTM assay using 2M sodium hydroxide. Other techniques include tissue digestion; Gauthier and Fisher (1990) developed a hemolymph assay and Bushek et al. (1994) developed a quantitative body-burden assay using RFTM. Based upon the previous studies, Fisher and Oliver (1996) published a protocol for a quantitative whole-oyster enumeration that has been considered a standard procedure of RFTM for detecting P. marinus. RFTM remains the primary method of detection and enumeration of P. marinus. The enlargement of the hypnospores and the blue-black stain produced by the Lugolâ€™s iodine solution in the RFTM assay is a common feature of all species of Perkinsus, except for P. qugwadi (Blackbourne et al., 1998). Perkinsus atlanticus, the
10 species mostly studied on the east coast of Europe, is routinely diagnosed in the cultured clam, Ruditapes decussates, using the RFTM assay (Almeida et al., 1999). Almeida et al. (1999) developed a quantitative whole-clam RFTM assay similar to the whole-oyster assay developed by Fisher and Oliver (1996). Thus, it should be noted that RFTM is not species-specific, and other assays are needed for specificity. Molecular Biotechniques Although RFTM method is the most commonly used in diagnostic Perkinsus studies, it does not discriminate among species. In vitro culture methods (Gauthier and Vasta, 1993; Gauthier and Vasta, 1995; Kleinschuster and Swink, 1993; Kleinschuster et al., 1994) allowed for DNA extraction which led to the development of molecular biotechniques, such as using polymerase chain reaction (PCR) diagnostic assays to differentiate the species. In 1995, Gauthier and Vasta optimized their culture technique using commercially available media normally used for mammalian research. The sequencing of the ribosomal RNA (rRNA) gene has provided intraspecies, as well as interspecies-specific PCR assays (Casas et al., 2004; Pecher et al., 2004; Coss et al., 2001; Robledo et al., 2000; Kotob et al., 1999; Robledo et al., 1998; Marsh et al., 1995; Fong et al., 1993). As shown in Figure 2-2, the ribosomal RNA (rRNA) operon of P. marinus are arranged in the follow order: 5S subunit gene, intergenic spacer (IGS) [external transcribed spacer (ETS) and nontranscribed spacer (NTS)], small subunit (SSU) gene, internal transcribed spacer 1(ITS1), internal transcribed spacer 2 (ITS2), and large subunit (LSU) gene (Robledo et al., 2002). Fong et al. (1993) reported the sequence for the SSU rRNA for P. marinus; however, there was a 77% sequence identity between the P. marinus SSU rRNA and C. virginica SSU rRNA. Greater sequence divergence between host and parasite was needed for the ideal target domain for PCR-based
11 diagnostic assays. Therefore, the non-coding regions, such as the NTS and ITS, were examined (Marsh et al., 1995). The high variability of the NTS sequence makes it an excellent target for detection of the differe nces among intraspecific strains or types and the detection of differences between species (Marsh et al., 1995, Coss et al., 2001). The ITS sequences flanking the rRNA genes has also b een used as a reliable characteristic to distinguish between species from diff erent geographical locations (Goggin, 1994). Figure 2-2. A representati on of the rRNA locus of P. marinus showing the NTS region where (a) the P. marinus primers (Marsh et al., 1995) and (b) Perkinsus -genus specific primers (Robledo et al. , 2001) are located, a nd (c) the universal eukaryotic primers (Medlin et al., 1988). Because of the sensitivity of PCR, light in fections as well as latent infection in over-wintering populations, oyster se ed stocks, and even larvae and spat, can be detected. Coss et al. (2001) used PCR to identify and describe P. andrewsi, and Cases et al. (2004) developed a PCR assay for identification of P. mediterraneus . Other types of PCR used to study this parasite include multiple x PCR (MPCR),which was optimized for simultaneous detection of low levels of infection of both Perkinsus and another oyster parasite, Haplosporidium nelsoni (Penna et al. , 2001). This method wa s developed to be sensitive and species-specific as well as fast and cost-effective. The primers for P. marinus were adapted from Marsh et al., 1995, which were derived from the NTS sequence. Another PCR development was the quantitative competitive PCR (QCPCR), whereby the target DNA competes with a d ilution series of known concentrations of competitor DNA (Yarnell et al. , 2000). Primers for the assay were derived from published P. marinus DNA sequences for the rRNA gene of Fong et al. (1993) and the
12 adjacent ITS1 region published by Goggin (1994) and amplified a 1, 210 bp-fragment from within the SSU to within the ITS1. The quantitation of the unknown target DNA is based upon the equivalence point at a known concentration of competitor DNA. Although Yarnell et al. (2000) reported the successful quantitation of P. marinus DNA in oyster tissue and hemolymph, there were several disadvantages: the use of expensive DNA polymerase is recognized, and the labor intensive process of performing multiple reactions per sample requires expertise to successfully perform this method. Another development for diagnosis of P. marinus and P. atlanticus is a PCR-enzyme-linked immunosorbent assay (ELISA) (Elandalloussi et al., 2004). The assay is performed using a commercially available PCR ELISA kit that incorporates a PCR â€“digoxigenin (DIG) labeling mix in the PCR assay so that the newly synthesized DNA has DIG incorporated into the sequence. The DIG â€“labeled amplified products are hybridized with biotin end-labeled oglionucleotides that bind and capture the product to streptavidin-coated microtiter plates. These products are detected by standard ELISA procedures using an anti-DIG peroxydase-labeled conjugate antibody and a peroxydase substrate that permit a colorimetric reaction dependent on the presence and amount of amplified product. The objective of Elandallousi et al. â€˜s study was to develop a new assay that was species-specific for multiple species within a host, sensitive to low levels of infection, and less labor-intensive and faster then current methods. This assay was compared to standard and multiplex PCR using Perkinsus genus primers, as well as species-specific primers for P. marinus and P. atlanticus. Analysis of the amplified products using this assay was compared with the Southern blot hybridization and the standard ethidium bromide (EtBr)-stained 1% agarose gel. The southern blot hybridization was
13 approximately 100 times more sensitive than the EtBr -stained agarose gels. A linear correlation between the amount of P. atlanticus and P. marinus DNA and the ELISA signal was observed (r= 0.8407 and 0.9139, respectively). Elandallousi et al. also compared the RFTM assay to the PCR-ELISA assay. RFTM and PCR-ELISA assays provided similar results in 39 of 45 samples (87%) with twenty-one samples tested negative and 18 samples were positive by both assays. One sample was positive by RFTM but negative by PCR-ELISA and five samples were negative by RFTM but positive by PCR-ELISA suggesting greater sensitivity of PCR compared to RFTM. The most recent development in molecular methods was a real-time PCR assay designed to detect low levels of P. marinus in environmental water samples (Audemard et al., 2004). Real-time PCR assay measures the exponential production of PCR amplicons as the copies of DNA are being made, thereby providing a relatively rapid and quantitative detection method. The quantification during the logarithmic phase is more indicative of the initial target concentration than quantification during the plateau phase, which is analyzed by standard PCR (Audemard et al., 2004). The objective of Audemard et al.â€™s study was to develop a real-time PCR assay to detect and quantify P. marinus in environmental water samples since few data have been collected on the quantity of Perkinsus isolates in environmental water samples. DNA extraction methods were also optimized with two concerns: 1) DNA recovery from water samples and 2) limiting the effect or presence of PCR inhibitors. The real-time PCR assay was able to detect levels of DNA concentrations as low as the equivalent of 3.3 X 10 -2 cells per 10l of reaction mixture. Results suggested that real-time PCR assay can be used for determining the relative abundance of P. marinus cells in the environment. However, the absolute
14 abundance of the parasitic cells may not be accurately measured, and interpretation of the published environmental abundance data is questionable. The P. marinus cells used to develop the standard curve were cultured cells and could potentially differ from cells found in water samples in the field due to genetic variation in the environmental populations (Audemard et al., 2004). Pathogenesis Paynter (1996) studied the physiological effects of P. marinus on C. virginica. He concluded that P. marinus reduced the hemolymph pH causing acidosis, which affected nutrient absorption, respiration and excretion of waste, resulting in decreased growth rate. Acidosis could inhibit calcification and shell deposition, accounting for the cessation of shell growth associated with infection. Heavy infection may also have a harmful effect on reproduction, but oysters can redirect energy from growth to gametogenesis to minimize the effects of infection on egg quality (Paynter, 1996). La Peyre (1996) has shown that P. marinus cells produce extracelluar proteases, which are thought to play a role in damaging host tissue by protecting the parasitic cells from the hostâ€™s immune system. Host genetic variability may determine differences in susceptibility and tolerance to P. marinus. Bushek and Allen (1996) also suggested that different races of the parasite have different levels of virulence. In their review, they described several studies that suggested persistence of the parasite in low salinity areas, tolerance of low temperatures, the rate of infection spreading from area to area, and significant enlargement of different isolates in RFTM may denote different races of P. marinus. In another study, Bushek and Allen (1996) inoculated over 500 uninfected oysters with 4 different isolates of P. marinus. The results suggested that there are differences in virulence among isolates of P. marinus.
