ABSTRACT Title of Document: ECOLOGICAL AND EXPOSURE ASSESSMENT OF VIBRIO VULNIFICUS AND VIBRIO PARAHAEMOLYTICUS IN AREAS OF IMPORTANCE FOR HUMAN USE IN CHESAPEAKE BAY Kristi Stevens Shaw, Doctor of Philosophy, 2013 Directed By: Dr. Byron C. Crump Associate Professor of Oceanography University of Maryland Center for Environmental Science Current microbial surveillance of water quality in marine and estuarine environments focuses on fecal indicator concentrations to determine suitable conditions for swimming or fishing, including commercial harvest of seafood. However, there are many pathogens in our waters, such as Vibrio vulnificus and V. parahaemolyticus, and it remains unclear how well fecal indicator surveillance protects the public from infection. This dissertation studied V. vulnificus and V. parahaemolyticus at locations in Chesapeake Bay where human contact is likely, in order to quantify dermal transmission to humans, describe the impact of storms on pathogen concentrations in oysters, and quantify antimicrobial resistance. Swim studies at four public beaches in Chesapeake Bay in 2009 and 2011 were the first of their kind to quantify Vibrio exposure by recreating swimmers and to qualify exposure in terms of dermal dose. Estimated exposures correlated with surface water Vibrio concentrations and suggested that the public could be exposed to V. vulnificus and V. parahaemolyticus at rates that may cause illness. To better protect human health, estimates of non-consumption dose-response would be helpful in completing a quantitative microbial risk assessment to calculate relative risk of swimming in waters known to harbor Vibrio bacteria. Oysters, water, and sediment were sampled at an aquaculture facility before and after Hurricane Irene impacted the Chesapeake Bay in 2011. Results indicated no difference in Vibrio uptake between oysters positioned on floats and on bottom sediments, but showed a difference in Vibrio species uptake, with V. parahaemolyticus increasing 1 day post-Irene, unlike V. vulnificus. This study suggests that storm events may increase V. parahaemolyticus in oyster tissue, and that virulent sub-types of both Vibrio species may increase in percent abundance within oysters following a storm event. Antimicrobial susceptibility testing showed that a large percentage of isolates from surface waters in the Chesapeake Bay displayed intermediate resistance to chloramphenicol. Most antimicrobial agents recommended for treatment of Vibrio illness by CDC were effective at controlling growth of V. vulnificus and V. parahaemolyticus. Results suggest treatment of pediatric illness with trimethoprim- sulfamethoxazole and the aminoglycoside, gentamicin, which was the only aminoglycoside 100% effective in controlling Vibrio growth in this study. ECOLOGICAL AND EXPOSURE ASSESSMENT OF VIBRIO VULNIFICUS AND VIBRIO PARAHAEMOLYTICUS IN AREAS OF IMPORTANCE FOR HUMAN USE IN CHESAPEAKE BAY By Kristi Stevens Shaw Dissertation submitted to the Faculty of the Graduate School of the University of Maryland, College Park, in partial fulfillment of the requirements for the degree of Doctor of Philosophy 2013 Advisory Committee: Dr. Byron C. Crump, Chair Dr. Raleigh R. Hood Dr. John M. Jacobs Dr. Erin K. Lipp Dr. Amy R. Sapkota Dr. Reginal M. Harrell, Dean?s Representative ? Copyright by Kristi Stevens Shaw 2013 ii Dedication I dedicate this work to my children, Ruby Katherine and Oliver James. May you always follow your inner voice and find passion in the work that you pursue. Passion will give you the endurance to push forward when other resources are in short-supply. iii Acknowledgements This work has been made possible by funding from National Oceanic and Atmospheric Administration funding of ?Microbial Community Assessments? (EA133C07CN0163), several small research grants from the Horn Point Laboratory Education Committee and a Rivers Fellowship from Horn Point Laboratory. Due to the unconventional framework of this funding, I am indebted. It allowed me to develop a research project that had to be adaptive to both the needs of my curiosities and the questions that were asked of me while I formulated this project. Thank you to the Oceans and Human Health community. You are truly a family of scientists and from my first EcoHealth meeting in 2006, when I sat in a small session with Dr. Rita Colwell, Dr. Mark Strom, Dr. Paul Sandifer and Dr. Juli Trtanj, I knew I had found kindred spirits in the research world. Subsequent Gordon Research Conferences and various meetings and workshops have served as a means to keep my passion for this important and timely field alive and well. Thank you to Dr. Gary Richards and Dr. Salina Parveen, who invited me into their laboratories to learn methods. Thank you to some of my newest colleagues, such as Dr. Jessica Jones and Dr. Craig Baker-Austin, who have had numerous helpful conversations with me regarding research direction and appropriate methods. No doubt I am forgetting the names of numerous others who have helped me along the way. Please accept my thanks and my apologies. It really does take a village. Thanks to my committee for supporting my ideas and interests, coming together as a homogenous group of experts across several disciplines. Thank you for embracing my interests and curiosities, as unconventional as I?m sure they appeared iv to be in respect to the more traditional dissertation projects that others have completed. Thank you to Dr. Raleigh Hood, who branched out from the field of Oceanography, to support my interest in this meld of Oceanography with Public Health. Thank you to Dr. Erin Lipp for offering your advice and time from several hundred miles away. Thank you to Dr. Amy Sapkota for your enthusiastic support and mentorship in applying Public Health principles to Marine Science problems. I greatly appreciate the time and guidance that Dr. John Jacobs has provided, and for answering my many, many perplexed phone calls. To both Amy and John, thank you for the generous use of your lab space, time and personnel. Without your guidance, I would not have been able to move these research questions forward to completed projects. Most importantly, thank you to Dr. Byron Crump, my advisor and friend. You have been my biggest supporter from Day 1, when I came to you with a hair- brained idea of looking for antimicrobial resistance in oysters and we went about, unsuccessfully at first, trying to complete a pilot project. You worked to secure funding that would allow me to develop a research project that did not fit neatly into any funding boxes, or at least any funding boxes that we could successfully draw upon, although we did get that Biosafety Level II Lab that neither you nor I will probably ever get the chance to use! Thank you for allowing me the opportunity to use my excitement for these new interdisciplinary fields to shape my dissertation focus and to travel for so many meetings and workshops, to be with others who understood my interest in Oceans and Human Health and served as mentors from the sidelines. Most importantly, my family and I are in debt to you for your patience, v understanding and commitment for supporting me not only as a student, but also as a mother. You gave me the time and flexibility to be with my babies when they have needed me most and have been compassionate in knowing that I was working very hard at two very different jobs. Thank you. Thank you to my children, who have been patient with my schedule and levels of stress, both in and out of utero. You have attended meetings, traveled internationally, and tempered physically demanding sampling trips and lab work, sometimes only days before your impending births. You have shared your needs for my time and attention with my work, and I know that was not easy for neither you, nor I. I promise to be patient and support your dreams, because I know the support it takes to achieve your goals. You are the most important creations of my time in this Doctoral program, and you have no doubt taught me more than anything I could have learned in a classroom or from a stack of journal articles. Thank you lastly, and most importantly, to my ever-supportive husband, Bryan. You were by my side when I read ?From Monsoons to Microbes? that summer day on the beach, celebrated my acceptance letter into a public health program two days before our wedding, listened to countless hours of research plans that no doubt mystified (and bored) you, and learned just enough microbiology to help me get home at night. You kayaked for hours during sample collections, carried lab supplies that were too heavy for my pregnant body and juggled the entertainment of two children so I could write my dissertation. You are truly my life partner and I am forever indebt to you for making my dreams, yours. vi Table of Contents Dedication ..................................................................................................................... ii Acknowledgements ..................................................................................................... iii Table of Contents ......................................................................................................... vi List of Tables ................................................................................................................ ix List of Figures ............................................................................................................... xi CHAPTER 1: INTRODUCTION .................................................................................. 1 Water Quality and Vibrio .......................................................................................... 2 Vibrio in the Environment ......................................................................................... 3 Virulence Factors of Vibrio ....................................................................................... 4 Vibrio epidemiology .................................................................................................. 6 Advances in Monitoring ............................................................................................ 7 Vibrio in a Warming Chesapeake Bay ...................................................................... 8 Research Significance ............................................................................................. 10 CHAPTER 2: RECREATIONAL SWIMMERS? EXPOSURE TO VIBRIO VULNIFICUS AND VIBRIO PARAHAEMOLYTICUS IN CHESAPEAKE BAY, MARYLAND, USA .................................................................................................... 13 Abstract .................................................................................................................... 14 Introduction ............................................................................................................. 15 Institutional Review Board .................................................................................. 17 Study population .................................................................................................. 18 Randomization of volunteers ............................................................................... 18 Site selection ........................................................................................................ 18 Controls ............................................................................................................... 20 Surface water collection ...................................................................................... 20 Fecal indicator measurements ............................................................................. 20 DNA extraction, detection and quantification ..................................................... 21 Physical/chemical measurements ........................................................................ 22 Data analysis ........................................................................................................ 23 Conversion of handwash qPCR results to cells cm-2. .......................................... 23 Results ..................................................................................................................... 24 Environmental conditions .................................................................................... 24 Enterococci counts .............................................................................................. 24 Sample size calculation for 2011 swim studies ................................................... 36 Swim Study Results ............................................................................................. 25 Total body surface area exposures ...................................................................... 27 vii Discussion ................................................................................................................ 27 Estimate of Exposure: Total Body ...................................................................... 28 Other routes of entry ............................................................................................ 30 Conclusions ............................................................................................................. 30 Tables?????????????????????????????...43 Figures?????????????????????????????..47 CHAPTER 3: IMPACT OF STORM EVENT, HURRICANE IRENE, ON VIBRIO VULNIFICUS AND VIBRIO PARAHAEMOLYTICUS CONCENTRATIONS IN SURFACE WATER, SEDIMENT AND OYSTERS AT AQUACULTURE FACILITY IN CHESAPEAKE BAY, MARYLAND, USA ...................................... 41 Abstract .................................................................................................................... 42 Materials and Methods ............................................................................................ 45 Sampling site ....................................................................................................... 45 Environmental sample collection ........................................................................ 46 Oyster sample collection ..................................................................................... 47 Physical/chemical measurements ........................................................................ 47 Sample size .......................................................................................................... 48 Oyster processing ................................................................................................ 48 DNA extraction, detection and quantification ..................................................... 48 Most Probable Number (MPN) calculation using qPCR results ......................... 50 Statistical analysis ............................................................................................... 50 Results ..................................................................................................................... 51 Hurricane details .................................................................................................. 51 Physical/chemical conditions .............................................................................. 51 Resuspension calculations ................................................................................... 52 Surface water and on-bottom oyster MPN .......................................................... 64 Vibrio vulnificus .................................................................................................. 