15 Crassostrea virginica Gmelin first identified Crassostrea virginica, the eastern oyster, in 1791. Its natural range is the western Atlantic coast from the Gulf of St. Lawrence in Canada to the Gulf of Mexico, Caribbean, and the coasts of Brazil and Argentina. It was introduced to the west coast of North America, Hawaii, Australia, England, and Japan. It is common in estuaries and coastal areas with lower salinities. Oysters play a functional role in the environment, providing hard substrate for a multitude of invertebrate macrofauna and serving as biological filters for coastal areas. Oyster beds are diverse habitats where small fish, crabs, worms and other invertebrates can be found. They also serve as bioindicators of the environment since they have the capability to bioaccumulate organic and non-organic contaminates. Because of high recruitment rates, oyster populations have never been in danger of total depletion; however, over-harvesting, other anthropogenic factors, and disease have affected the oyster fishery industry. Because of its widespread distribution and persistence in lower salinity areas of the Chesapeake Bay, Perkinsus has been the most studied oyster pathogen in this area since 1987 (Burreson and Ragone-Calvo, 1996). The Chesapeake Bay was once the greatest oyster producing area in the world. In the late 1880â€™s, this area produced 615,000 tons of oysters, which is approximately 20% of the current worldwide production (Goulletquer et al., 1994a), 39% of the United Statesâ€™ oyster harvest, 17% of all United Statesâ€™ fisheries and employed 20% of all Americans in the fishery industry (Kennedy and Breisch, 1983). Now, the average oyster population density in the bay is estimated to be 4% of what it was in the 1880â€™s (Rothschild et al., 1994). By 1959, the Eastern oyster industry was declining due to fishing pressure, reduced water quality, habitat destruction and diseases caused by H. nelsoni and P. marinus. The parasites were the main cause of the
16 devastating decline in the oyster population in the 1970s and 1980s. Current efforts to restore the oyster industry and the Chesapeake Bay as a whole includes the Chesapeake 2000 agreement signed by Maryland, Virginia, Washington DC, Pennsylvania, Chesapeake Bay Commission and the United States Environmental Protection Agency. This agreement was a comprehensive watershed restoration plan for the Chesapeake Bay that focused on protecting and restoring living resources and habitats, improving water quality, managing land, and engaging the community. In Florida, the eastern oyster is cultured on over 500 acres of state owned submerged land leases as constructed and maintained oyster reef (Florida Aquaculture Plan 2003-2004, Florida Department of Agriculture and Consumer Services). The oyster fishery, along with other aquatic animals such as crawfish, eels, snails, turtles, crabs, and frogs, produced less than 1% ($799,000) of all Florida aquaculture sales in 2001 (Florida Agricultural Statistics Service, 2002). Mercenaria mercenaria Hard clams, Mercenaria mercenaria, are marine bivalves that live in coastal areas. Natural distribution of hard clams included the Atlantic coast from Nova Scotia to Florida. They have been introduced in the Gulf of Mexico from Florida to Yucatan, Mexico. The hard clam is the largest and most valuable of the clam species (Lorio and Malone, 1995). They support major commercial fisheries all along the New England and Mid-Atlantic coasts. In Florida, clams are aquacultured in both the Atlantic and Gulf of Mexico coastal areas. The state of Florida provides leases of submerged bottom land in suitable harvesting water where other resources such as sea grasses, reefs and natural harvesting beds are absent (www.hboi.edu/aqua/pdfs/clam.pdf). These areas are called High Density Lease Areas (HDLAs). Farmers buy seed clams and place them into 16
17 squarefoot bags that are placed on the bottom of these leases, with the natural sediments serving as substrate. Individual sites may vary in productivity because of available food, water flow, predators and the bottom type, which can all have an effect on growth rate and survival. The clam leases are easiest to work in shallow water where low tides allow for tending the bags of clams. The hard clam industry has become very important in Florida over the last 20 years; growing from the interest of a few farmers to the preeminent source of U.S. farm-raised hard clams (Florida Aquaculture Plan 2003-2004, Florida Department of Agriculture and Consumer Services). In 2001, cultured clams had an economic impact in Florida of $34 million (Florida Agricultural Statistics Service, 2002) and that year, clams ranked third highest in value of Florida aquaculture sales at 18.4%. Economic output made an additional $17 million in state income and $12 million in valueâ€“added activity in 2001 (Florida Agricultural Statistics Service, 2002). However, sales of clam, clam seed and oysters decreased to $13 million in 2003, from $18.3 million in 2001 (Florida Agricultural Statistics Service, 2004). The demand for clams has been lower than previous years and some producers were impacted by recent poor weather conditions, such as hurricanes. Despite the decrease in sales, the hard clam industry is still the third most lucrative aquaculture industry for the state of Florida. Hypotheses The hypotheses of this present study are 1) Mercenaria mercenaria, the hard clam aquacultured in the Cedar Key area of the Gulf of Mexico is a reservoir host for Perkinsus marinus, and 2) Multiple species of Perkinsus are found in C. virginica and M. mercenaria in the Cedar Key area of the Gulf of Mexico.
18 Specific Aims 1. Survey natural infection of P. marinus in clam high density lease areas and naturally populated oyster reefs in the Cedar Key area of the Gulf of Mexico by Rayâ€™s Fluid Thioglycollate Media (RFTM) assay and by polymerase chain reaction (PCR). 2. Determine the susceptibility of M. mercenaria to infection by cultured P. marinus cells. 3. Survey clams and oysters for other Perkinsus species, using genus-specific PCR.
CHAPTER 3 MATERIALS AND METHODS Environmental Study Area and Sampling Plan The study area consisted of six sampling sites in the Cedar Key, Florida area. These sites included three naturally populated oyster reefs (Bad Luck Reef, Corrigan Reef and Derrick Key Gap) and three high density sites (Gulf Jackson, Pelicanâ€™s Reef and Dog Island high-density lease areas (HDLAs), supporting intensive clam farming (Figure 3-1). Clams and oysters (n=12 each) were collected monthly from the specified sites from February to August 2002, and then in September 2003 and June 2004. Oysters were taken directly from the reefs (Bad Luck, Corrigan and Derrick Key Gap). Clams were collected from individual clam farmers at their leased areas on the specified sites. Salinity, temperature, dissolved oxygen and depth were measured at each oyster and clam sampling site using a multi-parameter water quality monitor (Model 600XLM, YSI Incorporation, Yellow Springs, OH). The average salinity of all sites was 26.2ppt 3.37; water temperatures averaged 26.9C 4.36; the average dissolved oxygen content was 6.64mg/L 1.08 and the average depth was 0.84m 0.51. 19
20 Figure 3-1. Oyster and clam sampling sites in the Cedar Key area. Oyster sites included 1) Derrickâ€™s Gap, 2) Bad Luck Bar, and 3) Corrigan Reef. Clam sites included 4) Gulf Jackson HDLA, 5) Pelican Reef HDLA, and 6) Dog Island HDLA. Culture of Perkinsus Species Cultures of Perkinsus marinus were obtained from Dr. Julie Gauthier at Nichols State University, Louisiana. Cultures were maintained in the laboratory and incubated using the optimized methods of Gauthier and Vasta (1995) for DNA extraction and use in infection studies. Reagents were purchased from Fisher Scientific (Pittsburgh, PA) unless otherwise stated. Briefly, the Perkinsus cells were incubated at 28C in a broth media, which consisted of Dulbeccoâ€™s Media E/Hamâ€™s F12 (Gibco, Invitrogen Corporation, Carlsbad, California) dissolved in 20 ppt artificial seawater (ASW, INSTANT OCEAN,
21 Aquarium Systems, Mentor, Ohio). Antibiotics (100 units of penicillin G/streptomycin sulfate), 50 mM of Hepes (Gibco, Invitrogen Corporation, Carlsbad, California), 3.7 mM of sodium carbonate and sterile 1% fetal bovine serum (Sigma-Aldrich, St. Louis, Missouri) were added to ASW. Five milligrams of the original Perkinsus cells were inoculated into 50 ml of media and incubated for 5 days before 5ml were transferred to new media. The remaining cells were placed in a -70 C freezer with the broth media for future use. Infection Studies To test the susceptibility of M. mercenaria to P. marinus, laboratory infection trials were performed. Clams were collected from HDLAs in Cedar Key, Florida and placed in two open trays enclosed within wooded frames with a closed circulating system at 20-23 ppt salinity and 22-23C. Each treatment had a separate circulating system in order to prevent cross-contamination. After a 1-week acclimation to laboratory conditions, holes were drilled in shells (n=200), and parasites were injected into the body cavity at either high (10 5 ml -1 ) or low (10 2 ml -1 ) concentrations. Control clams received injections of artificial seawater. The holes were closed with water resistant non-toxic putty purchased at a local aquarium store. The clams were fed 2% of the estimated dry body weight of Diet C (Coast Seafoods Company, Quilcene, WA) daily. Mortality was determined by gaping and was monitored daily, and 16 clams from each treatment were collected every 2 weeks for 2 months to determine level of infection and intensity using Rayâ€™s Fluid Thioglycollate Media (RFTM) method.