52 Vibrio parahaemolyticus ..................................................................................... 54 Discussion ................................................................................................................ 56 Tables?????????????????????????????...72 Figures?????????????????????????????..75 CHAPTER 4: ANTIMICROBIAL SUSCEPTIBILITY OF VIBRIO VULNIFICUS AND VIBRIO PARAHAEMOLYTICUS RECOVERED FROM RECREATIONAL AND COMMERCIAL AREAS OF THE CHESAPEAKE BAY AND COASTAL BAYS .......................................................................................................................... 72 Abstract .................................................................................................................... 73 Introduction ............................................................................................................. 74 Materials and Methods ............................................................................................ 76 Sampling sites ...................................................................................................... 76 Sample collection ................................................................................................ 76 Physical and chemical water quality measurements ............................................ 77 Fecal indicator measurements ............................................................................. 77 Vibrio isolation .................................................................................................... 77 Vibrio species confirmation ................................................................................. 78 viii Clinical isolates ................................................................................................... 80 Antimicrobial susceptibility testing ..................................................................... 80 Statistical analyses ............................................................................................... 81 Results ..................................................................................................................... 81 Physical, chemical and bacterial water quality .................................................... 82 Species and virulence identification .................................................................... 82 Prevalence of antimicrobial resistance in V. vulnificus. ...................................... 82 Antimicrobial resistance in vcg+ V. vulnificus .................................................... 83 Prevalence of antimicrobial resistance in V. parahaemolyticus .......................... 83 Antimicrobial resistance in tdh/trh+ V. parahaemolyticus .................................. 84 Friedman two-way ANOVA ............................................................................... 84 Kruskal-Wallis one-way ANOVA ...................................................................... 85 Clinical V. parahaemolyticus .............................................................................. 85 Discussion ................................................................................................................ 87 Treatability of Chesapeake Bay related Vibrio illness in Maryland .................... 87 AST as compared to fecal indicator measurements ............................................. 88 Comparison to other U.S. studies of V. vulnificus and V. parahaemolyticus antimicrobial susceptibility ................................................................................. 89 Study sites and influences of pollution ................................................................ 90 Conclusions ............................................................................................................. 91 Tables?????????????????????????????.101 Figures????????????????????????????...107 CHAPTER 5: CONCLUSIONS ............................................................................... 102 APPENDIX ............................................................................................................... 109 References ................................................................................................................. 116 ix List of Tables Chapter 2 Table 1: Results of all swim studies, including surface water CFU mL-1, and handwash CFU cm-2 with standard deviation (+/-). Table 2: PCR conditions for V. vulnificus and V. parahaemolyticus and associated virulence assays. Table 3. Estimated CFU cm-2 of body surface area, by swim site, for Total Body Surface Area (TBSA) and per cm2 body surface area. Table 4: Mean and 97% upper percentile (UP; in parentheses) of estimated oral ingestion of surface water and Vibrio cells during swimming activity (EPA 2011). Chapter 3 Table 1. Vibrio vulnificus (Vv) and V. parahaemolyticus (Vp) (n=3) concentrations in oysters, surface water and sediment. Table 2. Correlation table of environmental parameters and Vibrio concentrations in oysters, sediment and surface water. Table 3: PCR conditions for V. vulnificus and V. parahaemolyticus and associated virulence assays. Chapter 4 Table 1. PCR conditions for V. vulnificus and V. parahaemolyticus and associated virulence assays. Table 2. Clinical isolates provided by Maryland Department of Health and Mental Hygiene. Sample type, infection source and associated, if any, antimicrobial resistance. Table 3. Physical, chemical and bacterial water quality. Table 4. Comparison of environmental and clinical isolates and their respective associated antimicrobial resistance to a subset of antibiotics to which highest resistance within tested isolates was displayed. x Table 5. Antibiotic resistance (AR) and multiple antibiotic resistance (MAR) by virulence (5A), site (5B) and month (5C). Table 6. Antimicrobial intermediate resistance and resistance, for respective number and percent, denoted for antibiotic class and specific antibiotic. xi List of Figures Chapter 2 Figure 1. Map of swim study locations in Chesapeake Bay. From: Tracey Saxby, Kate Boicourt, Integration and Application Network, University of Maryland Center for Environmental Science (ian.umces.edu/imagelibrary/displayimage-127-5815.html) Figure 2A, 2B. Vibrio parahaemolyticus and V. vulnificus average handwash CFU cm-2 in relation to surface water concentrations for all swim studies. Figure 3. Vibrio parahaemolyticus and V. vulnificus log CFU cm-2 for each swim time point during 2009 swim study. Figure 4A, 4B. Vibrio parahaemolyticus (3A) and V. vulnificus (3B) CFU cm-2, normalized to surface water concentration, in relation to swim time during 2009 swim study. Figure 5A, 5B. Surface water CFU mL-1 of V. parahaemolyticus and V. vulnificus for each swim study. Most CFU mL-1 peaked at approximately the third swim. Chapter 3 Figure 1. (A) Changes over time of log-transformed CFU mL-1 V. vulnificus and V. parahaemolyticus in surface waters; (B) Log-transformed MPN g-1 V. vulnificus and V. parahaemolyticus in oysters; (C) Changes over time of MPN g-1 V. vulnificus and V. parahaemolyticus in oysters based on position in water column and overall averages. Figure 2. Turbidity in Chesapeake Bay during Hurricane Irene. Figure 3. Best track positions for Hurricane Irene, 21 -28 August 2011. Track during the extratropical stage is based on analyses from the NOAA Hydrometeorological Prediction Center. (Avila & Cangialosi 2011) Figure 4A, 4B. Wind speed and direction at study site during Hurricane Irene Data from NOAA station CAMM2. Figure 5. Changes in Vibrio virulence during study. Figure 6. Physical and chemical measurements of the environment. xii Chapter 4 Figure 1. Sampling sites. (Tracey Saxby, Kate Boicourt, Integration and Application Network, University of Maryland Center for Environmental Science (ian.umces.edu/imagelibrary/ displayimage-127-5815.html) Figure 2. 16S rRNA sequencing analysis of a subset of Vibrio isolates tested. Figure 3. Antimicrobial intermediate resistance and resistance by Vibrio species, for number of isolates and number of antibiotics. 1 CHAPTER 1: INTRODUCTION 2 Water Quality and Vibrio Current microbial surveillance of water quality in marine and estuarine environments focuses primarily on fecal indicator concentrations to determine suitable conditions for swimming or fishing, including the commercial harvest of seafood products. Fecal indicator species have been chosen by regulatory agencies, such as the Environmental Protection Agency, as surrogates for harder-to-detect pathogenic bacteria, especially those arising from fecal pollution. However, there are many pathogens in our waters that occur in the absence of fecal pollution, such as Vibrio vulnificus and V. parahaemolyticus, and it remains unclear how well fecal indicator surveillance protects the public from infection (Harwood et al., 2005). Reports of illness and death caused by virulent Vibrio species are increasing, but little is done to protect the public from deleterious health effects associated with these organisms, especially in recreational settings. Moreover, public health records of infection are limited to a small database of reported adverse health outcomes; possibly because most infections of healthy, immunocompetent individuals result in diarrheal disease, which tends to be self-limiting (i.e., not reported to a healthcare provider, and is resolved on its own). For this reason, monitoring reported health outcomes is an ineffective strategy for understanding risk of infection by these pathogens. The foci of this dissertation research was to study V. vulnificus and V. parahaemolyticus in select locations of the Chesapeake Bay region where human contact is likely, in order to quantify dermal transmission to humans based upon exposure time and surface water concentrations, describe the impact of adverse weather on the concentration of these pathogens in surface water, sediment, and oysters, and, determine the prevalence of 3 bacterial antimicrobial resistance. Vibrio in the Environment Vibrio parahaemolyticus and V. vulnificus are Gram-negative, heterotrophic members of natural marine bacterioplankton communities, and, like the other members of these communities, they are subject to natural environmental factors that control their abundance and distribution. Temperature and salinity are key determinants in the occurrence and growth of both Vibrio species although V. parahaemolyticus tolerates a broader range of salinities relative to V. vulnificus (Johnson et al. 2012). A normal salinity range for the two organisms has been reported between 5 and 25 ppt (Motes et al. 1998). For example, a recent drought in North Carolina from 2007-2009 caused an increase in Neuse River Estuary salinity and concurrent loss of detectable V. vulnificus (Froelich et al. 2012). Additionally, high-salinity exposure is considered a viable means to depurate V. vulnificus from retail oysters (Audemard et al. 2011). Vibrio vulnificus and V. parahaemolyticus have been isolated in water temperature ranges from ca 7-36?C (Motes et al. 1998, Parveen et al. 2008, Johnson et al. 2012), although optimal temperature for growth is typically above 17?C (Vezzulli et al. 2013). Based on early studies by Kaneko and Colwell (1973), Vibrio spp. have long been thought to survive in sediments during the winter and move into the water column during the spring in response to warmer water temperatures and nutrient availability. Vibrio bacteria are believed to attach to zooplankton species (e.g., copepods), rich in chitin, which are used as a food source by the bacteria as they travel into surface waters (Kaneko & Colwell 1973). Once in the photic zone of surface waters, these heterotrophic bacteria are sustained by dissolved organic matter (DOM) from phytoplankton, supplied by the 4 processes of excretion, exudation and cellular death (Smith et al. 1995). Vibrio metabolism is broad, with the ability to utilize not only a wide variety of carbon sources, but nitrogen sources (Criminger et al. 2007) and even polycyclic aromatic hydrocarbons (Hedlund & Staley 2001). As a means to survive in nutrient limited conditions, such as often occurs in the water column, Vibrio have developed fitness-associated mechanisms of nutrient acquisition (Asplund et al. 2011). Vibrio may contribute to the aquatic cycling of organic matter, adding their biomass to the cycle after they are grazed upon by flagellates (Beardsley et al. 2003). Vibrio are one of many bacterial genera capable of using a protective strategy, referred to as a ?viable but nonculturable? (VBNC) state, against harsh conditions (Colwell 2000). The VBNC state allows cells to become metabolically dormant, limiting nutrition requirements, and then emerge from dormancy when environmental conditions improve and are favorable for bacterial growth (Nowakowska & Oliver 2012). Environmental triggers for the VBNC state among Vibrio have been recognized to include temperature, salinity, oxygen concentrations and nutrient deprivation (Colwell 2000, Oliver 2005). This strategy has become an accepted reason why Vibrio have historically not been detected during winter sampling efforts. Virulence Factors of Vibrio Vibrio vulnificus and V. parahaemolyticus are opportunistic pathogens of humans. The virulence mechanisms that they utilize are thought to have been developed to acquire nutrients in the environment (Johnson 2013), but are also deleterious to human health. For example, toxic hemolysins allow Vibrio to lyse host erythrocytes or access cell- bound nutrients such as iron (Johnson 2013). In V. parahaemolyticus, the genes 5 thermostable direct hemolysin (tdh), thermostable related hemolysin (trh) and thermolabile hemolysin (tlh) are responsible for three known virulence factors (Johnson 2013). While tdh and trh are commonly associated with clinical isolates, tlh, found in all strains of V. parahaemolyticus, has been found to upregulate in a mimicked human intestinal environment (Gotoh et al. 2010). The Type 3 Secretion Systems (T3SS1 and T3SS2) are also virulence factors found in V. parahaemolyticus that inhibit host?s immune response by way of effectors that cause enterotoxicity and cytotoxicity (Broberg et al. 2011, Johnson 2013). Varying degrees of virulence are associated with V. parahaemolyticus depending on the combination of the virulence factors they carry in their genomes (Broberg et al. 2011). Vibrio parahaemolyticus also possess two flagellae that allow for swimming and swarming, siderophores to chelate iron from the host, and the ability to form an antiphagocytic capsule (Broberg et al. 2011). Vibrio vulnificus is classified into three biotypes (Strom & Paranjpye 2000). Biotype 1 is associated with human infections, Biotype 2 is related to infections of eels and Biotype 3 was recently discovered in fish handlers in Israel (Bisharat et al. 1999). Vibrio vulnificus posseses many extracellular proteins, two of which are known hemolysin/cytolysin molecules (HlyIII, VvhA) similar to those in V. parahaemolyticus (Johnson 2013). Vibrio vulnificus has a single flagellum and is able to encapsulate. Most important to the virulence of V. vulnificus is the lipopolysaccharide (LPS) in its outer cell membrane, which has been linked with human death (Bowdre et al. 1981, Oliver 2012). Estrogen has been shown to be a protective mechanism against the effects of LPS and it 6 has been deduced that this is the main reason why the epidemiology of V. vulnificus infections tends to be dominated by male patients (Merkel et al. 2001, Oliver 2012). Vibrio epidemiology Routes of infection associated with V. parahaemolyticus and V. vulnificus include consumption of seafood and contact with seawater and, in the case of V. vulnificus, animal to human transmission (CDC 1999a, 1999b, 2000). In the United States, illness associated with Vibrio exposure occurs most frequently during the warmer months of the year (United States Food and Drug Administration 1992a). While non-consumption infectious dose is largely unknown for both V. vulnificus and V. parahaemolyticus (FDA 2012), modeled risk assessment results from the United States Food and Drug Administration (FDA) suggest the 50% probability of illness? (ID50) infective consumption dose for V. parahaemolyticus to be approximately 106 to 108 CFU g-1 (CFSAN (Center for Food Safety and Applied Nutrition 2005). Risk of illness modeled by Food and Agricultural Organization of the United Nations/World Health Organization (FAO/WHO) in a 2005 risk assessment approximated an infective consumption dose of 103 to 107 CFU g-1 oyster tissue (World Health Organization. & Food and Agriculture Organization of the United Nations. 2005). Vibrio vulnificus is responsible for 95% of seafood-related deaths in the United States, with the highest rate of mortality among cases of food-borne illness, followed by wound infection, septicemia, and other presentations of infection (Lipp & Rose 1997, Mead et al. 1999), and a 50% mortality rate for individuals at increased risk (e.g., immunocompromised, liver disease) (Oliver 1995). There are approximately 93 serious (requiring hospitalization) cases of V. vulnificus reported in the U.S. annually (Scallan et 7 al. 2011). A study of nonfoodborne Vibrio infections (NFVI) from 1997-2006, before Vibriosis became a nationally notifiable disease, reported V. vulnificus was responsible for 35% of all NFVI illness and 78% of NFVI deaths in the United States (Dechet et al. 2008). By contrast, V. parahaemolyticus is not as lethal as V. vulnificus, rarely progressing to septicemia (5%), but its clinical manifestation of wound infection (34%) is comparable to V. vulnificus (45%) in terms of proportion to other illness classifications (Dechet et al. 2008). While V. vulnificus is rarely reported as gastroenteritis (5%), a large percentage of V. parahaemolyticus infections are gastroenteritis (59%) (Dechet et al. 2008). Vibrio parahaemolyticus has been implicated in a number of recent outbreaks in the United States (CDC 1998, 1999b, United States Food and Drug Administration 1992), and was estimated to be responsible for 19% of all NFVI (Dechet et al. 2008). In the Chesapeake Bay, the Centers for Disease Control and Prevention (CDC) reported 59 illnesses associated with Vibrio spp. infections in 2009 (Maryland and Virginia) (CDC 2011). Preliminary data from CDC shows a 115% increase in reported Vibrio infections in 2010 in relation to 1996-1998 FoodNet data (CDC 2012c). Advances in Monitoring Recent advances in efforts to monitor and predict Vibrio in the estuarine and marine environment will be useful tools in protecting human health. Modeling efforts have been made to predict the likelihood of occurrence of Vibrio in the Chesapeake Bay (Brown et al. 2012). Remote sensing technologies have been developed to link epidemiological and environmental data to predict the presence of Vibrio and associated illness (Ford et al. 2009, Baker-Austin et al. 2010, Baker-Austin et al. 2012). The ability 8 to pair these advances in predicting the occurrence and abundance of V. vulnificus and V. parahaemolyticus with risk assessment models (Dickinson et al. 2013) will move the science of monitoring forward to management implementation. Vibrio in a Warming Chesapeake Bay It is likely that Vibrio populations will spread geographically and increase in response to global climate change (Lipp et al. 2002, Vezzulli et al. 2013). Average global temperatures have increased by almost 1?C since the late nineteenth century and by approximately 0.2?C per decade for the last 25 years (IPCC 2007). Oceans are predicted to acidify, with an expected decrease in pH of 0.4 by the end of the century (Orr et al. 2005). It is not known how increased environmental temperatures will affect important Vibrio reservoirs or hosts, but a recent study suggests that plankton will be resilient to the predicted decrease in surface water pH (Nielsen et al. 2012). Salinity is the primary determinant for the spatial distribution of Vibrio spp. (particularly V. vulnificus, Jacobs et al. 2010) in the Chesapeake (i.e., positioning up and down the Bay), while temperature increases influence when Vibrio spp. are present in significant quantities during the year. If salinity decreases due to projected increases in precipitation events, there will be a broader range of areas where V. vulnificus will be able to grow in the Chesapeake. Paired with a temperature increase, it is reasonable to assume that V. vulnificus and V. parahaemolyticus will have longer active seasons in the Chesapeake Bay as a result of climate change. Additionally, storm events have been thought to be an important mechanism for distributing benthic Vibrio populations into the water column via resuspension of sediments caused by high winds and flushing due to large volumes of precipitation 9 (Randa et al. 2004, Fries et al. 2008, Wetz et al. 2008, Johnson et al. 2010). The frequency and intensity of storm events are predicted to escalate due to global climate change (Goldenberg et al. 2001), with increases in peak wind intensities and near-storm precipitation (Meehl et al. 2007) likely impacting mid-Atlantic areas such as the Chesapeake Bay. Therefore, relatively moderate wind speed and associated wave action in Chesapeake Bay storm events could increase the overall Vibrio density in surface waters. Based on modeled projections of increased precipitation, temperature and sea- level rise, stratification in Chesapeake Bay is projected to increase (Boesch et al. 2007), allowing for more pronounced algal and bacterial blooms in the eutrophic, warm upper waters (Shiah & Ducklow 1994). This feature of global climate change is likely to increase Vibrio in surface waters of the Chesapeake. Benthic Vibrio concentrations will also likely increase, in response the increased organic material available for decomposition resulting from algal blooms. It is conceivable that, as the climate warms, the human population will seek refuge in waterfront activities, including swimming and boating, to relieve themselves from the heat, possibly over a longer recreational season. Such a situation presents itself as a dangerous intersection of higher concentrations of human pathogens with a higher proportion of the public seeking out exposure to the water. Paired with the possibility that virulence may increase in response to global warming, clinical cases may be expected to increase (Oh et al. 2009, Mahoney et al. 2010). 10 Research Significance This research describes the ecological and environmental conditions favoring the transmission of these pathogens to humans. This development of disease transmission theory (i.e., exposure assessment of humans recreating in a Vibrio dense environment, treatability of Vibrio infection, impact of storm events on the concentration of Vibrio in surface waters, sediment and oysters) will benefit public health and environmental management policy-making decisions. Moreover, given the magnitude of importance that climate change commands in making connections between ecological data and human exposure data related to thermophilic pathogens such as Vibrio spp., conducting a baseline exposure assessment of human interaction with waterborne pathogens is an important consideration in protecting public health in the future. Specific Aim 1: Investigate the magnitude of exposure to V. vulnificus and V. parahaemolyticus in select locations of the Chesapeake Bay region where human contact is likely, in order to quantify dermal transmission to humans based on exposure time and surface water concentrations of these pathogens. In order to protect human health in estuarine and marine communities, it is critical to quantify the numbers of microorganisms to which individuals are exposed during routine activities in the marine environment, such as swimming. Swimming exposure assessments were conducted to define and quantify important routes of exposure, to advance the understanding of how people become ill after environmental exposure, and to be included in risk assessments to protect the general public and important sub- populations (i.e., fishermen, boaters, swimmers). Benefits from this study include 11 understanding the relationship between exposure time and dermal acquisition of pathogens in the estuarine-marine environment in relation to the surface water concentrations of those organisms at the times of exposure. Such estimates are needed for constructing quantitative microbial risk assessments of the risk of infection to such pathogens following such exposure. Specific Aim 2: Provide estimates of storm-related V. vulnificus and V. parahaemolyticus density changes in oyster tissues, sediment and surface water at an aquaculture facility in the Chesapeake Bay. According to the Environmental Protection Agency, the Chesapeake Bay is home to 25% of the total shellfish harvesting waters in the United States (EPA 2011a). Recently, the Chesapeake Bay has become the site of interest for oyster aquaculture production to supplement the dwindling wild harvest of bottom dwelling oysters through on-bottom (submerged land) and off-bottom (water column) leases. Summer is generally considered to be a viable oyster-harvest season in Maryland, but summer is also when Vibrio populations reach their peak in the Bay (Wright et al. 1996, Parveen et al. 2008, Jacobs et al. 2010, Johnson et al. 2012). Oysters may concentrate Vibrio up to 100-fold higher than surrounding waters and it is expected that up to 100% of oysters may be contaminated with V. vulnificus and V. parahaemolyticus during summer months (Morris 2003). Thus, the harvest of oysters during seasons when surface water Vibrio populations are at high densities has the possibility to become a pressing issue for seafood safety. 12 Specific Aim 3: Evaluate the degree to which V. vulnificus and V. parahaemolyticus isolates from the Chesapeake Bay are susceptible to common antimicrobial treatments. In the natural aquatic environment, environmental bacteria provide an unlimited source of resistance genes and determinants, and these genes and determinants can be passed to pathogenic bacteria sharing the aquatic environment, producing newly resistant pathogens (Baquero et al. 2008). In one study it was shown that more than 90% of seawater-derived bacterial strains were resistant to at least one antibiotic and 20% were resistant to at least five antibiotics (Martinez 2003). Human infection intensity and associated morbidity-mortality rates for V. vulnificus and V. parahaemolyticus would be greatly increased by resistance to antimicrobial drugs. The spread of antimicrobial resistance in the microbial communities of our waterways is of concern for treatment of waterborne bacterial infections, especially as these infections can progress quickly and are likely lethal if not treated within a short period of time. A compelling study by Baker-Austin et al., (2009) indicated high levels of antimicrobial resistance in V. vulnificus and V. parahaemolyticus isolated from environmentally degraded sites in South Carolina. It was hypothesized that high levels of resistance may stem from antimicrobial compounds produced by naturally occurring environmental bacteria or horizontal transfer of resistance factors from anthropogenically introduced taxa (Baker-Austin et al., 2009). Widening the geographic scope of such studies will offer greater insight to the mechanisms of antibiotic resistance in these taxa. 13 CHAPTER 2: RECREATIONAL SWIMMERS? EXPOSURE TO VIBRIO VULNIFICUS AND VIBRIO PARAHAEMOLYTICUS IN THE CHESAPEAKE BAY, MARYLAND, USA (Formatted for submission to Environmental Health Perspectives) 14 Abstract Background: Vibrio vulnificus and Vibrio parahaemolyticus are ubiquitous in the marine-estuarine environment, but the magnitude of human non-consumption exposure to these waterborne pathogens is largely unknown. Objective: To evaluate the magnitude of exposure to V. vulnificus and V. parahaemolyticus among swimmers recreating in Vibrio-populated waters. Methods: Swim studies were conducted at four individual swimming locations in the Chesapeake Bay in 2009 and 2011. Volunteers swam for set time periods (2-20 minutes (2009), 8 minutes (2011)), and surface water and handwash samples were collected. Vibrio concentrations were determined using quantitative PCR for each timed swim exposure. Descriptive statistics and regression analysis (linear and logistic) were used to evaluate factors associated with exposure. Results: Mean surface water V. vulnificus (Vv) and V. parahaemolyticus (Vp) concentrations were 1128 (95% confidence interval (CI) 665.6, 1591.4) CFU mL-1 and 18 (95% CI: 9.8, 26.1) CFU mL-1, respectively, across all sampling locations. Vibrios in handwash samples (Vv mean 180 (95% CI: 136.6, 222.5) CFU cm-2; Vp mean 3 (95% CI:2.4, 3.7) CFU cm-2) were significantly associated with Vibrio concentrations in surface water (P=<0.01 (Vv), P=<0.01 (Vp)), but not with salinity or temperature (adjusted R2=0.067 (Vv), adjusted R2=0.026 (Vp)). Handwashing reduced Vibrios on subjects? hands by 93.9% (Vp) and 89.4% (Vv). 