22 Rayâ€™s Fluid Thioglycollate Media Assay Rayâ€™s Fluid Thioglycollate Media (RFTM) assay was used to detect Perkinsus species in the samples (Figure 3-2). The oysters and clams were opened with an oyster knife or clam opener, respectively. Each animal was rinsed with sterilized ASW (35 ppt), followed by 10% bleach solution and rinsed again with ASW. Approximately 100 mg of tissue from the digestive tract, mantle, or gill were placed into separate wells filled with 1 ml of RFTM on a 48-well plate. Three wells within a single plate were allocated for each sample; there was a single plate for each site. The plates were incubated at room temperature inside a cabinet for 5-7 days. Afterwards, each piece of tissue was stained with Lugolâ€™s iodine solution and examined for Perkinsus at 100X with immersion oil. Infection intensity was based on the Mackin (Ray et al., 1953) scale, and ranged from 0 indicating no infection to 5 indicating heavy infection, as indicated in Table 3-1. 1 2 1 2 4 3 Figure 3-2. RFTM experiment set up: 1)12 samples, oysters or clams;2) 48 well plate with 1ml of FTM in wells;3) Fluid Thioglycollate Media;4) Sample tubes Table 3-1. Mackin scale used to measure infection intensity of Perkinsus in samples Category Definition Score Negative No blue staining sporocysts visible 0 Light A few definite but isolated cells or clusters of cells visible 1
23 Table 3-1. Continued Category Definition Score Light/Moderate Parasites occupy about 25% of tissue preparation 2 Moderate Parasites occupy about 50% of tissue preparation 3 Moderate/Heavy Parasites occupy about 75% tissue preparation 4 Heavy Parasites occupy about 100% tissue preparation 5 (Ray et al., 1953) DNA Extractions DNA was extracted from the digestive tract of samples collected from February though August 2002 using the Qiagen DNeasy tissue kit (Qiagen, California). DNA was extracted from the gills from February 2002 and March 2002 samples and both gills and digestive tract from September 2003 and June 2004 samples using the QIAamp Kit (Qiagen, California). These kits are identical, but the names are different for clinical validation purposes (Qiagen, personal communication). Samples extracted with the DNeasy Kit used 25mg of tissue, which was lysed with 180l Buffer ATL and 20 l Proteinase K for 24 hours. DNA was bound to a column with 200l Buffer AL and 200l EtOH and washed twice with 500 ml ethanol buffers, AW1 and AW2. Finally, the DNA was eluted with 200 ml of AE buffer. Samples extracted with the QIAamp Kit used 60 mg of tissue, which was lysed with 180 l Buffer ATL and 48 l of Proteinase K for 24 hours. Samples were treated with 8l of RNase A and incubated at room temperature for 2 minutes. DNA was bound to a column with 200l Buffer AL and 480l EtOH and washed twice with 500 ml ethanol buffers, AW1 and AW2. Finally, the DNA was eluted with 100 ml of AE buffer. All DNA extracts were ethanol precipitated, and DNA yield
24 and purity were determined using a spectrophotometer (SPECTRA max , Plus 384, Molecular Devices, Sunnyvale, CA) at absorbance 260 and 280 nm. Polymerase Chain Reaction Diagnostic Assay The polymerase chain reaction (PCR) assay was used to determine the presence of Perkinsus infection in oysters and clams. Perkinsus genus-specific (Robledo et al., 2002) and P. marinus species-specific primers (Marsh et al., 1995) were used to determine infection in C. virginica and M. mercenaria. Both sets of primers were derived from the NTS region of the rRNA operon. Primers were obtained from Sigma-Genosys Laboratories (The Woodlands, TX). Each PCR mixture contained autoclaved water, 100mM each of dATP, dCTP, dGTP, dTTP (Eppendorf Scientific, Hamburg, Germany), 10x Taq polymerase buffer with Mg 2+ (500mM KCl, 100mM Tris-HCl pH 8.3 at 25C, 15mM Mg (OAc) 2 ) (Eppendorf Scientific, Hamburg, Germany), 100nM each of forward and reverse primers and 1.25U of Taq DNA polymerase (Eppendorf Scientific, Hamburg, Germany), and 1 g of DNA template, in a total volume of 25 l for each DNA sample. PCR reactions for the monthly samples of April through August 2002, September 2003 and June 2004 included HotMaster Taq DNA polymerase (Eppendorf, Hamburg, Germany) instead of the Taq polymerase previously mentioned. DNA extracted from cultured parasites was used as the positive control, while Vibrio vulnificus DNA and distilled water (no template) were negative controls. Once the sample DNA was amplified using a Mastercycler Gradient thermocycler (Eppendorf Scientific, Hamburg, Germany), the sample was visualized in a 1% agarose gel stained with ethidium bromide for electrophoresis at 80-85 volts.
25 The universal rRNA primers, ss3 and ss5, were used to confirm the stability of the DNA template (Medlin et al., 1988). The expected fragment size was 1792 base pairs. The thermal parameters for this assay were: 93C was held as samples were placed in the thermocycler, followed by 30 cycles of 93C for 1:30 min, 58C for 1:30 min, 72C for 2:00 min and then a final extension time of 10 min at 72C. Sequences for primers PER1 and PER2 were designed to amplify a 319base pair target which is a partial sequence of the intergenic spacer (IGS) in P. marinus, P. atlanticus, and P. andrewsi, making this set of primers â€œgenus-specificâ€(Robledo et al., 2002). The thermal parameters for this assay were: Hold at 94 as samples were placed, followed by 94C for 4 min, then 35 cycles of 91C for 1 min, 60C for 1 min, 72 C for 1 min and then a final extension time of 7 min at 72 C. The primers used for the P. marinusâ€“specific assay were PM300F and PM300R (Marsh et al., 1995). These primers amplify a 307-base pair target region within the non-transcribed spacer (NTS) of the rRNA operon from P. marinus. The samples were processed as follows: Hold at 93 as samples were placed, followed by 3 min at 93C, 35 cycles of 93C for 1 min, 60C for 1 min, and 72C for 1 min with a final extension at 72C for 10 min. This process was adapted from Robledo et al. (1998), in which these primers were tested to determine their sensitivity and speciesspecificity in comparison to the fluid thioglycollate assay. A temperature gradient experiment was performed in order to optimize this PCR assay for our thermocycler, raising the annealing temperature to 60C from 55C.
26 DNA Sequencing To confirm the results of the PCR assays, selected samples were sent to the Interdisciplinary Center for Biotechnology Research (ICBR) at the University of Florida, Gainesville, FL to be sequenced. To prepare amplified samples for sequencing, the GENECLEAN kit (Qbiogene, Carlsbad, CA) was used. Three volumes of sodium iodide (NaI) were added to one volume of the amplified DNA. Then the DNA was vortexed to bind to GLASSMILK, an aqueous solution of silica matrix (Qbiogen, Carlsbad, CA). This pellet was washed 3 times with an ethanol-based solution, NEW wash (Qbiogene, Carlsbad, CA). After the washes, the DNA was eluted with sterilized water. DNA samples (6l) were placed in a 1% agarose gel to determine DNA concentration. Sequences were confirmed by comparison to published sequences using an online program, BLAST (http://www.ncbi.nlm.nih.gov/BLAST/) to determine identity. Statistics Significant differences between months and sites were determined by analysis of variance (ANOVA) at a level of significance of p<0.01. If ANOVA analysis was significant, then the Tukey test was used to determine significant difference between individual data points of months or sites at a level of significance of p < 0.01. Both statistical analyses were performed with the SAS program. Statistical analyses were done by Samara Strauber and Dr. Ramon Little from the Department of Statistics, University of Florida.