15 Conclusions: During months when surface waters are host to an elevated abundance of Vibrio cells, a person recreating or working in those waters should expect a significant dermal exposure, highlighting the potential for illness associated with such an exposure. Introduction Vibrio parahaemolyticus and V. vulnificus are normal functioning members of natural bacterioplankton communities in estuarine and marine waters routinely used for recreation. Current microbial surveillance of water quality in these environments focuses primarily on fecal-indicator concentrations to determine suitable conditions for swimming (EPA 2011b). However, it remains unclear how well fecal-indicator surveillance protects the public from Vibrio infection (Harwood et al. 2005). There are approximately 93 serious (requiring hospitalization) cases of V. vulnificus reported in the U.S. annually (Scallan et al. 2011). A study of non-foodborne Vibrio infections (NFVI) from 1997-2006, before Vibriosis became a nationally notifiable disease, reported V. vulnificus was responsible for 35% of all NFVI illness and 78% of NFVI deaths in the United States (Dechet et al. 2008). For individuals at increased risk (e.g., immunocompromised, liver disease), there is a 50% mortality rate (Oliver 1995). By contrast, V. parahaemolyticus is not as lethal as V. vulnificus, rarely progressing to septicemia (5%), but its clinical manifestation of wound infection (34%) is comparable to V. vulnificus (45%) (Dechet et al. 2008). While V. vulnificus is rarely reported as gastroenteritis (5%), a large percentage of V. parahaemolyticus infections are gastroenteritis (59%) (Dechet et al. 2008) Vibrio parahaemolyticus has been implicated in a number of recent outbreaks in the United States (CDC 1998, 1999b, United States Food and Drug Administration 1992), including 16 responsibility for 19% of all NFVI (Dechet et al. 2008). In the Chesapeake Bay, the Centers for Disease Control and Prevention (CDC) reported 59 illnesses associated with Vibrio spp. infections in 2009 (Maryland and Virginia) (CDC 2011). Preliminary data from CDC shows a 115% increase in Vibrio infections in 2010 in relation to 1996-1998 FoodNet data (CDC 2012c). Approximately 50% of all Maryland V. parahaemolyticus and V. vulnificus infections originate from non-foodborne exposures (Maryland Department of Health and Mental Hygiene (DHMH), personal communication). Routes of infection associated with V. parahaemolyticus and V. vulnificus include consumption of seafood and contact with seawater and, in the case of V. vulnificus, animal (fish) to human transmission (CDC 1999a, 1999b, 2000). In the United States, illness associated with Vibrio exposure occurs most frequently during the warmer months of the year (United States Food and Drug Administration 1992a). While non-consumption infectious dose is largely unknown for both V. vulnificus and V. parahaemolyticus (FDA 2012), modeled risk assessment results from the United States Food and Drug Administration suggest that the infective consumption dose producing 50% probability of illness (ID50) for V. parahaemolyticus is approximately 10 6 to 108 CFU g-1 (CFSAN (Center for Food Safety and Applied Nutrition 2005)). Risk of illness modeled by the Food and Agricultural Organization of the United Nations/World Health Organization (FAO/WHO) in a 2005 risk assessment approximated a infective consumption dose of 103 to 107 CFU g-1 oyster tissue (World Health Organization & Food and Agriculture Organization of the United Nations 2005). It is conceivable that non- consumption dose, encountered from direct contact between an open wound and Vibrio- 17 populated media (e.g,, water, surfaces, seafood products), may require a fraction of the consumption-based infectious dose. While reports of illness and death caused by virulent Vibrio species, including V. vulnificus and V. parahaemolyticus, are increasing (Scallan et al. 2011b), very little is known about the magnitude of human exposure to these environmental pathogens. In order to protect human health in estuarine and marine communities it is critical to quantify the numbers of organisms to which individuals are exposed during routine activities in the marine environment, such as swimming. This study investigates the magnitude of exposure to V. vulnificus and V. parahaemolyticus in select locations of the Chesapeake Bay region where human contact is likely, in order to quantify dermal transmission to humans based upon exposure time and surface water concentrations of these pathogens. It also assesses the efficacy of washing in clean water to remove Vibrio from the surface of skin following dermal exposure. This study also describes the surface water conditions favoring the transmission of these pathogens to humans, including the study conditions under which virulent species were encountered. The estimates of exposure produced by this assessment will help quantify possible disease transmission from surface waters to humans in estuarine and marine environments. Institutional Review Board Study design and participant informed consent forms were reviewed and approved by the University of Maryland Institutional Review Board (Protocol: 11-0442). 18 Study population The study population was a convenience sample of individuals recruited from a local academic institution. The initial 2009 swim (Sandy Point) included 19 participants, and subsequent 2011 swims (Choptank, Tred Avon, Chester) included four participants for each study, based upon a power analysis calculation for sample size performed using the 2009 data. Randomization of volunteers Subjects were assigned random letters from A to S, which were associated with each of their samples. Subject names were not associated with those sample letters and no identifiable information was recorded to associate samples with subjects. Site selection Swimming sites were selected based on differing salinities and geographic location to ensure a range of surface water Vibrio spp. concentrations in order to test the correlation of Vibrio spp. concentrations with overall degree of exposure. Swimming beaches on four different rivers in the Chesapeake Bay were chosen: Choptank River, Chester River, Tred Avon River and Chesapeake mid-Bay (Sandy Point State Park) (Fig. 1). All sites are regularly used by recreational swimmers and thus provide a realistic snapshot of exposure levels. Swims were conducted approximately 1-2 hours post high tide to standardize tidal cycle across swims. Swim study times and description of activity In 2009, a total of ten, independent-timed swims were conducted at each site for the same group of swimmers, ranging from 2 to 20 minutes, increasing incrementally. In 2011, a standardized swim time of 8 minutes per swim was chosen based upon the 2009 19 data. The average concentration of Vibrio in handwash samples stabilized at an approximate exposure duration of 8 minutes. Swimmers were requested to keep their hands submerged for the full time they were in the water. Other activity was not restricted. Swimmers were allowed to swim, wade, float, etc., to account for normal swimming behavior and thus, normal exposures. Time between handwash collection and the next subsequent timed swim was approximately 5 minutes. Description of handwash stations and collection Handwash stations were assembled on rectangular, plastic resin folding tables and shaded completely by a tent. Sterile phosphate buffered saline (PBS, pH 7.4, 500 mL) (FDA 1998) was aliquoted into Ziploc freezer bags (1 gallon size) and stored at 4?C until use (<24 h storage), including storage on ice until sample collection. Bags corresponding to each swim volunteer were clipped to a central holding apparatus and opened approximately 1 minute before each discrete swim time was completed. Each volunteer completely submerged their hands and rubbed them together in a vigorous manner for 60 seconds in the bag of PBS following the guidance of Larson et al. (1998), Brower et al. (2000) and Chen et al. (2001). During the Choptank 2011 swim study, an additional handwash sample was taken after each initial handwash to assess the efficiency of handwashing in the reduction of Vibrio concentrations on hands. All bags were immediately sealed upon handwash completion. Samples were either filtered in the field or frozen and filtered in the lab using 0.22 ?m Sterivex-GP polyethersulfone filters (Millipore, Billerica, MA), wrapped in Parafilm M laboratory wrapping film (Bemis Flexible Packaging, Oshkosh, WI), sealed in a labeled 7 oz Whirlpak bag (Nasco, Fort Atkinson, WI) and stored at -20?C until extraction. 20 Controls Control handwash samples were collected (one per person) before individuals entered the water for the first time to account for any background Vibrio spp. on their hands. Also, control sample bags (n=2 at each time point) of sterile PBS were clipped onto the board and opened at the same time as each handwash collection bag to account for any potential airborne contamination. In 2011, swim studies at Tred Avon River and Chester River were conducted on the same day with the same study participants. Participants liberally applied Purell? brand ethyl alcohol antimicrobial hand sanitizer (GOJO Industries, Akron, OH) to their hands before the start of the second swim study to control for any cross-contamination between swim sites and handwash samples. Surface water collection Surface water samples were collected at each sampling location in sterile wide mouth polypropylene 1 L bottles (Nalgene Thermo Scientific, Waltham, MA). Bottles were rinsed three times with surface water and then dipped below the surface for final 1 L collection volume. Surface water (200 mL) was filtered through a 0.22 ?m Sterivex-GP polyethersulfone filter (Millipore, Billerica, MA) using a 60 mL BD luer lock syringe (BD, Franklin Lakes, NJ). Air was pushed through the filter to remove as much water as possible, then wrapped in Parafilm M laboratory wrapping film (Bemis Flexible Packaging, Oshkosh, WI) and sealed in a labeled 7 oz Whirlpak bag (Nasco, Fort Atkinson, WI). Filters were stored on ice until return to the laboratory (approximately 1 h), where they were stored at -20?C until extraction. Fecal indicator measurements Fecal indicator measurements were made following the standard methods for 21 enumerating Enterococci in Standard Methods for the Examination of Water and Wastewater (Eaton et al. 1998). Briefly, surface water samples were filtered in triplicate volumes onto sterile 0.45 um pore size, 47 mm diameter, gridded membrane filters, and plated onto DifcoTM m Enterococcus (BD, Franklin Lakes, NJ) agar. They were incubated for 48 hours at 35?C before inspection for isolate growth. All light to dark red colonies were recorded as presumptive Enterococci. DNA extraction, detection and quantification DNA was extracted following a modified MO BIO Powersoil extraction protocol (Jacobs et al. 2009) and stored at -80?C until use. A Bio-rad CFX96 Touch? Real-Time PCR Detection System (Bio-rad, Hercules, CA, USA) was used to detect total Vibrio vulnificus (Panicker & Bej 2005) and total Vibrio parahaemolyticus (Nordstrom et al. 2007) in each sample using TaqMan chemistry. Samples testing positive for either species were subjected to further qPCR testing for virulence genes (Vv: virulence correlated gene vcg (Baker-Austin 2010); Vp thermostable direct hemolysin (tdh) and thermostable related hemolysin (trh) (Nordstrom et al. 2007)). Quantitative PCR was performed by using 2.50uL of 10X PCR Buffer (Qiagen, Valencia, CA), 1.25uL of 25 mM MgCl2 (Qiagen), 0.50uL of 10 mM dNTP?s solution (Qiagen), 5uL Q solution (Qiagen), 0.45 uL of 5U uL-1 TopTaq DNA polymerase (Qiagen), 0.188 uL of 10 uM internal control primers (each), 0.375 uL of 10uM internal control probe, 2uL internal control DNA, 0.50 uL of 10 uM primer (each), 0.188 uL of 10 uM probe and 3 uL DNA template per reaction, with the exception of the Vv vcg assay, in which 5uL of DNA template was used. DNase/RNase free water was added to bring the total reaction volume. Two-stage qPCR cycling parameters are presented in Table 2. 22 A unique internal control assay, including a primer set, probe with unique fluorochrome, and internal control DNA, was added to each tube, excluding the vcg analyses, to test for the presence and influence of inhibitors (Nordstrom et al., 2007). Positive controls were also run in a separate well of each qPCR assay plate. Strains used were: V. parahaemolyticus USFDA TX2103 and V. vulnificus ATCC 27562. Standard curves were constructed as reported in Jacobs et al. (2010) from spiked environmental matrices and used during each qPCR analysis with the appropriate qPCR parameters. Cycle threshold (Ct) value was plotted against the slope of the standard curve to determine PCR unit quantity of cell equivalents (CFU). Physical and chemical measurements Physical and chemical measurements were taken at each swim time point, including just before the first swim commenced. Measurements were taken with a YSI 556 Multiprobe System (YSI Incorporated, Yellow Springs, OH). Salinity measurements from July 10, 2011 at Choptank River, Cambridge were retrieved from the Maryland Department of Natural Resources monthly sampling on July 13, 2011, collected 1.23 nautical miles from the swim study site. According to almanac data records (http://www.wunderground.com/history/airport/KSBY/2011/7/10/DailyHistory.html), there was no precipitation between July 10 and July 13, so it can be deduced that the salinity was likely similar on July 10. Sample size calculation for 2011 swim studies Sample size was calculated for a desired power of 0.90, preferred detection level of 25 CFU and an alpha of 0.05, using standard deviation calculations from the 2009 swim study 4.89 (between swim), 10.5 (between swimmer) (Vv) and 3.31 (between 23 swim), 4.4 (between swimmer)(Vp) CFU mL-1, respectively. It was determined that three swims were needed per site and three swimmers were needed for each swim. Based on these results, each 2011 study consisted of five swims per location, with four swimmers. Data analysis Quantitative PCR data was exported to Excel spreadsheet format using BioRad CFX Manager? Software (Bio-Rad, Hercules, CA, USA). Statistical analysis was completed using Intercooled Stata 9.1 for Macintosh statistical software (StataCorp LP, College Station, TX). Descriptive statistics include means, standard deviations and range (min to max). A sample size calculation was conducted for 2011 swims based on the 2009 results (type I error rate=0.05; power=0.90). Linear regression was completed to determine degree of variance between handwash concentration by swim length, individual swimmer and surface water concentration. Handwash concentrations were then divided by the corresponding surface water concentration to normalize data before additional linear regression analysis for associations between exposure and salinity- temperature. Logistic regression analysis was conducted to evaluate the association of virulent strains occurrence in handwash samples in relation to surface water density and environmental conditions. Conversion of handwash qPCR results to CFU cm-2. Previously calculated total body surface area (TBSA) averages for adults and children, including the ratio of hand and palm surface area in relation to the TBSA, were used to qualify dermal exposure from the data collected in this study. Measurements of patient hands are routinely employed by physicians to estimate the area of a burn injury (Amirsheybani et al. 2001). The average adult hand (distal wrist to finger tips) is ~1% of 24 total body surface area (TBSA) and palm (distal wrist to base of fingertips) is ~0.5% (Mosteller 1987). A rough, and likely conservative, estimate of the entire area of the hand (palm, fingertips and back of hand) would therefore be approximately double the average percentage of TBSA for a hand, equaling ~2% of TBSA for calculation purposes. The average TBSA for adult males and females is 1.9 m-2 and 1.6 m-2, respectively, with a combined average of 1.73 m-2 (Mosteller 1987). If average handwash densities of each Vibrio species are interpreted as CFUs per hand area, an estimate of density for total body surface area can be calculated by dividing the PCR unit quantity by average hand area such that CFU cm-2 = CFU / (0.04 *17,300 cm2). Results Environmental conditions Average salinity and water temperature (+/-standard deviation) for each of the four swim sites was as follows: 9.9 ppt (+/-0.01), 27.7 ?C (+/-0.22) (Sandy Point); 6.1 ppt (+/-0.00), 31.4?C (+/-0.26) (Choptank); 7.5 ppt (+/-0.48), 31.0 ?C (+/-0.59) (Tred Avon); 5.5 ppt (+/-0.05), 30.9 ?C (+/-0.21) (Chester). Each site experienced small changes in salinity (0-1 ppt) and temperature (0.5-1?C) over the course of each swim study. Sandy Point mean salinity was calculated by averaging the first three swim salinities, as salinity values collected after those times were unreliable. Enterococci counts Enterococci counts confirmed that all swim study sites were appropriately open for recreational swimming according to Maryland?s single sample maximum allowable density at a recreational beach (COMAR 2013), which is less than 104 CFU 100-mL. The geometric mean (+/-standard deviation) of the Enterococci counts (CFU 100 mL-1) for 25 each swim study site were as follows: 22.2 (+/-1.3) (Sandy Point); 9.9 (+/-13.7) (Choptank); 8.8 (+/-9.1) (Tred Avon); 22.5 (+/-7.9) (Chester). Swim Study Results: Surface and Handwash Concentrations Average concentrations (+/- standard deviation) of Vibrio CFU mL-1 in surface water and handwash samples are presented in Table 1. Mean surface water V. vulnificus and V. parahaemolyticus concentrations were 1128 (95% CI: 665.6, 1591.4) CFU mL-1 and 18 (95% CI: 9.8, 26.1) CFU mL-1, respectively, across all sampling locations. Mean Vibrio in handwash samples were 180 (95% CI:136.6, 222.5) CFU cm-2 (V. vulnificus) and 3 (95% CI:2.4, 3.7) CFU cm-2 (V. parahaemolyticus). During the Choptank swim sub-study of handwash efficiency in removal of Vibrio spp. there was an overall average reduction of 93.9% (95% CI: 86.5%, 101.3%) for V. parahaemolyticus and 89.4% (95% CI: 80.1%, 98.7%) for V. vulnificus concentrations in handwash samples. Data were log transformed (log10) to equalize variance before statistical testing with regression analyses. Linear regression analysis demonstrated a significant positive association between V. parahaemolyticus handwash CFU cm-2 and surface water CFU mL-1, predicting the log handwash CFU cm-2 as y= 0.3563*(log surface water CFU mL-1) ? 