CHAPTER 4 RESULTS Physical Parameters of Sampling Sites As both the incidence of shellfish disease and the prevalence of Perkinsus spp. have been shown to vary with environmental conditions, various physical parameters were observed at six sampling sites. Salinity, temperature, dissolved oxygen and depth were recorded monthly at each site, as described in the Materials and Methods section. Three of these sites (Corrigan Reef, Bad Luck Bar and, Derrick Key Gap) were natural oyster beds, and three were sites for commercial clam aquaculture (Dog, Island, Pelican Reef, and Gulf Jackson), as described in Materials and Methods. Mean depths of the various sampling sites ranged from 0.4m to 1.4m and averaged 0.84m 0.51 (Tables 4-1, 4-2), but there were no significant differences among the monthly depths for each site (Table 4-3). However, the mean depth of one clam site (Pelican Reef) was significantly greater (p < 0.01) than all three oyster sites (Table 4-4). Dog Island, another clam site, was significantly (p < 0.01) deeper than Bad Luck Bar and Derrick Key Gap. There were no significant differences among the mean depths of the clam leases or among the mean depths of the oyster sites. Mean temperatures ranged from 17.5C to 32.9C and increased throughout the spring but began to decline in August (Figure 4-1, Tables 4-1, 4-2). The highest mean temperature was observed in July 2002, which was significantly (p < 0.01) higher than all other monthly mean temperatures (Table 4-3). The lowest mean temperature was 27
28 recorded in February 2002, which was also significantly (p < 0.01) lower than all other monthly mean temperatures. Table 4-1. Monthly means of physical parameter of all sampling sites Months Salinity Temperature Dissolved Oxygen Depth February 2002 28.37 0.58 17.49 0.23 7.30 0.71 0.8 0.46 March 2002 25.32 2.09 23.01 0.53 6.86 0.44 1.3 0.84 April 2002 25. 48 2.35 26.31 0.31 5.73 1.32 0.8 .39 May 2002 27. 98 2.55 27.70 0.55 5.85 0.48 0.8 0.52 June 2002 30.83 2.02 30.55 0.61 8.13 1.08 0.9 0.62 July 2002 27.34 1.37 32.86 1.27 6.95 1.01 0.7 0.27 August 2002 22.48 2.04 28.97 0.52 5.86 0.66 0.7 0.36 September 2003 22.42 0.93 27.41 0.41 6.48 0.33 0.6 0.35 June 2004 23.27 3.45 29.06 0.29 5.93 0.55 0.9 0.49 Table 4-2. Site means of physical parameters of all sampling sites Sites Salinity Temperature Dissolved Oxygen Depth Corrigan Reef 26.98 4.09 27.17 4.59 6.19 1.33 0.6 0.3 Bad Luck Bar 25.66 3.00 27.48 4.79 7.05 1.07 0.5 0.48 Derrick Key Gap 23.54 2.97 27.06 4.55 6.79 0.98 0.4 0.17 Gulf Jackson 26 2.97 26.63 4.28 5.99 0.60 1.0 0.16 Dog Island 27.53 4.06 26.23 4.69 6.44 1.53 1.2 0.61 Pelican Reef 26.94 2.26 26.51 4.56 6.45 0.65 1.4 0.33 Table 4-3. Effect of months on parameters as determined by ANOVA Variable Pairs of significantly different months Salinity (df=8; F=16.95; p<0.01) June 2002 is higher than March, April, August 2002, September 2003, and June 2004 February 2002 is higher than August 2002, September 2003, and June 2004 May 2002 is higher August 2002, September 2003, and June 2004 July 2002 is higher August 2002, September 2003, and June 2004 Temperature (df=8; F=415.82; p<0.01 July 2002 is higher than all other months February 2002 is lower than all other months March 2002 is lower than April to August 2002, September 2003 and June 2004 but higher than February 2002
29 Table 4-3. Continued Variable Pairs of significantly different months Temperature (df=8; F=415.82; p<0.01) April 2002 is higher than February and March 2002 but lower than May to August 2002, June 2004 May 2002 is higher than February to April of 2002 but lower than June to August 2002 and June 2004 June 2002 is lower than July 2002 but higher than all other months August 2002 is higher than February to May 2002 and September 2003 but lower than June and July 2002 September 2003 is higher than February and March 2002 but lower than June to August 2002, and June 2004 June 2004 is higher than February to May 2002 and September 2003 but lower than June and July 2002 Dissolved Oxygen (df=8; F=6.65; p<0.01) June 2002 is higher than April, May, August 2002, September 2003, and June 2004 February 2002 is higher than April 2002 Depth (df=8; F=1.81; p<0.01) Not Significant a) For ANOVA, df = degrees of freedom, F= F value, p = significance value Table 4-4. Effect of site on parameters as determined by ANOVA Variable Pairs of significantly different months Salinity (df=5; F=5.45; p<0.01) Derrick Key Gap is lower than Dog Island, Corrigan Reef and Pelican Reef Temperature df=5; F=2.90; p<0.01) Bad Luck Bar is higher than Pelican Reef and Dog Island Dissolved Oxygen df=5; F=1.89; p<0.01) Not Significant Depth df=5; F=11.23; p<0.01) Pelican Reef is higher than Corrigan Reef, Bad Luck Bar, and Derrick Key Gap Dog Island is higher than Corrigan Reef, Bad Luck Bar and Derrick Key Gap Gulf Jackson is higher than Derrick Key Gap a) For ANOVA, df = degrees of freedom, F= F value, p = significance value Significant differences were also noted among other months, and these differences are summarized in Table 4-3. Bad Luck Bar was the only oyster site at which temperature was significantly (p < 0.01) higher than two clam sites (Pelican Reef and Dog Island (Table 4-4). There were no significant differences among the mean temperatures of clam sites or among the oyster sites.
30 Dissolved oxygen fluctuated throughout the sampling period, averaging 6.6 mg/L 1.08 and ranging from 5.7 to 8.13 mg/L (Tables 4-1, 4-2). June 2002, in which the highest dissolved oxygen level was observed, was significantly (p < 0.01) higher than April, May, and August 2002, September 2003, and June 2004. The lowest dissolved oxygen level was in April 2002 and was only significantly lower than June 2002 (Tables 4-1, 4-3). All other monthly mean dissolved oxygen levels were not significantly different from each other. Bad Luck Bar and Gulf Jackson had the highest and lowest mean dissolved oxygen levels, respectively; however, no significant differences were observed among the mean dissolved oxygen levels of each site (Tables 4-1, 4-2). Figure 4-1. Physical parameters for the sampling sites. Salinity (A), temperature (B) and dissolved oxygen (C) are from all six sampling sites in Cedar Key, FL, from February through August 2002 and September 2003 and June 2004. The oyster sites are Corrigan Reef, Bad Luck Bar, and Derrick Key Gap. The clam sites are Gulf Jackson, Dog Island, and Pelican Reef.
31 0.05.010.015.020.025.030.035.040.0February 2002March 2002April 2002May 2002June 2002July 2002August 2002September 2003June 2004Temperature (C)B 0.02.04.06.08.010.0February 2002March 2002April 2002May 2002June 2002July 2002August 2002September 2003June 2004MonthDissolved Oxygen (mg/L) Corrigan Reef Bad Luck Bar Derrick Key Gap Gulf Jackson Dog Island Pelican Reef O y ster Sites:Clam Sites:C Figure 4-1. Continued RFTM Analysis of Oyster Samples In order to determine if the hard clam, Mercenaria mercenaria, is a possible reservoir host for Perkinsus species, parasite loads of clams and oysters from Cedar Key, FL were compared by two independent methods: Rayâ€™s Fluid Thioglycollate Media (RFTM) assay and Polymerase Chain Reaction (PCR) assay. RFTM analysis was used to
32 evaluate both the percentage of positive samples, as well as the infection intensity of Perkinsus spp. as determined by the Mackin scale. Tissue from the digestive tract was examined by the RFTM and speciesâ€“specific PCR assay for all nine months of the study. Gill tissues were examined by genus-specific and species-specific PCR assays for four months (February, March, 2002, September 2003 and June 2004). RFTM was not used to examine gill tissues. The percentage of Perkinsus-positive oysters exhibited seasonality and increased dramatically from winter to summer as determined by RFTM of digestive tract tissues (Figure 4-2). During the months of February and March of 2002, no infections were detected by RFTM in oyster samples from any site. 0%20%40%60%80%100%February 2002March 2002April 2002May 2002June 2002July 2002August 2002September 2003June 2004MonthsPercent Positive0.05.010.015.020.025.030.035.0Salinity (ppt) and Temperature (C ) Monthly mean Salinity Temperature Figure 4-2. Monthly means of the percentage of Perkinsus positive oyster samples as determined by RFTM. Monthly samples (n=36) were collected from three oyster sites. The percent of positive samples was examined by RFTM. July, August 2002, September 2003 and June 2004 were significantly different from February â€“June 2002 samples (p < 0.01).
33 The months with the highest mean percentages of positive oyster samples were July and August of 2002, September 2003, and June 2004 with 86%, 86%, 100%, 78%, respectively. The mean percentages of positive samples of these months were significantly higher (p < 0.01) than the mean percentages of other months (February to June of 2002). As shown in Figure 4-3, infection intensity, as measured by the relative numbers of parasites in infected tissue based on the Mackin scale, corresponded well with the percentage of infected oysters, r 2 = 0.94 (p < 0.01). The mean infection intensities of the summer months were also significantly (p < 0.01) higher than the mean infection intensities of the winter and spring months. Individually, the mean infection intensity of September 2003 was significantly (p < 0.01) higher than the means from all the other months. 012345February 2002March 2002April 2002May 2002June 2002July 2002August 2002September 2003June 2004MonthsMackin Score0.05.010.015.020.025.030.035.0Salinity (ppt) andTemperature ( C ) Monthly Mean Salinity Temperature Figure 4-3. Mean infection intensity of Perkinsus positive oyster samples as determined by RFTM using the Mackin scale. Monthly samples (n=36) were collected from three oyster sites. Each point is a mean Mackin score from each oyster site for each month. The infection intensity is measured by the Mackin scale which ranked from 0-5, 0 as uninfected to 5 as heavy infection.