0.0896 (adjusted R2 = 0.3071; P=<0.002) (Figure 2A). When a similar regression model was fit for V. vulnificus, a significant positive association was also found (adjusted R2 = 0.6139; P=<0.001) and log handwash CFU cm-2 were predicted as y= 0.808*(surface water cells CFU mL-1) - 0.4192 (Figure 2B). Average proportion of CFU cm-2 in handwash samples, in relation to surface water CFU mL-1, were 17.8% (95% CI: 8.5%, 27.2%) (Vp) and 13.1% (95% CI: 9.3%, 17.0%) (Vv) CFU cm-2: CFU mL-1. 26 Since estimated exposure (CFU cm-2) was significantly associated with surface water concentrations, handwash concentrations (CFU cm-2) were normalized to surface water concentrations (CFU mL-1) associated with the time of testing. Once normalized, linear regression models incorporating the independent variables of salinity and temperature were performed to test for a relationship with handwash CFU cm-2. The models accounted for less than 3% of the variability for exposure to V. parahaemolyticus (adjusted R2=0.026;l P=0.0345) and only 6% for exposure to V. vulnificus (adjusted R2=0.067; P=<0.001). Handwash concentrations of Vibrio CFU tended to increase until approximately the third swim of the day and remained fairly constant (V. vulnificus) or decreased (V. parahaemolyticus) for subsequent, longer-timed swims during the 2009 swim study (Fig. 3). There was approximately 10 minutes between each swim interval. During each of the four swim studies (Fig. 4), there was an appreciable increase in surface water concentration of both Vibrio species at approximately the third swim. Time was not found to be a significant predictor of exposure when 2009 swim study data were analyzed with regression analysis. Vibrio CFU cm-2, normalized to surface water CFU mL-1, were plotted against time and demonstrated low to minimal regression coefficients for V. vulnificus (adjusted R2=0.207, P=<0.001) and V. parahaemolyticus (adjusted R2=0. 003, P=<0.01) (Fig. 5). ANOVA tests determined that individual swimmers did not contribute to the variance in the 2009 data (P=0.134(Vp), 0.282(Vv)). Virulence genes Thermostable direct hemolysin positive strains of V. parahaemolyticus were detected in 4.1% of handwash samples (10/243) and 7% of surface water samples (2/28). 27 No thermostable related hemolysin (trh) positive strains were detected. Of the samples positive for tdh, nine handwash samples were from Sandy Point and one was from Choptank River. Sandy Point and Choptank River each had one tdh positive surface water sample. Vibrio vulnificus virulence correlated gene (vcg) was not detected in any of the handwash samples or surface water samples Vibrio parahaemolyticus virulence gene tdh presence was not statistically associated with salinity, temperature or surface water cell concentrations when tested in a logistic regression model (P=0.134). Total body surface area exposures Based on the range of Vibrio concentrations seen in handwash samples, the highest estimated exposure was 43 CFU cm-2 (Vp) and 3060 (Vv) CFU cm-2 and the average estimated exposure was 3 (95% CI: 2.4, 3.7) CFU cm-2 (Vp) and 180 (95% CI: 136.6, 222.5) CFU cm-2 (Vv) (Table 3). Discussion Handwash samples collected during this study suggest that the public is exposed to Vibrio while recreating in waters where such bacteria naturally occur. Secondary handwash studies conducted in 2011 at Choptank River confirm that the handwash methods employed in this study successfully removed a representative sample of Vibrio from swimmers? hands. The positive correlation between surface water concentrations and handwash samples provides a quantitative model to assess the degree of exposure and potential risk while recreating in waters harboring these bacteria. Moreover, virulent strains of V. parahaemolyticus were detected in surface waters and handwash samples, indicating that virulent species are present and the recreating public could be exposed. 28 While data regarding virulent strains was not quantitative, the presence of such strains raises concerns regarding the risk of infection from recreating in waters harboring Vibrio, especially given that the dose of non-virulent strains--let alone virulent strains--needed to cause illness is largely unknown for dermal exposure. Predictive models of surface water V. vulnificus and V. parahaemolyticus concentrations have been developed for the Chesapeake, using the variables of salinity and temperature as the key determinants of surface water bacterial presence and abundance (Jacobs et al. 2010). Other studies have also determined that these are important environmental variables when modeling V. vulnificus and V. parahaemolyticus surface water concentrations in other geographical areas (Zimmerman et al. 2007, Johnson et al. 2010, Baker-Austin et al. 2012, Johnson et al. 2012). By coupling surface water predictive models with regression models of dermal exposure for these Vibrio species, it is possible to estimate an individual?s level of dermal exposure when encountering water of known surface water concentration. These models may provide a powerful predictor of overall exposure for use by public health managers to protect public health. Estimate of Exposure: Total Body A key component to understanding the overall risk to an individual is to qualify the exposure in units of a predicted dose. The first step in translating the overall dermal exposure by an individual, based on these study findings, is to estimate an individual?s total body exposure. The dose-response mechanism for V. vulnificus and V. parahaemolyticus is poorly understood and creates an obstacle in estimating true overall risk. Additionally, genetic virulence markers and overall mechanism of virulence for each 29 Vibrio species are debated within the scientific community, resulting in a level of uncertainty when depending only on virulence markers to estimate overall risk of illness (Staley & Harwood 2010, Thiaville et al. 2011, Jones et al. 2012). By estimating the size of a typical wound for an adult or a child, one can begin to appreciate the relative exposure in terms of dose. For instance, if an adult experiences the average V. vulnificus handwash from this study of 180 CFU cm-2 and a wound is 2 cm2, the person will be exposed to an estimated dose of 360 CFU. This initial estimate of dose is only a ?snap-shot? of overall dose. Depending on the period of time that the person is immersed in Vibrio-laden surface waters, this dose might be multiplied many times as cells move from the environment into the wound and internal body. It is unknown if this dose would cause an infection in an immuno-competent individual, much less someone with compromised immune function or a pre-existing condition known to increase susceptibility to Vibrio illness (e.g., liver cirrhosis). With immuno-compromised populations growing at a rapid pace, it is conceivable that a greater proportion of the population will be susceptible to illness at lower levels of exposure. While overtly immuno-compromised populations stand out (e.g., patients with HIV-AIDS, cancer, organ-transplant recipients), there are emerging populations in rising numbers that should be considered immuno-compromised, including diabetics (CDC 2012b) and those taking steroidal medications (e.g., to control asthma, rheumatoid arthritis, inflammatory bowel disease, etc.) (Myasoedova et al. 2010, Akinbami et al. 2012, Molodecky et al. 2012). Given the increasing sub-population of children becoming ill with asthma and diabetes, it is prudent to consider the most sensitive populations when formulating 30 recommendations for recreational water use associated with Vibrio spp. Children do not have the same robust immune system of an adult and would therefore be susceptible to infection at lower dosages of an infectious organism. Children are also more likely to have skin abrasions from outdoor play, especially during the summer, when they would also be most likely to be exposed to recreational surface waters. While pediatric Vibrio case reports are limited, perhaps due to limited encounters between children and raw seafood, future wound infection and otitis (ear inflammation or infection) cases may be anticipated to increase as surveillance and detection improves for the two studied Vibrio species, as well as the emerging pathogen, V. alginolyticus. Other routes of entry Oral ingestion rates during swimming have been estimated by (Dufour et al. 2006) and are used in the Environmental Protection Agencies Exposure Factors Handbook (U.S. EPA, 2011). Based on these rates, the ingestion of surface water V. vulnificus and V. parahaemolyticus can be estimated using the average bacterial levels found in this study (Table 4). According to these estimates, a child (younger than 18 years of age) may ingest an average of 42,000 V. vulnificus CFU per swimming event or 55,296 CFU per hour. A dose at this level could lead to illness in a child or an immunocompromised adult, although ingestion rates are likely at the lower end of the dose-response continuum and symptoms, if any, may be limited to mild gastroenteritis. Conclusions This study of recreational water exposure to V. vulnificus and V. parahaemolyticus is the first of its kind to quantify the number of bacteria to which recreating swimmers are exposed and qualify that exposure in terms of dermal dose. Due 31 to a lack of information regarding non-consumption dose-response for V. vulnificus and V. parahaemolyticus, it is unknown if current levels of exposure in the Chesapeake Bay are likely to cause illness, but the public is being exposed to V. vulnificus and V. parahaemolyticus at rates for which illness is conceivable. It was confirmed in this study that washing ones hands following exposure to marine water is a useful practice to reduce the number of Vibrio on a person?s skin by a large percentage. In order to better protect human health, estimates of non-consumption dose-response would be helpful in completing a quantitative microbial risk assessment to calculate relative risk of swimming in waters known to harbor Vibrio bacteria. Additionally, these data should be paired with models of surface water Vibrio concentration to predict exposure at local and regional scales. Finally, data should be incorporated into global climate change models to predict ?tipping points? of sea-surface temperature and salinity that may result in an escalation of recreationally acquired illness. 32 Table 1: Results of all swim studies, including surface water CFU mL-1, and handwash CFU cm-2 with standard deviation (+/-). Site Swim # Swim time (min) Surface water Vp CFU mL-1 Vp CFU cm-2 handwash (standard deviation) Surface water Vv CFU mL-1 Vv CFU cm-2 handwash (standard deviation) Sandy Point 1 2 15.86 3.90 (5.74) 631.85 18.68 (26.93) Sandy Point 2 4 61.81 2.17 (1.84) 2411.39 86.90 (83.63) Sandy Point 3 6 70.29 8.70 (10.11) 4699.56 159.40 (195.45) Sandy Point 4 8 56.60 4.68 (4.04) 2544.68 170.94 (302.31) Sandy Point 5 10 10.81 0.90 (0.70) 1546.65 161.44 (162.73) Sandy Point 6 12 14.29 1.39 (1.96) 1792.27 218.45 (204.59) Sandy Point 7 14 32.47 2.93 (8.06) 2373.90 529.57 (814.08) Sandy Point 8 16 17.45 1.89 (4.31) 2839.74 406.41 (574.60) Sandy Point 9 18 15.42 2.39 (3.74) 1742.38 216.01 (248.91) Sandy Point 10 20 28.76 4.67 (3.15) 982.31 193.13 (196.50) Choptank 1 8 19.44 2.32 (1.34) 644.40 88.21 (69.62) Choptank 2 8 29.31 1.37 (1.58) 153.18 17.89 (12.20) Choptank 3 8 26.11 2.88 (2.94) 703.10 79.17 (63.83) Choptank 4 8 12.25 1.30 (1.01) 297.63 26.82 (21.94) Choptank 5 8 10.51 0.60 (1.19) 477.23 19.53 (26.21) Tred Avon 1 8 7.08 3.74 (3.23) 316.39 51.90 (49.43) Tred Avon 2 8 3.00 1.09 (1.55) 164.69 13.21 (14.62) Tred Avon 3 8 10.12 8.17 (10.90) 988.06 133.45 (41.37) Tred Avon 4 8 6.70 0.41 (0.81) 330.67 104.39 (196.15) Tred Avon 5 8 0.00 0.82 (0.84) 123.12 15.28 (7.16) Chester 1 8 0.00 1.45 (2.13) 874.13 70.57 (72.54) Chester 2 8 0.00 2.06 (3.20) 621.47 62.56 (38.87) Chester 3 8 0.00 1.71 (3.42) 239.45 109.54 (104.24) Chester 4 8 0.00 0.28 (0.56) 327.58 52.22 (34.96) Chester 5 8 0.00 0.49 (0.99) 387.00 39.96 (26.57) 33 Table 2: PCR conditions for detecting V. vulnificus and V. parahaemolyticus virulence genes. Primer Primer (forward & reverse)/Probe Concentrations (nM) PCR conditions Vibrio vulnificus/vvh 400/240 1x: 95 ?C for 60 s; 41x: 95 ?C for 5 s, 59 C for 45 s Vibrio vulnficus/vcg 250/180 1x: 95?C for 10 m; 40x: 95?C for 15 s, 60?C for 90 s Vibrio parahaemolyticus/tlh 200/150 1x: 95?C for 10 m; 45x: 95?C for 5 s, 66?C for 45 s Vibrio parahaemoylticus/tdh trh 200/75 1x: 95?C for 60 s; 50x: 95?C for 5 s, 59?C for 45 s 34 Table 3. Estimated CFU cm-2 of body surface area, by swim site, for handwash (HW), Total Body Surface Area (TBSA) and per cm2 body surface area. Swim site Cells CFU per: Overall Vp average overall Vp std dev Highest HW Vp Overall Vv average overall Vv std dev Highest HW Vv Choptank HW 1,315 480 4,844 32,843 34,957 116,795 TBSA 32,881 11,988 121,104 823,914 575,401 2,919,865 cm2 1.9 0.7 7.0 47.5 50.5 169 Tred Avon HW 2,286 1,902 16,909 44,044 66,393 275,770 TBSA 57,152 47,552 422,736 1,032,290 828,803 6,894,238 cm2 3.3 2.8 24.4 59.7 47.9 398.5 Chester 2,346 1,541 4,731.56 46,343 41,962 179,643 TBSA 58,649 38,514 118,289 1,158,582 457,061 4,491,080 cm2 3.4 2.2 6.8 67.0 60.6 259.6 Sandy Point HW 2,633 1,590 29,816 125,290 176,781 1,675,186 TBSA 65,818 39,754 745,403 1,474,215 1,585,427 52,943,859 cm2 3.8 2.30 43.1 218.5 382.6 3,060.3 35 Table 4: Mean and 97% upper percentile (UP; in parentheses) of estimated oral ingestion of surface water and Vibrio CFU during swimming activity (EPA 2011). Surface water ingestion V. vulnificus ingestion V. parahaemolyticus ingestion mL event-1 mL hour-1 CFU event-1 CFU hour-1 CFU event-1 CFU hour-1 Children 37 (90) 49 (120) 41,754 (101,565) 55,296 (135,421) 663 (1,613) 878 (2,151) Adult 16 (53) 21 (71) 18,056 (59,811) 23,698 (80,124) 286 (950) 376 (1,273) 36 Figure 1. Map of swim study locations in Chesapeake Bay. From: Tracey Saxby, Kate Boicourt, Integration and Application Network, University of Maryland Center for Environmental Science (ian.umces.edu/imagelibrary/displayimage-127-5815.html) 37 Figure 2A, 2B: Vibrio parahaemolyticus and V. vulnificus average handwash CFU cm-2 in relation to surface water concentrations for all swim studies. 38 Figure 3. Vibrio parahaemolyticus and V. vulnificus log handwash CFU cm-2 for each swim time point during 2009 swim study. 39 Figure 4A, 4B: Surface water CFU mL-1 of V. parahaemolyticus and V. vulnificus for each swim study. Most CFU mL-1 peaked at approximately the third swim. Individual swim times were not statistically significant (Fig. 5A, 5B). 40 Figure 5A, 5B. Vibrio parahaemolyticus (5A) and V. vulnificus (5B) CFU cm-2, normalized to surface water concentration, in relation to swim time during 2009 swim study. 41 CHAPTER 3: IMPACT OF STORM EVENT, HURRICANE IRENE, ON VIBRIO VULNIFICUS AND VIBRIO PARAHAEMOLYTICUS CONCENTRATIONS IN SURFACE WATER, SEDIMENT AND OYSTERS AT AN AQUACULTURE FACILITY IN THE CHESAPEAKE BAY, MARYLAND, USA (Formatted for submission to Applied Environmental Microbiology) 42 Abstract To determine if storm events, (i.e., high winds, large volumes of precipitation) increase surface water, sediment and oyster concentrations of Vibrio vulnificus and Vibrio parahaemolyticus, this study followed a sampling timeline before and after Hurricane Irene impacted the Chesapeake Bay in late August 2011. Oysters were sampled from two levels in the water column (surface water defined as the upper 0.3 m of the water column, and just above sediment layer) to determine if there was a difference in uptake of either Vibrio species based on location of the oyster. Sediment and surface water samples were directly extracted and tested with real-time PCR, while oyster samples were tested by a combination of real-time PCR and most probable number enrichment methods. Results indicated no difference in Vibrio uptake between surface water and near bottom oyster samples, but did show a difference in species uptake, with V. parahaemolyticus increasing 1 day post-Irene, unlike V. vulnificus. Vibrio concentrations in surface water samples decreased at 1 day post-Irene, but increased at 4 days post-Irene, at the same time oyster concentrations decreased. At 8 days post-Irene, surface water and sediment concentrations were only a fraction of their initial values before the hurricane impacted the area, but oyster concentrations were similar (V. vulnificus), if not higher (V. parahaemolyticus), in comparison to their pre-storm concentrations. This study suggests that storm events may cause a temporally limited increase in V. parahaemolyticus in oyster tissue and that virulent sub-types of both Vibrio species may increase in percent abundance within oysters following a storm event. 43 Introduction Storm events are thought to be important mechanisms for the distribution of benthic Vibrio populations into the water column via resuspension of sediments associated with high winds, and flushing due to large volumes of precipitation (Randa et al. 2004, Fries et al. 2008, Wetz et al. 2008, Johnson et al. 2010). Frequent storm events in the Chesapeake Bay are associated with the summer season, when V. vulnificus and V. parahaemolyticus, autochthonous bacteria known to cause human illness, are at their highest densities in surface waters (Wright et al. 1996, Parveen et al. 2008, Jacobs et al. 2010, Johnson et al. 2012). The frequency and intensity of storm events are predicted to escalate in response to global climate change (Goldenberg et al. 2001), with increases in peak wind intensities and near-storm precipitation (Meehl et al. 2007) likely impacting mid-Atlantic areas such as the Chesapeake Bay. In the Chesapeake Bay, a shallow, partially mixed estuary prone to tidal circulation (average depth 6.5 m), storm events may increase the overall Vibrio density in surface waters with relatively moderate wind speed and associated wave action. According to the U.S. Environmental Protection Agency, the Chesapeake Bay is home to 25% of the total shellfish harvesting waters in the United States (EPA 2011a). Recently, the Chesapeake Bay has become a site of interest for oyster (Crassostrea virginica) aquaculture production to supplement the dwindling wild harvest of bottom dwelling oysters, both through on-bottom (submerged land) and off-bottom (water column) leases (Maryland Department of Natural Resources, Shellfish Aquaculture Program). As of September 2010, 169 aquaculture operation permit applications (~4000 44 acres) were submitted to Maryland Department of Natural Resources for water-column and submerged-land leases (Donald Webster, University of Maryland Extension, personal communication), and a total of 300 submerged-land leases (~3500 acres) and 23 water- column leases (~94 acres) permitted. A small number of new aquaculture operations are in year-round production of retail oysters, with the supposition that many new operations will soon be joining their ranks. Summer is generally considered to be a viable oyster harvest season in Maryland, but summer is also when Vibrio populations reach their peak in the Bay (Wright et al. 1996, Parveen et al. 2008, Jacobs et al. 2010, Johnson et al. 2012). Studies are currently being conducted to determine ways to reduce Vibrio concentrations in oysters (e.g., high salinity relay), but factors influencing the accumulation of high numbers or virulent strains of Vibrio in oysters are not completely understood (Warner 2008, Johnson et al. 2010). Thus, the harvest of oysters during seasons when surface water Vibrio populations are at high densities could become a pressing issue for seafood safety. If Vibrio density in oysters increases after storm events, shellfish managers may need to institute shellfish harvest closure periods to allow for oyster depuration. At present, shellfish harvest areas are closed after heavy rainfall to account for increased fecal coliform levels, and while a recent study found a weak, but significant positive correlation between total Vibrio counts and the occurrence of rainfall two days prior, a negative relationship was found between fecal coliforms and total Vibrio counts (Yamazaki & Esiobu 2012). Thus, shellfish harvest area closures based upon fecal coliform levels are likely not protective against increased Vibrio concentrations in shellfish. 45 This study was conducted to test the hypothesis that a storm event, generating enough wave energy to cause resuspension of sediment, would cause an increase in oyster-tissue density of V. vulnificus and V. parahaemolyticus. Oysters were tested in Taylor-style surface-water floats (Luckenbach et al. 1999) and in on-bottom cages, to determine if there was an accumulation difference based on water column position. Results from this study provide the first estimates of storm-related Vibrio density changes in oyster tissues, sediment and surface water at an aquaculture facility in the Chesapeake Bay. Materials and Methods Sampling site The study was conducted at an oyster aquaculture facility in a mesohaline tributary of the Chesapeake Bay. The oyster facility was approximately 250,000 m2 (6 acres) with a water depth of approximately 1.2 m (4 ft) at low tide and 2.1 m (7 ft) at high tide. The oyster farm had sediment types ranging from predominantly sand to predominantly silt. The sampling location within the oyster farm was chosen for the predominance of silty sediment (20.4% sand: 66.6% silt: 13.0% clay)(Micheal Owens, Jeffrey Cornwell, University of Maryland Center for Environmental Science, personal communication). Three sampling sub-locations were chosen along the outermost matrix of oyster floats, which covered approximately 1 acre, both for the sediment composition and the likelihood of the area being unprotected from wind events and resultant resuspension activity. Estimates of wind speeds and resultant wave height were made using equations from Young and Verhagen (1996). Calculations of maximum bottom- sheer stress were made according to Sanford (1994) incorporating an approximate bottom 46 depth of 1m and sand grain roughness of 0.0005 m. Erosion rate was calculated using the equation E (g m-2 hr-1) = Mo (kg m-2 s-1 Pa-1) * 3600 sec hr-1 * 1000 gm kg-1 * (taub ? tauc) (Pa), with site-specific estimates of tauc = 0.025 Pa and Mo = 0.000315 kg m -2 s-1 Pa-1 (Taub: bottom-related sheer stress; Tauc: current-related shear stress; Pascal (Pa); Mo is erosion rate constant) (Sanford, Kwon, University of Maryland Center for Environmental Science, personal communication). These calculations do not acknowledge the potential for a wave-dampening effect by the large array of oyster floats tied together at the aquaculture site, although a physical oceanographer conducting experiments at the same site shares that long period waves at the bottom of the water column are damped out by perhaps as much as 50% by the floats, but not so much that resuspension would be negated (Lawrence P. Sanford, University of Maryland, personal communication). Environmental sample collection Baseline surface water, oyster and sediment samples were collected from the field location on 26 August 2011, the day before Hurricane Irene, and any associated storm impacts, was forecast to be present along the Maryland coastline. Subsequent samples were taken at time points 1, 4 and 8 d after Hurricane Irene. All samples were collected at approximately 10:00 h to approximate a uniform water and air temperature at the time of sampling due to solar irradiation. Surface-water samples were collected at each sampling location in sterile wide mouth polypropylene 1 L bottles (Nalgene Thermo Scientific 2105-0032) following the methods described by Jacobs et al. (2009). Briefly, surface water (200 mL) was filtered through a 0.22 ?m Sterivex-GP polyethersulfone filter (Millipore, Billerica, MA) using a 47 60 mL BD luer lock syringe (BD, Franklin Lakes, NJ), wrapped in Parafilm M laboratory wrapping film (Bemis Flexible Packaging, Oshkosh, WI) and sealed in a labeled 7 oz. Whirlpak bag (Nasco, Fort Atkinson, WI). Filters were stored on ice until return to the laboratory (approximately 1 h), where they were stored at -20?C until extraction. Oyster sample collection Oyster samples (a composite of six oysters (Kaufman et al. 2003)) were collected from the top (n=3) and bottom (n=3) of the water column on each of the four sampling dates. Oysters had shell heights (oyster hinge to opposite edge periphery) of approximately 8 cm (3.1 in). Surface water oyster samples were collected from Taylor- style floats and bottom-water oyster samples were enclosed in 1.3 cm mesh bags deployed inside of crab pots to keep the oysters at the bottom of the water column, but out of the sediment layer. Upon removal, oysters were immediately placed in a refrigerated cooler (ice covered by insulation material) and transported to the lab within an hour, where they were immediately processed. Crab pots consistently had a coating of top layer sediment on the bottom of the pot from being deployed in the sediment. That sediment was collected at each of the three sites by filling a 50 mL Falcon sterile polypropylene conical centrifuge tube (BD Vacutainer Labware Medical 352070). Sediment samples were placed on ice and transported back to the laboratory within 1 h, where they were frozen at -20?C until defrosted and extracted following the PowerSoil extraction method. Physical/chemical measurements Temperature, salinity, conductivity, and dissolved oxygen were sampled using a YSI Model 85 (YSI, Yellow Springs, OH). Secchi depth was recorded to the nearest 0.05 48 meter. Total suspended solids (TSS) measurements were completed using 250-400 mL of surface water, filtered onto pre-weighed 47mm glass fiber-filter membranes. Sample size Based on standard deviations reported in Johnson et al. (2010), sample size needed was calculated for a statistical power of 0.8, significance criterion of 0.05, and preferred detection difference of 500 CFU gram-1. Based upon this calculation, three samples were required for each depth (top and bottom), per sampling period. Oyster processing Six oysters (Kaufman et al. 2003), collected from each sampling location (top (n=3), bottom (n=3)), were homogenized following the three-tube MPN method as described in the U.S. Food and Drug Administration Bacteriological Analytical Manual (BAM) methods (DePaola & Kaysner 2004) with slight modifications. Briefly, oysters were scrubbed, shucked with a sterile knife into a sterile blender, diluted with an equal weight of sterile phosphate-buffered-saline (FDA 1998) and blended for 90 s to create a 1:1 (wt:wt) shellfish:diluent homogenate. A 1:20 dilution of oyster homogenate was prepared in triplicate by adding 1 mL of the 1:1 diluted homogenate to 9 mL alkaline peptone water (APW; 1% peptone, 1% NaCl, pH 8.5 ?0.2). Additional 10-fold dilutions to 5 x10-7 were prepared volumetrically by transferring 3 x 1 mL portions into 10 mL APW. Following overnight incubation at 35 ? 2?C, the top 1 mL of tubes showing growth were collected and frozen at -20?C. DNA extraction, detection and quantification Extraction of surface water DNA was completed following a modified MO BIO Powersoil extraction protocol (Jacobs et al. 2009). The standard MO BIO Powersoil 49 extraction protocol was used for sediment samples. Extracted DNA was stored at -80?C until use. Quantitative PCR was used to quantify CFU mL-1 and CFU g-1 from each environmental matrix, respectively. DNA template was obtained from MPN cultures by producing crude cell lysates by boiling 1 mL aliquots of APW cultures in 2 mL micro-centrifuge tubes for 10 min. Following boiling, tubes were plunged into ice until cool and then centrifuged at 14,000 xg for 2 min. Supernatant template was added to real-time PCR reactions at concentrations of 3-5 uL (see PCR methods) to determine presence or absence of V. vulnificus and V. parahaemolyticus in cultured samples. Bio-rad CFX96 Touch? Real- Time PCR Detection System (Bio-rad, Hercules, CA, USA) was used to confirm the species with primers designed to detect V. vulnificus (Panicker & Bej 2005) or V. parahaemolyticus (Nordstrom et al. 2007). Following initial detection, samples testing positive for either species were subjected to further PCR testing for virulence genes (V. vulnificus: virulence correlated gene (vcg) (Baker-Austin 2010); V. parahaemolyticus: thermostable direct hemolysin (tdh), thermostable related hemolysin (trh) genes (Nordstrom et al. 2007). Quantitative PCR was performed on surface water and sediment sample extracts by using 2.50uL of 10X PCR Buffer (Qiagen, Valencia, CA), 1.25uL of 25mM MgCl2 (Qiagen), 0.50uL of 10mM dNTP?s solution (Qiagen), 5uL Q solution (Qiagen), 0.45uL of 5U/uL TopTaq DNA polymerase (Qiagen), 0.188 uL of 10uM internal control primers (each), 0.375 uL of 10uM internal control probe, 2uL internal control DNA, 0.50 uL of 10uM primer (each), 0.188 uL of 10 uM probe and 3uL DNA template per reaction, with the exception of the V.vulnificus vcg assay, in which 5uL of DNA template was used. 50 DNase-RNase free water was added in a quantity sufficient for a 25uL total reaction volume. Two-stage qPCR cycling parameters were optimized to the conditions presented in Table 3. A unique internal control, including a primer set, probe and internal control DNA, was incorporated simultaneously into each assay, excluding V. vulnificus vcg, to test for the presence and influence of inhibitors (Nordstrom et al., 2007). Positive controls used for each qPCR were V. parahaemolyticus USFDA TX2103 and V. vulnificus ATCC 27562. Standard curves were constructed as reported in Jacobs et al. (2010) from spiked environmental matrices and used during each qPCR analysis with appropriate parameters. Cycle threshold (Ct) value