34 Seasonality of P. marinus infections in oysters may be related to differences in the physical parameters that were observed in winter (February and March 2002) compared to summer months (July and August 2002, September 2003 and June 2004). For example, February and March were significantly lower in temperature than all other months. As temperature increased from February to July 2002, increases were observed in both the percentage of positive samples and in infection intensities. Although August mean temperature decreased, this shift was apparently not sufficient to affect infections of Perkinsus in oysters. These results were also confirmed by observations of high levels of infections in September 2003 and June 2004. In general, summer months with high levels (<70%) of infection (July and August 2002, September 2003, and June 2004) also exhibited significantly (p < 0.01) lower average salinity (23ppt) compared to other months (27ppt).On the other hand, the month with the highest mean salinity (30.8ppt in June 2002) showed less than 17% infection, which was significantly less (p < 0.01) than the RFTM positive samples that were detected in samples collected at lower salinities, 22.5-27.3ppt (July and August 2002, September 2003 and June 2004). Further, the mean salinity of February 2002 was significantly higher than September 2003 (p < 0.01) when corresponding infection levels were 0% (no detection) and 100%, respectively. Finally, dissolved oxygen did not appear to affect the number of P. marinus infected oyster samples, as the month with highest dissolved oxygen (June 2002) showed less than 20% infection, and one of the months with lowest oxygen (August 2002) had an infection level greater than 80%. There was no significant difference in mean dissolved oxygen among the oyster sites.
35 Examination of differences among sampling sites over the course of this study did not reveal any significant differences in the levels of Perkinsus spp. infections, as determined by RFTM analysis of oysters; and high levels (>65%) of RFTM positive samples were detected from all three sites during the months of July and August 2002 and in September 2003 and June 2004 (Table 4 -5). However, differences among sites were noted during individual months. For example, during May 2002, only one site (Bad Luck Bar) was positive for Perkinsus spp. infections by RFTM, while, in June 2002, Bad Luck Bar was the only site where infections were not detected. At the oyster sites, there was a significant difference between the mean salinities of Derrick Key Gap and Corrigan Reef (p < 0.01); however, neither site was significantly different in the level of P. marinus infections compared to Bad Luck Bar. Table 4-5. Monthly and site percentage of positive oyster digestive tract samples by RFTM assay (n=12 oysters per site per sampling month) Oyster Sites Month Derrick Key Gap Bad Luck Bar Corrigan Reef Monthly mean February 2002 0% 0% 0% 0% March 2002 0% 0% 0% 0% April 2002 33% 25% 8% 22% 0.13 May 2002 0% 8% 0% 3% 0.05 June 2002 8% 0% 17% 8% 0.09 July 2002 92% 83% 83% 86% 0.05 August 2002 83% 92% 83% 86% 0.05 September 2003 100% 100% 100% 100% June 2004 67% 100% 67% 78% 0.16 Site Mean 43% 0.43 45% 0.47 40% 0.42 Overall, infection intensity was highest for Bad Luck Bar with a mean Mackin score of 1.28 for all months. This score, however, was not significantly higher than the mean Mackin scores of the other sites, Corrigan Reef and Derrick Key Gap (Table 4-6). With the exception of August 2002, significant differences in infection intensity between
36 particular sites were observed in the summer months but only for particular months. For example, in July 2002, Corrigan Reef (1.1) had significantly lower (p < 0.01) infection intensity than Bad Luck Bar (3.1) and Derrick Key Gap (3.3). However, in September 2003, Derrick Key Gap (2.3) was significantly lower (p < 0.01) than Corrigan Reef (3), but neither site was significantly different from Bad Luck Bar (2.8). In June 2004, Bad Luck Bar (2.3) had significantly higher (p < 0.01) infection intensity than Corrigan Reef (1.1) and Derrick Key Bar (1.5). Table 4-6. Mackin scale scores of the monthly and site infection intensity of positive oyster digestive tract samples by RFTM assay (n=12 oysters per site per sampling month) Oyster Sites Month Derrick Key Gap Bad Luck Bar Corrigan Reef Monthly mean February 2002 0 0 0 0 March 2002 0 0 0 0 April 2002 0.4 0.25 0.1 0.25 0.15 May 2002 0 0.1 0 0.03 0.06 June 2002 0.1 0 0.4 0.2 0.21 July 2002 3.3 3.1 1.1 2.5 1.22 August 2002 2.5 3 2.6 2.7 0.26 September 2003 2.3 2.8 3 2.7 0.36 June 2004 1.5 2.3 1.1 1.6 0.61 Site Mean 1.12 1.30 1.28 1.46 0.92 1.16 PCR Analysis of Oyster Samples Oyster samples examined by RFTM from the various sites and months were also evaluated by P. marinusâ€“specific PCR assay. DNA was extracted from digestive tract tissues of oysters and clams from all the monthly samples. DNA was also extracted from gill tissues from the samples collected in February and March of 2002, September 2003 and June 2004. A universal eukaryotic primer was used to determine if the stability of the extracted DNA was suitable for PCR analysis. Only positive samples by the universal
37 primers were examined further, while negative samples were excluded. At least four samples were examined for each site with an average of ten samples per site. P. marinus-specific PCR detected 47% and 50% of samples as positive from oyster digestive tract collected in the winter months of February and March of 2002, respectively. These samples were negative for P. marinus infection by RFTM for the same time period (Figure 4-4). Increased detection of positive samples by PCR compared to RFTM was also observed for April, May and June of 2002. Thus, the PCR results for these months indicated greater sensitivity of the P. marinus-specific PCR compared to RFTM. However, increased sensitivity of PCR detection compared to RFTM was not observed in summer months. For example, 88% of the samples were positive during the summer months (July 2002, August 2002, September 2003, and June 2004) by the RFTM assay versus only 39% positive by species-specific PCR. In fact, 44% of the summer oyster samples that were positive for RFTM were negative for P. marinus-specific PCR. In the months of February through June 2002, there was significantly (p < 0.01) more detection of P. marinus by species-specific PCR assay than by RFTM; however, even though all summer months were positive for both assays, RFTM detected 49% more infected samples than PCR. Despite the differences among specific months, there was no significant difference between the two assays in overall infection levels, as determined by the total mean detection for all months. Results from P. marinus-specific PCR assay also did not differ significantly by site: Bad Luck Bar (59%), Corrigan Reef (43%) and Derrick Key Gap (42%). However, differences in site mean percentages within specific months were noted. In June 2002, Corrigan Reef (17%) was lower than Derrick Key Gap (75%) and Bad Luck Bar (75%).
38 In September 2003, Bad Luck Bar (80%) was higher than Derrick Key Gap (0%) and Corrigan Reef (33%). 0%20%40%60%80%100%February 2002March 2002April 2002May 2002June 2002July 2002August 2002September 2003June 2004MonthsPercent Postive RFTM P. marinus PCR Figure 4-4. Monthly mean percentage of positive samples from RFTM assay versus P. marinus PCR assay for oyster digestive tract samples. The X-axis consists of the months and the Y-axis consists of the percentage of positive samples. Standard deviations are shown for both assays. In RFTM assay, means from summer months (July 2002, August 2002, September 2003 and June 2004 were significantly higher (p <0.01) than means from other months (February â€“June 2002). In PCR assay (n 20 per month), there was no significant difference among the monthly means. Several samples that were positive by RFTM were negative by species-specific PCR, suggesting that Perkinsus spp. other than P. marinus may infect oysters in Cedar Key, FL. Therefore, oyster digestive tract samples were also examined using a Perkinsus genus-specific PCR assay. Comparison of PCR assays to RFTM results for oyster digestive tract indicated that the Perkinsusâ€“genus PCR assay detected more positive samples in the months of February and March of 2002 and June 2004 (Figure 4-5). However, the RFTM assay detected Perkinsus spp. in 100% of the samples collected in
39 September 2003, but compared to genus-specific and species-specific assays, 49%, 38%, respectively. Digestive Tract0%20%40%60%80%100%February2002March 2002September2003June 2004Percent Positive Gill0%20%40%60%80%100%February 2002March 2002September2003June 2004MonthsPercent Postive Perkinsus genus PCR P. marinus PCRAB Figure 4-5. Comparisons of monthly mean percentages of positive samples of oyster (A) digestive tract tissue and (B) gill tissue by Perkinsusâ€“genus specific and P. marinusspecific PCR assays. The xaxis consists of the months. The yaxis consists of the percentage of positive samples. Standard deviations are shown.