was plotted against the slope of the standard curve to determine PCR unit quantity. Most Probable Number (MPN) calculation using qPCR results Corresponding qPCR-MPN values were derived using the U.S. Food and Drug Administration MPN calculator, downloaded from the online publication ?Bacteriological Analytical Manual, Appendix 2: Most Probable Number from Serial Dilutions? (http://www.fda.gov/Food/scienceResearch/LaboratoryMethods/BacteriologicalAnalytica lManualBAM/ucm109656.htm). Statistical analysis Oyster MPN g-1, sediment and surface water data (CFU mL-1) were log transformed (log10) to equalize variances. Multivariate analysis of variance (MANOVA) was conducted to test for differences in sampling location (top vs. bottom oyster concentrations), sampling date (oyster, surface water, sediment) and the interaction effect of sampling location and date for each species of Vibrio. Pearson pairwise correlation 51 analysis was conducted for the experimental variables of oyster MPN g-1, surface water CFU mL-1, sediment CFU g-1, virulence genes (tdh and vcg) MPN g-1, salinity, temperature, total suspended solids, dissolved oxygen, tidal height and secchi depth. Results Hurricane details During the early morning hours of 28 August 2011, Hurricane Irene was just off the Delmarva coastline and the associated winds and rain impacted the Chesapeake Bay region (Figure 1). At the study site, there were approximately 18.4 cm (7.23 inches) of rainfall (NOAA 2011). Barometric pressure over the area reached a minimum of 976.2 (mb) at approximately 18:40 h on 28 August 2011. Wind gusts were recorded in excess of 26 m s-1 (58 MPH). Highest sustained winds were measured at 19.5 m s-1 (44 MPH) at 23:30 h on 27 August 2011 (Avila & Cangialosi 2011) (Figure 2A). Tidal height did not deviate from the predicted normal height on the first day of sampling, so there was no Hurricane related tidal forcing at the first sampling time point. Physical and chemical conditions After Hurricane Irene, salinity at the study site dropped from 10.6 to 8.0, and by Day 8 increased to 9.9. Dissolved oxygen increased from 5.01 mg L-1 to 6.37 mg L-1 after the storm, and remaining above 6 mg L-1. Water temperature decreased from 25.6?C to 24.1?C after the storm, and by Day 8 increased to 25.7?C. Secchi depth increased from 0.4 m to 0.45 m on the day after the storm, returned to 0.4 m on Day 4, and increased to 0.55 m on day 8 (Figure 3). Total suspended solids (TSS) started at 25.1 mg L-1 and decreased over the course of the study to 19.5 mg L-1 (Day 1), 14.7 mg L-1 (Day 4) and 14.9 mg L-1 (Day 8). Tidal height ranged from low tide during initial sampling efforts 52 (pre-storm: 0.20 m above mean lower low water (MLLW), Day 1: 0.15 m MLLW) to close to high tide (Days 4: 0.38 m MLLW; Day 8: 0.55 m MLLW). While changes in temperature, salinity, dissolved oxygen, secchi depth and TSS were small, tidal height was significantly correlated with temperature (P=0.001), TSS (P<0.001), and secchi depth (P<0.001). Resuspension calculations Rates of erosion were calculated based on highest wind gusts (26.9 and 22.6 m s- 1) and highest sustained wind speeds (9-9.8 m s-1). Most winds during the storm were moving in a NNE or NE direction. Erosion rates were predicted to range from 2,343 to 3,616 g m-2 hr-1 during periods of wind gusts and 487 to 730 g m-2 hr-1 during highest sustained winds. Given the lowest wind speed (m s-1) during the height of the storm, the oyster farm would have expected an erosion rate of approximately 3x105 g sediment hr-1. Vibrio vulnificus Oyster MPN Average V. vulnificus in oysters (MPN g-1) changed little between the first sampling pre-storm (26 August 2011) and 1 day after the storm (29 August 2011), with an average 6% increase (Table 1, Figure 4B). Average V. vulnificus decreased approximately 83% between Day 1 and Day 4 post-storm, but increased again between Day 4 and Day 8. This pattern of average V. vulnificus in oysters was driven by samples collected from the top of the water column. Oysters collected from the bottom of the water column had approximately the same average number of V. vulnificus MPN g-1 over the time series, with a small increase 1d post-storm (~9%). Overall, V. vulnificus in oysters decreased by only 4% during the study period. A multivariate analysis of 53 variance found no statistical difference between the sampling locations, sampling dates or an interaction effect of sampling date and location for V. vulnificus MPN values of oysters (P=0.7960). Correlation analysis of oyster V. vulnificus MPN g-1 showed no significant associations with any of the sampled environmental variables (Table 2). Surface water and sediment Vibrio vulnificus decreased in surface waters and sediment on Day 1 post-storm, increased markedly on Day 4, and decreased again to very low concentrations on Day 8 (Table 1). One-way ANOVA analysis of sediment and surface water CFU mL-1 determined no statistically significant difference between dates for either variable (sediment, P=0.1261; surface water, P=0.8219). Correlation analysis of sediment V. vulnificus revealed significant negative relationships with the environmental variables of salinity (P=0.0224, R= -0.4641), secchi depth (P=0.0000, R= -0.9343) and tidal height (P=0.0256, R= -0.4548). Correlation analysis of surface water V. vulnificus found significant associations with sediment V. vulnificus concentrations (P=0.0000, R= 0.9882) and secchi depth (P=0.0000, R= -0.8917) (Table 2). Vibrio vulnificus virulence correlated gene (vcg) Vibrio vulnificus? vcg was detected in oysters during each of the sampling dates, but concentrations were reduced during the Day 1 and 4 sampling time points (393 and 105 MPN g-1, respectively) relative to concentrations pre-storm (789 MPN g-1) and on Day 8 (622 MPN g-1) (Table 1, Fig. 5). Interestingly, while concentrations of vcg decreased, the presence of vcg increased from 50% of sampled oysters with detectable virulence (pre-storm and Day 1) to 83% at Day 4 post-Irene. Overall percentage of sampled oysters (2/6) positive for vcg was at its lowest percentage on Day 8 (33%). The 54 percentage virulent V. vulnificus MPN g-1 of overall V. vulnificus MPN g-1 was highest on Day 4 (0.6%). Vibrio vulnificus vcg was detected in both surface and bottom sampled oysters, but not in sediment or surface waters during this study. No statistically significant correlations were found associated with V. vulnificus vcg concentrations. Vibrio parahaemolyticus Oyster MPN Average overall V. parahaemolyticus MPN g-1 approximately doubled between pre-storm and 1 d post-storm (+54% top oysters, +862% bottom oysters), and then decreased by 88% 4 d post-storm (-84% top, -92% bottom) amounting to a 64% reduction from the initial MPN g-1 by day 4. By 8 d post-storm, V. parahaemolyticus MPN g-1 increased by 86% from 4 d post-storm (+508% top, +790% bottom), amounting to a 162% increase from the pre-storm measurements. Highest oyster V. parahaemolyticus MPN g-1 were approximately 54% greater at the end of the study and bottom oyster V. parahaemolyticus MPN g-1 were over six times greater than pre-storm values. Analysis using multivariate analysis of variance found no statistical difference between the sampling locations, sampling dates or the interaction of sampling location and date for V. parahaemolyticus MPN values of oysters (P=0.5415). Oyster V. parahaemolyticus MPN g-1 did not correlate significantly with any of the environmental variables tested (Table 2). Surface water and sediment Vibrio parahaemolyticus decreased in surface waters, but increased in sediment, 1 day after the storm. Surface water V. parahaemolyticus then increased on Day 4 post- 55 storm and decreased on Day 8 post-storm. Conversely, sediment V. parahaemolyticus decreased on Day 4 and decreased further on Day 8 (Table 1). Sediment and surface water Vibrio abundances (CFU mL-1) were log transformed (log10) to equalize variances. One-way ANOVA analysis of difference among sampling dates for sediment and surface water CFU mL-1 showed no statistically significant difference between dates for either variable (sediment, P=0.8080; surface water, P=0.6978). Correlation analysis of sediment V. parahaemolyticus CFU g-1 revealed significant associations with the environmental variables of temperature (P=0.0124, R= - 0.5019), total suspended solids (P=0.0000, R=0.8569), dissolved oxygen (P=0.0094, R=- 0.5187), secchi depth (P=0.0161, R=-0.4856) and tidal height (P=0.0000, R=-0.9592). Correlation analysis of surface water V. parahaemolyticus CFU mL-1 found a significant negative relationship with salinity (P=0.0414, R= -0.4193), secchi depth (P=0.0000, R= - 0.9727) and tidal height (P=0.0024, R=-0.5903). Conversely, a strong positive association was found between surface water V. parahaemolyticus CFU mL-1 and surface water and sediment V. vulnificus (P<0.0001, R=0.9595, R=0.9866, respectively)(Table 2). Vibrio parahaemolyticus tdh/trh The trh gene was not detected in any of the oyster MPN cultures, nor the sediment or surface water samples. The tdh gene was detected in oyster MPN cultures at all time points except on Day 8. Two samples were positive for tdh during pre-storm sampling (average 658 MPN g-1), and three samples were positive post-storm (Day 1, 1239 MPN g- 1; Day 8, 294 MPN g-1). Concentrations of tdh decreased over the sampling period (Figure 5), although overall percent V. parahaemolyticus tdh MPN g-1, when compared to 56 total V. parahaemolyticus MPN g-1, was greatest at Day 4 (2.9%). The percent of sampled oysters positive for tdh was lowest on Day 8 ((2/6) = 33%). Vibrio parahaemolyticus tdh MPN g-1 correlated significantly with surface water V. vulnificus (P=0.0093, R=-0.0097), sediment V. vulnificus (P=0.0400, R=-0.9600), surface water V. parahaemolyticus (P=0.0152, R=-0.9648) and tidal height (P=0.0463, R=-0.9537) (Table 2). Discussion Hurricane Irene produced a significant wind event for the Chesapeake Bay region and wave action was sufficient to cause sediment resuspension at the aquaculture study facility, according to estimates of erosion based on wind speed and direction. Additionally, there was a large amount of precipitation (18 cm) during the storm event. Although our data lacks a sampling time point during the storm, in situ continuous monitoring data archives of turbidity (accessed at Maryland Department of Natural Resources ?Eyes on the Bay;? http://mddnr.chesapeakebay.net/eyesonthebay/index.cfm) depict sharp spikes in nephelometric turbidity units (NTU) during the peak of the storm winds and a rapid subsequent decrease of NTU, most likely due to the large amount of rainfall experienced during the storm and a resultant flushing effect (Figure 6). This flushing affect may be the cause of reduced turbidity and lowered surface water CFU mL- 1 for both Vibrio species 1 d after the storm. Many concentrations of V. vulnificus and V. parahaemolyticus detected during this study were greater than similar studies documenting the detection of these species in the same sampled matrices in the Chesapeake Bay. Maximum concentrations of Vibrio detected in previous studies of oyster tissue were considerably lower (V. 57 parahaemolyticus: 6.0 x 102 CFU g-1 (Parveen et al. 2008), 1.0 x 104 CFU g-1(Johnson et al. 2012, a); V. vulnificus: 1.2 x 104 CFU g-1 (Johnson et al. 2012)) than the findings of this study (V. parahaemolyticus: 5.0 x 104 MPN g-1; V. vulnificus: 2.5x105 MPN g-1). Johnson et al. (2012) detected lower surface water and sediment V. vulnificus concentrations (surface water: 150 CFU mL-1 vs. 1.2 x103 CFU mL-1 (this study); sediment: 3.5 x 104 CFU g-1 vs. 3.6 x105 MPN g-1 (this study)), while V. parahaemolyticus concentrations found in Johnson et al. (2012) were approximately double the concentrations detected in this study (surface water: 60 CFU mL-1 vs. 17.5 CFU mL-1 (this study); sediment: 1.5 x 104 CFU g-1 vs. 6.0 x 103 MPN g-1(this study). The lower oyster MPN g-1 and surface water/sediment V. vulnificus values from previous studies may be due to a difference in sampling depth for oysters (i.e., natural oyster bar depth and open water versus near shore shallows) or a difference in recovery efficiencies of methodologies used in either study, such as under-detection (culture-based methods, previous studies) or detection of non-viable cells by qPCR (direct detection, this study) in sampled surface water and sediment matrices. While there were changes in the average V. vulnificus and V. parahaemolyticus cell densities in oysters, surface water, and sediment, the values quantified in each of these substrates was not significantly different over the course of the study. There was a species difference in oyster tissue absorption immediately after the storm, with V. parahaemolyticus increasing substantially, but V. vulnificus increasing only slightly. Unlike oyster V. vulnificus MPN counts, V. parahaemolyticus MPN counts were similar at each time point for oysters sampled from the top and the bottom of the water column, with levels increasing 2 to 9 times, respectively, from pre-storm MPN concentrations. 58 Previously, it has been shown that V. vulnificus outnumbers V. parahaemolyticus in sediment, oyster tissue and the water column (Johnson et al. 2010). During this study, V. parahaemolyticus cell g-1 was approximately 5% of the total V. vulnificus cell g-1 in sediment, which supports the Johnson et al. findings (Johnson et al. 2010). However, despite the relative dominance of V. vulnificus in sediments, post-storm increases in Vibrio were dominated by V. parahaemolyticus, suggesting species-specific variation in the degree to which these bacteria are resuspended from sediments or are retained in oyster tissues, perhaps differing from V. vulnificus in properties of adhesion to marine aggregates, which may have been subsequently filtered by oysters. Interestingly, on Day 4 post-storm, oyster tissue Vibrio MPN g-1 decreased precipitously from pre-storm concentrations (-82%, V. vulnificus; -64% V. parahaemolyticus), while surface water CFU mL-1 and sediment CFU g-1 increased substantially (+337% and +84%, respectively). On Day 8, oyster tissue V. vulnificus concentrations returned to pre-storm concentrations, while V. parahaemolyticus MPN g-1 concentrations approximately tripled. Conversely, surface water and sediment concentrations decreased to a fraction of their original concentrations at Day 8 post-storm (-92%, -66% V. vulnificus, respectively; -100% for both sediment and surface water, V. parahaemolyticus). One possible explanation for this observation is a bacterial response to the flushing effect from the wind and rain at the study site, but the most likely is changes in filtration rates of the oysters over the course of this study. Oysters have been shown to reduce or halt filtration during periods of high suspended solids (Loosanoff & Tommers 1948), which was probable during the height of the storm (Figure 6). Oysters may have responded to the increase in suspended solids 59 during the storm by reducing filtration until some time between Day 1 and Day 4 post- storm. During the period between Day 1 and Day 4, the dramatic decrease of both Vibrio species in the oysters may have been due to an increase in their filtration rate, possibly depurating the Vibrio from their tissues. Approximately 4.5 million oysters are present at the aquaculture site, in various stages of maturity, and it is conceivable that their associated filtration would produce detectable