40 Although, the genus-specific PCR detected 37% more positive digestive tract samples than by RFTM and species-specific PCR, there were no significant differences among the three assays for overall detection based on the mean total for all months. In order to determine whether Perkinsus spp. other than P. marinus were detected by genus-specific PCR, selected PCR products (n=3) from the genus-specific amplification were sequenced. Strains that were either positive for P. marinus-specific primers (n=2) or were positive for genusâ€“specific primers but did not amplify with P. marinusâ€“specific primers (n=1) were purified as described in Materials and Methods. All sequences showed 99% identity to previously published P. marinus sequence (Accession number: AF497479) and differed by only one base over the 500 to 540 bp examined. Interestingly, an identical polymorphism was identified in the sequences of all three samples, with a substitution or addition of a thymine base for guanine base at the same site. Comparison of Infections from Digestive Tract vs. Gill Tissues To determine the distribution of P. marinus in oyster tissues, we also compared the infections in gill versus digestive tract tissues using both PCR assays (speciesand genus-specific PCR). Neither the species-specific nor genus-specific PCR assay detected any differences in infection levels between the two tissues (Figure 4-5). Comparison of the results from the P. marinus species-specific assays indicated that there was no significant difference in infection levels between the two tissues in the months examined. Mean percentages of infection in digestive tract tissues were not significantly different among the months or sites. However, infection level in the gill tissues varied by both month and site. For example, in February 2002, speciesâ€“specific PCR detected infection
41 in 80% of the gill samples at Corrigan Reef while infection was detected in only 22% from Bad Luck Bar. Overall, the genus-specific PCR assay did not detect significant differences in infection in the gill samples compared to digestive tract samples. Examination of oyster tissues by genus-specific PCR indicated a higher percentage of infected samples in digestive tract samples (94%) collected in February and March 2002 compared to infected gill samples (39%), although these differences were not significant. In the gill samples, significantly more samples were positive in September 2003 and June 2004 than in February and March 2002, p < 0.01 (Figure 4-5B). There were no significant differences among the site mean percentages for the Perkinsus spp. digestive tract or the gill tissue by genus-specific PCR assay. Overall, 59% of oyster gill samples that were determined to be negative for P. marinus by species-specific PCR (n=62) were positive by genus-specific PCR, while 83% of oyster gill samples that were determined to be positive for P. marinus by species-specific PCR (n= 60) were positive by genus-specific PCR. RFTM and PCR Analysis of Clam Samples In contrast to oysters, clams exhibited no association with P. marinus, as none of the samples, from either digestive tract or gill tissue, were positive by the species-specific assay. In order to determine the incidence of Perkinsus infections in M. mercenaria in Cedar Key, FL, clams were examined for the presence of Perkinsus species by genus-specific PCR and RFTM and specifically for P. marinus by species-specific PCR. Clam digestive tract samples were examined monthly at three sites over nine months for species-specific PCR and RFTM. The digestive tract samples that were collected in February and March of 2002, September 2003, and June 2004 were also examined by
42 genus-specific PCR assay. Gill tissues from the clams were also examined by species-specific and genus-specific PCR but not by RFTM, except for the gill tissues collected in June 2004 which was examined by all assays. Perkinsus spp. were detected by RFTM in clam digestive tract samples only from those collected in June 2002 and 2004. Perkinsus spp. were also detected in June 2004 by genus-specific PCR assay. In June 2002, RFTM infection was detected in 17% of clams from one site (Pelican Reef) with a mean Mackin scale score of 0.25, ranging from 0-2. In June 2004, RFTM detected Perkinsus spp. in 33% and 8% of the digestive tract samples collected at two sites (Pelican Reef and Gulf Jackson, respectively) with a Mackin score of 1 in all infected samples (Figure 4-6). Clam samples collected from Pelican Reef and Gulf Jackson were also positive by Perkinsus-genus PCR with 20% and 88%, respectively (Figure 4-6). Overall, 35% of the digestive tract samples collected in June 2004 were positive by Perkinsus genus-specific assay. Gill tissues were also negative by P. marinus-specific PCR. In June 2004, RFTM detected Perkinsus spp. at only one site (Gulf Jackson) at which 17% of the gill samples were positive with a Mackin score of 1(Figure 4-7). Perkinsus was also detected in the corresponding digestive tract of one of these gill samples by both RFTM and the genus-specific PCR. The Perkinsus genus-specific PCR assay detected infection at all three sites, Pelican Reef (11%), Gulf Jackson (22%) and Dog Island (45%). Of 29 samples, 28% were determined positive by the genus-specific PCR (Figure 4-7). Clam Infection Study In order to determine the susceptibility of clams to Perkinsus marinus, either 10 2 or 10 5 P. marinus cells in ASW (n=200) were artificially inoculated into the body cavity of clams (n=200). Control clams were inoculated with 1 ml ASW. Clams were examined
43 biweekly for 8 weeks using the RFTM assay. None of the clam samples exhibited infection by P. marinus following artificial inoculation, as determined by the RFTM assay. June 20020%20%40%60%80%100%Pelican ReefGulf JacksonDog IslandPercent Positive June 20040%20%40%60%80%100%Pelican ReefGulf JacksonDog IslandSitesPercent Positive RFTM Genus PCR P. marinus PCR A B Figure 4-6. Comparisons of percentage of positive clam digestive tract samples by RFTM, genus-specific PCR, and P. marinus PCR assays. Tissues were collected in (A) June 2002 and (B) June 2004. The X-axis consists of the assays and the Y-axis consists of the percentage of positive samples. For both PCR assays, n 5 per site. Genus-specific PCR was not done with clams collected in June 2002.
44 June 2004 Gill0%20%40%60%80%100%Pelican ReefGulf JacksonDog IslandSitesPercent Positive RFTM Genus PCR P. marinus PCR Figure 4-7. Percentages of positive June 2004 clam gill samples by RFTM, genus-specific PCR, and P. marinus-specific PCR assays. The xaxis consists of the assays. The yaxis consists of the percentage of positive samples. For genus-specific and P marinus-specific PCR assays, n 9 per site.
CHAPTER 5 DISCUSSION AND CONCLUSION Perkinsus marinus was first diagnosed in the Eastern oyster, C. virginica and can be found along the western Atlantic and Gulf Coasts. Few studies have specifically looked at the potential of P. marinus infections in other mollusks, such as the hard clam, M. mercenaria (Ray, 1954; Andrews and Hewatt, 1957; Cheng et al., 1995). Another Perkinsus species, P. atlanticus, is considered a primary parasite of clams native to Europe, along the eastern Atlantic Coast and the Mediterranean Sea; however, the extent of host range for these parasites is unclear. Coss et al. (2001) detected a new species, P. andrewsi, as well as P. marinus, in the hard clam and demonstrated that these Perkinsus species can coexist in the same host. Thus, the commercially important hard clam, often found sympatrically with oysters, could potentially be a vector or serve as a reservoir of infectious life stages of P. marinus. The first specific aim of the present study was to compare the infection level of P. marinus in M. mercenaria versus C. virginica from a region of high density clam aquaculture on the Gulf Coast of Florida. The second specific aim was to test the susceptibility of hard clams to cultured P. marinus infection. Finally, the third specific aim was to survey clams and oysters for other Perkinsus species, using genus-specific PCR. Two independent assays, RFTM and species-specific PCR, were performed to detect P. marinus in M. mercenaria and C. virginica. A genus-specific PCR assays was also performed to detect Perkinsus species. Clam and oyster samples were collected monthly from six sites in Cedar Key, FL area located in the Gulf of Mexico from 45
46 February 2002 to August 2002 and in September 2003 and June 2004. Three of the sites were naturally populated oyster reefs, and three were clam aquaculture sites. Perkinsus marinus Detection in Oysters Perkinsus marinus infections in oysters have been shown to generally increase with increasing water temperature and salinity (for a review see Soniat, 1985). The present study confirmed these results with respect to temperature, as the monthly means of the percentage of positive samples and infection intensities in the summer months (July 2002, August 2002, September 2003, and June 2004) were significantly higher than the winter months (February and March) as determined by RFTM. In the study presented herein, however, temperatures recorded in winter and spring months were not sufficiently low enough to explain differences in prevalence and infection intensity over the time sampled. For example, the significantly lowest (p<0.01) mean temperatures (17.5C and 23.1C) occurred in the winter months of February and March 2002. Although these temperatures were significantly lower (p < 0.01) than temperatures observed in the summer months (28C 34C), they still fall in the temporal range of growth for P. marinus. Chu and Greene (1989) reported that P. marinus is proliferative at temperatures of 20C in vitro, with the optimum range being 25 30C, and growth inhibition was not observed until temperature reached lows of 12-15C. The other major factor reported to influence P. marinus infection rate is salinity. Mean salinity measured at different times during the present study ranged from 21.8 to 30.8 ppt; however, individual samples were as low as 18.4 ppt and as high as 32 ppt. These levels should not inhibit P. marinus cell growth as Perkinsus growth is reported to be inhibited at or below 10 ppt (Chu and Greene, 1989).
47 Although RFTM analysis indicated the presence of P. marinus in oysters corresponded to months with increased temperatures, the P. marinus-specific PCR assay results indicated no significant difference in the infection rates among sampling months, sites or tissues. These PCR results suggested that site, temperature and/or salinity did not have an influence on infection. However, the results of the Perkinsus genus-specific PCR assay for gill samples exhibited significant differences in the percentage of positive samples in the summer months compared to winter months and more closely resembled results from RFTM. For instance, there was no detection of Perkinsus in February and March by RFTM, and lower prevalence and infection intensity were observed in the months of April to June 2002. Although experimental limits for growth at different temperature and salinity have been established, their relationship to the environmental survival of P. marinus is less clear. Quick and Mackin (1971) found no relationship between infection intensity and salinity, but observed a significant relationship between infection intensity and temperature during their study of 86 sites in Florida. Burreson and Ragone-Calvo (1996) also found significant correlation between temperature and both prevalence and infection intensity in their 4-year study in the James River (Virginia). On the other hand, several studies completed in the Gulf of Mexico reported correlations only with salinity. For example, Soniat (1985) found a correlation between salinity and weighted incidence (a measure of intensity infection), but not between temperature and incidence, in his 2-year study of oysters at one site. Craig et al. (1989) also reported a correlation between salinity and infection intensity but found no significant relationship between temperature and infection intensity in their three-month study of Gulf of Mexico oysters at 49 sites.
48 Soniat and Gauthier (1989) also found a positive correlation between infection intensity and salinity and no correlation with temperature. The Soniat (1985) and Soniat and Gauthier (1989) studies were much longer (2 and1.5 years, respectively) then the present study (9 months). Quick and Mackin (1971) and Craig et al. (1989) examined a greater number of sites (86 and 49, respectively) than the present survey of 3 oyster beds. Differences in study designs, site locations, and sampling plans, may account for the variability in correlations among these studies. The results obtained from this present study supported the idea that the variations in salinity and temperature among sites did not influence prevalence or infection levels in this study. Soniat (1996) stated â€œAlthough both factors and, in particular, their interaction are important (Soniat, 1985), most of the variation in levels of parasitism is not explained by variations in temperature and salinity. Other environmental and biological factors, of which we know little or nothing, must significantly affect the levels of parasitism observed in the field.â€ He compared field studies completed in the Gulf of Mexico and found that temperature and salinity can be inversely correlated and have opposing effects on the parasite, especially if the sampling period is short. In 1989, Soniat and Gauthier stated that whenever temperatures are consistently high and salinity varies, salinity appears to be the controlling factor of disease. Likewise, whenever salinities are relatively consistent and temperature varies, then temperature is the controlling factor. In the study presented herein, although both temperature and salinity showed significant variation at times, both parameters were at relatively high levels and probably did not contribute to prevalence of P. marinus.
49 RFTM and PCR Analysis of Clam Samples P. marinus was not detected in any of the clam samples by P. marinus-specific PCR assay. Only 5% of the hard clams in the entire study appeared to be infected with any Perkinsus spp. Detection of several species of Perkinsus in clams has been reported elsewhere. For example, Coss et al. (2001) detected P. marinus and P. andrewsi in several clam species, including the hard clam, in the Chesapeake Bay. Perkinsus atlanticus is recognized as a pathogen of Ruditapes decussates, a clam located in Portugal (Azevedo, 1989). Resistance of the host or in the virulence of the parasite as a result of geographic restrictions may affect exposure or susceptibility to infection (Bushek and Allen, 1996). The natural distribution of hard clams along the Atlantic coast from Nova Scotia to Florida overlaps the natural distribution of P. marinus. Native hard clam populations from the Atlantic coast of Florida were brought to the Gulf coast to improve the economic development of the area in the 1990â€™s. The first documentation of P. marinus in Florida was in Apalachicola Bay by Dawson in 1955. Therefore, clams have had ample opportunity to become infected. It has been reported that Gulf coast isolates of P. marinus are slightly less virulent in oysters than Atlantic isolates (Bushek and Allen, 1996). The P. marinus isolates found in the Gulf coast of Florida may not be virulent enough to cause major infections in hard clams. Alternatively, hard clams might have a higher level of resistance to the Gulf coast strains of P. marinus. Further, the grow-out time and reproductive turn over of clams is much more rapid in the Gulf compared to the clams on the Atlantic coast, which may lessen the impact of shellfish mortalities in this region. In our studies, none of the clams that were artificially inoculated with cultured P. marinus cells exhibited infection. Other studies have also examined the inoculation of the
50 hard clam with P. marinus cells. Both Ray (1954) and Andrews and Hewatt (1957) attempted to experimentally infect hard clams with P. marinus. Ray placed clams in epizootic waters in Louisiana in order to expose the clams to the parasite, which failed to cause detectable infections in either live or dead clams collected over 4 months, whereas adjacent oysters became infected within 13 weeks. Both studies fed infected minced tissue to clams but infection was still not produced. Injections of P. marinus from heavily infected oysters into clams did produce localized infections near the injection site, but they did not spread beyond the inoculation site (Ray, 1954). Andrew and Hewatt (1957) also inoculated large doses of parasites from infected oysters into clams and examined tissue distance from the site of inoculation to avoid the localization artifact. They found no evidence of infection in either live or gaping clams after 1 month. In 1995, Cheng et al. reported transmission of P. marinus from infected oysters to hard clams after a 10-day experiment in which clams and oysters were placed together in beakers filled with seawater. Parasitic infections between the two bivalves were morphologically similar and were found primarily in the connective tissue surrounding the digestive tract. These results indicated that P marinus can infect hard clams under experimental conditions; however, there is no evidence that parasites will spread and maintain persistent infections. Other studies have looked at the infectivity and pathogenicity of different life stages, the required dosage of parasitic cells, and the routes of transmission that are necessary for the oyster infection by P. marinus. These studies provide some insight into why the hard clams did not show infection. Mackin (1962) inoculated oysters with minced tissues containing various concentrations (10-10 6 ) of meronts that had been
51 incubated in RFTM for 24 hours. He found that a dose of 1.0 x 10 2 to 5x 10 2 cells was required to cause mortality within 41 days and that the mortality rate was positively correlated with the inoculated infective cell numbers. However, Mackin only monitored the oyster mortality caused by the inoculations, not the infection intensity. Mackin and Ray (1966) fed and injected infected minced tissue into oysters from Louisiana and Connecticut in order to produce infection in C. virginica. Both methods produced infections and mortalities. In 1994, Volety and Chu compared the pathogenicity and infectivity of two life stages, the meront vs. the prezoosporangia, by inoculating 10 4 of each into the shell cavity of individual oysters. In 40 days, infections developed in oysters inoculated with both forms. The infection rate and intensity increased with time in both groups; however, rate of infection was significantly higher in the oysters inoculated with the meronts compared to those with the prezoosporangia. In 75 days, the infection intensity ranged from light to moderate in the meront-inoculated oysters, but only light infections were detected in the prezoosporangiaâ€“inoculated oysters. Chu and Volety (1997) reported that the minimum dose required to infect oysters was the same for meronts and prezooporangia (10 2 per oyster) that were inoculated through the shell cavity and that temperature was the most important factor in determining the susceptibility of oysters to P. marinus. Differences in virulence between laboratory cultured cells and freshly isolated cells have also been studied. Oysters infected with cultured meronts did not exhibit an infection rate as high as when meronts were derived from freshly isolated tissue (Chu, 1996). Bushek (1994) examined the effects of feeding versus direct injection of in vitro cultured cells into the adductor muscle or shell cavity. He found that oysters fed as high
52 as 10 7 cells per oyster were not infected, whereas light infections developed within 50 days with a dosage above 10 4 cells per oyster when inoculated into the shell cavity or adductor muscle. LaPeyre et al. (1993) dosed oysters with 10 6 cultured cells per oyster and detected only light infections in 8 weeks; no mortality was noted. Chintala et al. (1995) also compared the infectivity of cultured cells and cells isolated from infected oysters. After 12 weeks, 75% of the oysters injected with natural isolates died with heavy infections, whereas only 7.5% of the oysters injected with cultured cells died. In contrast, Gauthier and Vasta (1993) injected uninfected oysters biweekly with 2 x 10 5 cultured cells. In 4 -5 weeks heavy infections occurred, and mortalities were observed in 6-8 weeks. The cultured cells used by Gauthier and Vasta appeared to be more virulent than those used by Bushek (1994), LaPeyre et al. (1993), and Chintala et al. (1995). The reason for differences in infectivity of the cells is unknown, but may be due to the differences in culture media, age of culture, or the origin of the isolates (Chu, 1996). The latter three studies used the same medium that was developed by LaPeyre et al. (1993), but Gauthier and Vasta developed and used a very different medium. In a three-part study, Ford et al. (2002) compared the virulence of wild-type P. marinus cells to cultured cells. They reported that the wild-type caused higher mortalities than the cultured cells. At the end of the 3-month experiment, most of the oysters that were inoculated with cultured cells were still alive. They also reported that log-phase parasites were significantly more virulent than those obtained from lag or stationary phase cultures; however, they reported that virulence of P. marinus cells was lost immediately when put in culture media. In the same study, Chintala et al. (2002) reported that injection of wild-type and cultured cells of P. marinus into the adductor-muscle
53 produced the heaviest infections followed by shell-cavity injections. Although the virulence of the wild-type cells was higher than the cultured cells, the ranking of the dosing methods was the same, indicating that the higher virulence of the wild-type P. marinus is not due to their ability to take advantage of one particular route or overcome a certain barrier. The cells used in the present study were cultured in the Gauthier and Vasta medium and injected into clams in order to optimize infection. However, infections still were not detected in clams. The culture phase was probably early stationary (2-3 day incubation); but differences in growth phase or route of inoculation were not compared. We chose to inject clams in the shell cavity with P. marinus as the literature indicated this method was generally more effective. For future studies on the susceptibility of clam to P. marinus, an experimental design similar to the 3-part study by Ford et al., 2002 should be done. Clams would be inoculated with a single dosage of wild type P. marinus at 10 6 g -1 wwt in both adductor muscle and shell cavity for comparison. Other variables, such as temperature, salinity and dose should be optimized for clam infections, as these parameters may differ from those reported for oysters. We used cells that we obtained from Dr. Gauthier, which had been shown previously to be virulent in oysters (Gauthier and Vasta, 1995). However, cultures were subcultured for months prior to inoculation which may result in decreased virulence. In future studies, fresh cultures probably should be obtained from frozen stocks or from infected oysters to ensure virulence. Perhaps the most important positive control for future studies would be to first assess virulence of P. marinus in oysters, as they are the primary hosts. Recovery and culture of strains from fresh infections should increase the probability of strain virulence.
54 Comparisons of Detection Assays Comparison of assays for detection of P. marinus in the present study suggested observed differences in correlations of P. marinus oyster infections with temperature and salinity may be an indication of the relative lack of sensitivity of a particular assay. For example, Ragone-Calvo and Burreson (1994) used an antibody detection method, while Bushek et al. (1994) used a full body burden FTM assay, and both methods showed that detection did not decline as dramatically during the winter as routine RFTM assay had suggested. In the present study, P. marinus-specific PCR assay detected more positive samples in the winter months than did the RFTM, but in the summer months, the RFTM detected more positive samples. The RFTM assay lacks species-specificity compared to the P. marinus-specific PCR, and thus may detect more than one Perkinsus species. In fact, RFTM assay has been used to help identify and describe other Perkinsus species such as P. mediterraneus (Casas et al., 2004), P. andrewsi (Coss et al., 2000) and P. atlanticus (Azevedo, 1989). Robledo et al., (1998) compared species-specificity and the sensitivity of P. marinus-specific PCR assay (Marsh et al., 1995) to the RFTM assay. They reported that the PCR assay detected infections in more samples than did the RFTM from oysters that were collected in December, a month in which infection intensity and prevalence should be low. They also examined the reliability of the species-specific PCR assay for detection of P. marinus by testing it against P. atlanticus and P. olseni. The P. marinus primers did not detect these other species. In 2002, Robledo et al. developed the Perkinsus genus-specific primers, which were used in this study. To test the primers, DNA extracted from cultured cells of P. marinus, P. andrewsi and P. atlanticus were used as well as DNA from dinoflagellates, uninfected clam hosts (but not the hard clam) and the Eastern oyster. Thus, it was shown that the genus primer could reliably detect the
55 cultures of different species, but the ability to detect genetically variable strains of species from environmental samples was not examined. In summary, both P. marinusâ€“specific PCR and Perkinsus genusâ€“specific PCR assays appeared to provide more sensitive detection in winter months as compared to RFTM analysis; however, this increased sensitivity was not apparent in the summer months. Overall, in our studies, both P. marinus-specific PCR and RFTM assays were positive for 45% of total samples, while the Perkinsus genus-specific PCR detected positive infections in 82% of all oyster samples. There are several possible explanations for the discrepancies among these assays. For example, samples that were positive for both RFTM and Perkinsus genusâ€“specific PCR but negative for P. marinusâ€“specific PCR may indicate detection of a species other than P. marinus. The genusâ€“specific assay detected significantly more infected samples than either the P. marinus-specific PCR or RFTM assay. The Perkinsus genus-specific PCR assay was designed to detect three species: P. marinus, P. andrewsi and P. atlanticus (Robledo et al., 2002), but may actually have a broader detection range outside of Perkinsus spp. Alternatively, some Perkinsus spp. may not be detected by RFTM but are still positive by the genus-specific PCR such as P. qugwadi (Blackbourn, 1998) Some oyster samples were positive by RFTM and negative for P. marinus-specific PCR during summer months when heavy infections were expected. A possible explanation for differences in PCR assays sensitivity between winter and summer samples could be genetic variability in the PCR target of the P. marinus-specific PCR assay. In other words, variation in the DNA sequence of the primer sites may lead to false negative results. Genetic variability within species derived from different geographical
56 locations is possible (Bushek and Allen, 1996). Robledo et al., (1999) reported 2 distinct sequence patterns that differed by 1-nucleotide from the published sequence amplified by the P. marinus primers developed by Marsh et al., 1995. In this present study, PCR products from Perkinsus genus-specific PCR positive samples that were either negative or positive by P. marinusâ€“specific PCR assay were sequenced. Although sequencing was limited, sequence identity to the previously published strains confirmed that these strains were P. marinus. Thus, we cannot explain the lack of amplification by P. marinus-specific PCR from our data. Divergent DNA at the primer sites may account for these discrepancies, but this sequence would not be detected in our assays and cannot be compared. Finally, discrepancies between assays may result from inhibition of the PCR reaction. Several different protocols were evaluated in an attempt to increase the sensitivity of the PCR assay. The DNA extraction protocol (Qiagen) was changed in the following ways: Tissue concentrations were increased from 30 to 60 mg, the proteinase K volume was increased to 48 l, and RNase was added in order to eliminate RNA carryover. These changes in the extraction method from the manufacturerâ€™s (Qiagen) instructions yielded increased DNA amounts but did not increase assay sensitivity. Bleach was used to sanitize tissues and was examined as a possible PCR inhibitor as stated in the Materials and Methods section. A separate study was performed to look at the effects of the 10% bleach rinse on fresh and frozen oyster tissues, and the results indicated that the bleach did not have an effect on the PCR assays. However it was noted that more DNA was extracted from the fresh tissue than from the frozen tissue samples. The majority of the samples examined in this study were frozen and loss of DNA
57 template may have reduced assay sensitivity. Finally, standard Taq polymerase was used in the â€œmaster mixâ€ for initial PCR assays but was replaced by â€œhotstartâ€ Taq polymerase, which increased detection in samples that were previously negative by PCR. Alternatively, sources of inhibition include containments from oyster tissues that may not be entirely excluded by DNA extraction such as metals, restriction enzymes, etc. Conclusions Our data indicated that M. mercenaria currently does not appear to be a reservoir host for P. marinus; however, the potential risk to hard clam aquaculture by P. marinus or other Perkinsus spp. cannot be negated by studies presented herein, as some positive cultures were detected by RFTM and genus-specific PCR. Future studies should include readdressing artificial inoculation of clams using methods optimized for clams. Isolation of Perkinsus spp. detected in clams should also be pursued in order to characterize these strains and assay pathogenicity. Perkinsus species have caused heavy mortalities in shellfish populations worldwide. Methods for prevention of Perkinsus infection have been described. Ford et al. (2001) suggested the use of particle filtration and UV irradiation to prevent infections of P. marinus and H. nelsoni in larvae and juvenile oysters. This method was effective; however, once the juveniles were moved from treated water in the hatchery to natural seawater, they became infected. To date, there are no available treatments in practice to protect aquaculture shellstock from Perkinsus infection. Therefore, monitoring programs, such as the one established in Maryland, are the only means to prevent areas with little to no infection from reaching enzootic levels (Krantz and Jordan, 1996). The standard PCR assay may not be suitable for surveying the prevalence of P. marinus from environmental samples. Although this assay showed increased sensitivity
58 compared to RFTM in winter samples, expected increases in P. marinus infections in summer months were not detected by the P. marinusâ€“specific PCR assay. Genetic variability within an area and/or PCR inhibition could provide false negatives in samples. On the other hand, RFTM assay is not species-specific and genus-specific PCR confirmed the possibility of species other than P. marinus. Further, specificity for the genusâ€“specific PCR is only defined for three species. Thus, none of these assays are currently suitable as the sole method for evaluation of infection in environmental samples. Continued monitoring of this parasite is important to the lucrative clam industry in Cedar Key and the entire state of Florida. The utilization of methods such as the MPCR or real-time PCR in conjunction with specific examination of other Perkinsus species is also recommended due to problems with the standard PCR assay that were observed in this study. In summary, improved methods may provide more reliable results and expedite the monitoring of infection levels in both hard clams and Eastern oysters.
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BIOGRAPHICAL SKETCH Ayana McCoy was born on July 28, 1977, and raised in Baltimore, MD. She graduated in 1995 from Western Senior High, the oldest all-female public high school in the nation. She received her Bachelor of Science in marine science from Savannah State University in May 1999. She began her masterâ€™s studies at the University of Florida in August 2000. While attending UF, she has been active in several organizations, including Delta Sigma Theta Sorority, Inc and the Black Graduate Student Organization. After graduation, she plans to pursue her goals of (1) increasing diversity in the field of marine science and other scientific fields through education and mentorship and (2) opening a bookstore. 67