changes in water concentrations of Vibrio. This depuration may have increased turbidity in the water column (i.e., high TSS, low Secchi depth) and increased surface water concentrations of both Vibrio species. On Day 8 post-storm, the oysters may have re-filtered the Vibrio from the surface water back into their tissues, reducing surface water and sediment concentrations by 1 to 2 orders of magnitude and decreasing turbidity to the lowest level seen during the study. Similar to Fries et al. (2008), who noted an increase in sediment concentrations of total Vibrio when Hurricane Ophelia impacted the Neuse River Estuary, NC, there was also an increase in the sediment concentrations of both Vibrio species during the first four days post-storm. However, this pattern then reversed with an overall decrease in sediment CFU g-1 (-100%, V. parahaemolyticus; -66%, V. vulnificus). Whether this was due to a change in oyster filtration or a difference in how each Vibrio species was introduced into the water column as a function of resuspension, and associated particle adhesion, remains to be understood. In contrast to other studies (Fries et al. 2008, Hsieh et al. 2008, Wetz et al. 2008, Johnson et al. 2010), surface water CFU mL-1 decreased following the storm, possibly due to oyster filtration. Notably, virulent V. vulnificus and V. parahaemolyticus were not detected in surface waters or sediment during the course of this study. This is counter to other study 60 findings, such as Johnson et al. (2010), which reported virulent V. parahaemolyticus at similar frequencies in sediment, surface water and oysters (Johnson et al. 2010). Virulent V. vulnificus was found routinely in oyster tissues, especially on Day 4 when virulence genes were detected in 5 of the 6 oyster samples (Figure 5). The incidence of V. vulnificus vcg in oyster samples increased by approximately 30% and the MPN g-1 of vcg doubled from pre-storm concentrations on Day 4. This finding is counter to previous, laboratory-based studies, examining the relationship between V. vulnificus? virulence in oysters. These previous studies found no change in V. vulnificus virulence during the passage through the oyster (Groubert & Oliver 1994, Staley et al. 2011). Similarly, the percentage of V. parahaemolyticus tdh MPN g-1 increased to 2.9% of total V. parahaemolyticus MPN g-1 at Day 4, although the percent of oyster samples positive for tdh was lower than pre-storm samples (16.7%). Incidence and concentration of virulent V. vulnificus and V. parahaemolyticus were at their lowest points at Day 8. Movement towards increased aquaculture production of oysters in the Chesapeake Bay, in combination with forecasted environmental responses to global climate change (e.g., warmer surface waters, increased frequency and/or intensity of storm events), may create a situation of higher Vibrio density in oysters, especially during the summer harvest season. Because oysters are routinely consumed raw, understanding how oyster concentrations of Vibrio might be impacted is vital. One relevant question posed by the aquaculture community is whether Vibrio concentrations in oysters differ based on their position in the water column. This study found no difference in Vibrio concentrations between oysters collected from the bottom and the top of the water column. The sampling location may have been too shallow to see a true difference in surface versus 61 bottom culture, but since much of the Chesapeake is shallow, and most aquaculture operations will likely be near-shore, this was probably a good indicator site. Our data shows that a temporally limited increase in V. parahaemolyticus may be expected after a storm, although impacts on V. vulnificus are not as clear. Post-storm, it can be anticipated that detectability of virulence genes may be increased in oysters for both Vibrio species. At the end of the study, in-oyster concentrations were approximately similar to the pre-storm concentrations, suggesting a possible ?ambient? concentration of summer Vibrio density. Further testing should be conducted to determine if these levels vary based on site, position in the water column, and after storm impacts. Ideally, further research would have the opportunity to sample before and after separate wind and precipitation events. Given that V. parahaemolyticus and V. vulnificus appeared to respond differently during post-storm in oyster samples, further research is needed to determine if patterns of adherence to oyster tissues is different between V. parahaemolyticus and V. vulnificus, as well as among virulent subsets of each species. On the spectrum of storm intensity, this study fell on the high end of impacts with Hurricane Irene. As Hurricane Irene consisted of both high winds and large amounts of precipitation, it would be useful to examine storm events with a range of wind speeds and precipitation to account for the individual response variables of resuspension and surface water flushing. Such information would help managers of shellfish harvest decide if there should be a cessation of harvest post- storm, what winds or rainfall would be significant for a given aquaculture site, and how long that suspension of harvest should be recommended. At this time, it is inconclusive whether a storm event should trigger closure of a shellfish fishery. However, 62 concentrations of Vibrio in oysters were very high throughout the study and the percentage of samples that were positive for virulent V. vulnificus and V. parahaemolyticus increased after the storm. From the results of this study, it can be concluded that sampling sediment and surface water for general concentrations of Vibrio, or pathogenic subspecies, may not be enough to predict the concentrations in oysters. Additionally, climate change estimates of increases in surface water temperature, changes in salinity and intensity or frequency of storm events may also drive changes to shellfish management practices. 63 Table 1. Vibrio vulnificus (Vv) and V. parahaemolyticus (Vp) (n=3) concentrations. Date Vv average MPN +/- std. err. Vv top average MPN +/- std. err. Vv bottom average MPN +/- std. err. Vv vcg1 average MPN +/- std. err. Vv SW CFU mL-1 +/- std. err. Vv sediment CFU g-1 +/- std. err. 26-Aug-11 313,320 170,001 206,819 0 419,820 258,030 789 353 827 108 363,767 172,175 29-Aug-11 332,936 165,909 206,819 0 459,053 246,702 393 321 318 76 296,857 106,683 1-Sep-11 56,913 30,280 30,394 6,121 419,820 43,624 105 39 3,616 1,216 669,908 431,266 5-Sep-11 302,089 173,302 145,126 43,624 419,820 246,702 662 52 68 9 122,769 91,153 Date Vp average MPN +/- std. err. Vp top average MPN +/- std. err. Vp bottom average MPN +/- std. err. Vp tdh2 average MPN +/- std. err. Vp SW CFU mL-1 +/- std. err. Vp sediment CFU g-1 +/- std. err. 26-Aug-11 28,426 20,281 46,032 2,901 10,820 9,484 658 56 14 1 9,754 6,204 29-Aug-11 87,604 124,368 71,093 117,551 104,116 155,024 1,239 0 7 0.5 14,791 5,555 1-Sep-11 10,235 9,285 11,694 8,898 8,777 11,401 293 0 49 28 20 7 5-Sep-11 74,641 102,537 71,149 117,503 78,133 111,539 0 0 0 0.3 7 5 64 Table 2. Correlation table of environmental parameters and Vibrio concentrations in oysters, sediment and surface water. 65 Table 3: PCR conditions for the detection of V. vulnificus and V. parahaemolyticus virulence genes. Primer Primer (forward & reverse)/Probe Concentrations (nM) PCR conditions Vibrio vulnificus/vvh 400/240 1x: 95 ?C for 60 s; 41x: 95 ?C for 5 s, 59 C for 45 s Vibrio vulnficus/vcg 250/180 1x: 95?C for 10 m; 40x: 95?C for 15 s, 60?C for 90 s Vibrio parahaemolyticus/tlh 200/150 1x: 95?C for 10 m; 45x: 95?C for 5 s, 66?C for 45 s Vibrio parahaemoylticus/tdh trh 200/75 1x: 95?C for 60 s; 50x: 95?C for 5 s, 59?C for 45 s 66 Figure 1. Best track positions for Hurricane Irene, 21 -28 August 2011. Track during the extratropical stage is based on analyses from the NOAA Hydrometeorological Prediction Center. (Avila & Cangialosi 2011) 67 Figure 2A, 2B. Wind speed and direction at study site during Hurricane Irene Data from NOAA station CAMM2. 0 5 10 15 20 25 30 26 -A ug -1 1 27 -A ug -1 1 28 -A ug -1 1 29 -A ug -1 1 30 -A ug -1 1 31 -A ug -1 1 01 -S ep -1 1 02 -S ep -1 1 03 -S ep -1 1 04 -S ep -1 1 W in d S p ee d ( m s -1 ) average wind (m/s) maximum gust (m/s) minimum gust (m/s) 970 980 990 1000 1010 1020 1030 0 5 10 15 20 25 30 35 26 -A ug -1 1 27 -A ug -1 1 28 -A ug -1 1 29 -A ug -1 1 30 -A ug -1 1 31 -A ug -1 1 01 -S ep -1 1 02 -S ep -1 1 03 -S ep -1 1 04 -S ep -1 1 at m o sp h er ic p re ss u re ( m b ) Te m p er at u re ( C ) air temp (C) water temp (C) atmospheric pressure (mb) 68 Figure 3. Physical and chemical measurements of the environment. 0 0.1 0.2 0.3 0.4 0.5 0.6 0 5 10 15 20 25 30 8/2 5/1 1 8/2 6/1 1 8/2 7/1 1 8/2 8/1 1 8/2 9/1 1 8/3 0/1 1 8/3 1/1 1 9/1 /11 9/2 /11 9/3 /11 9/4 /11 S ecc hi d ep th (m) S al in ity , D iss ol ve d O 2 (m g L- 1 , Te m pe ra tu re ( C ), T SS (m g L- 1 ) , Sampling Date salinity dissolved oxygen (mg L-1) water temperature (C) TSS (mg L-1) secchi (m) 69 Figure 4. (A) Changes over time of log-transformed CFU mL-1 V. vulnificus and V. parahaemolyticus in surface waters (+/- standard error); (B) Log-transformed MPN g-1 V. vulnificus and V. parahaemolyticus in oysters (+/- standard error);; (C) Changes over time of MPN g-1 V. vulnificus and V. parahaemolyticus in oysters based on position in water column and overall averages (+/- standard error);. 70 Figure 5. Changes in Vibrio virulence during study. 71 Figure 6. Turbidity in Chesapeake Bay during Hurricane Irene. 1 10 100 1000 26 -A ug -11 27 -A ug -11 28 -A ug -11 29 -A ug -11 30 -A ug -11 31 -A ug -11 1-S ep -11 2-S ep -11 3-S ep -11 4-S ep -11 W at er c o lu m n t u rb id it y (N T U ) Sassafras R. at Budds Landing Manokin R. at Westover Big Annemessex R. at Coulbourn Creek Corsica R. at Sycamore Point 72 CHAPTER 4: ANTIMICROBIAL SUSCEPTIBILITY OF VIBRIO VULNIFICUS AND VIBRIO PARAHAEMOLYTICUS RECOVERED FROM RECREATIONAL AND COMMERCIAL AREAS OF THE CHESAPEAKE BAY AND COASTAL BAYS (Formatted for submission to Water Research) 73 Abstract Vibrio vulnificus and V. parahaemolyticus in the estuarine-marine environment are of human health significance and may be increasing in pathogenicity and abundance. Vibrio illness originating from contact with waters of the Chesapeake Bay or through seafood originating from the Chesapeake, can cause deleterious health effects, particularly if the strains involved are resistant to clinically-important antibiotics. To our knowledge, little data exists regarding antimicrobial susceptibility patterns of V. vulnificus and V. parahaemolyticus isolated from the Chesapeake Bay. The purpose of this study was to evaluate antimicrobial susceptibility among these pathogens. Surface- water samples were collected from three sites, of recreational and commercial importance, from July to September 2009. Samples were plated onto species-specific media and resulting V. vulnificus and V. parahaemolyticus strains were confirmed using polymerase chain reaction assays and tested for antimicrobial susceptibility using the Sensititre? microbroth dilution system. Descriptive statistics, Friedman two-way Analysis of Variance (ANOVA) and Kruskal-Wallis one-way ANOVA were used to analyze the data. Vibrio vulnificus (n=120) and V. parahaemolyticus (n=77) were isolated from all sampling sites. Most isolates were susceptible to antibiotics recommended for treating Vibrio infections, although some isolates expressed intermediate resistance to chloramphenicol (78% V. vulnificus, 96% V. parahaemolyticus). Vibrio parahaemolyticus also demonstrated resistance to penicillin (68%). No location or month differences were detected in V. parahaemolyticus resistance patterns, but V. vulnificus isolates from St. Martin?s River had lower intermediate resistance than the other two sampling sites during the month of July (P=0.0166). Antibiotics recommended to treat 74 adult Vibrio infections were effective in suppressing bacterial growth, while some antibiotics recommended for pediatric treatment displayed intermediate resistance and resistance. Introduction Bacterial antimicrobial resistance is an important public health consideration with regard to coastal microbiology. Pathogenic bacteria and antimicrobial resistance genes are often released with wastewater discharges into aquatic environments (Baquero et al. 2008). Naturally occurring bacteria produce antibiotics in the environment for signaling and regulatory roles in microbial communities (Martinez 2008). Bacteria protect themselves from the toxicity of the compounds they generate by evolving antibiotic resistance elements (Wright 2007). Because of this resistance, naturally-occurring bacteria are also capable of serving as reservoirs of resistance genes and, coupled with the introduction and accumulation of antimicrobial agents, detergents, disinfectants, and residues from industrial processes, these bacteria may play an important role in the evolution and spread of antibiotic resistance in aquatic environments (Baquero et al. 2008). Vibrio bacteria in the estuarine-marine environment are of particular concern to human health and may be increasing in pathogenicity and abundance (Baker-Austin et al. 2010). In order to protect recreational and commercial users of estuarine-marine environments, and ensure the safety of locally-harvested seafood, the antibiotic resistance patterns among these pathogens need to be better understood. Identifying antibiotic resistance patterns among Vibrio will highlight potential treatment obstacles that the public may experience upon exposure to and infection with these microorganisms. In the 75 United States, previous studies exploring antimicrobial susceptibility of Vibrio vulnificus and V. parahaemolyticus have been conducted in South Carolina and the Gulf region (Han et al. 2007, Baker-Austin et al. 2008, Baker-Austin et al. 2009). However, to our knowledge, no studies have been completed in the Chesapeake Bay, which lies in a watershed where 17 million people work, live and play. Previous work has demonstrated that human recreational exposures to V. vulnificus and V. parahaemolyticus in the Chesapeake Bay are at significant enough concentrations to potentially elicit deleterious health effects, particularly among immunocompromised recreationists (Shaw et al. 2011). Moreover, current models predict that total tissue loading of shellfish and finfish with V. vulnificus and V. parahaemolyticus is associated not only with surface water concentrations but also with the risk of illness for those consuming contaminated seafood products (CFSAN (Center for Food Safety and Applied Nutrition) 2005, World Health Organization. & Food and Agriculture Organization of the United Nations 2005, 2011). Given these data, along with the knowledge that environmental conditions may be increasingly more favorable for Vibrio growth (Baker-Austin et al. 2012), it is not surprising that rates of Vibrio infections are increasing in Maryland and other U.S. states (Scallan et al. 2011a). In this context, it is critical to gain a better understanding of the antimicrobial susceptibility patterns of V. vulnificus and V. parahaemolyticus originating from estuarine-marine environments. This study evaluated the degree to which V. vulnificus and V. parahaemolyticus isolates from the Chesapeake Bay were susceptible to a broad range of antimicrobial treatments, and our findings provide the first data on antimicrobial resistance patterns 76 among Vibrio bacteria isolated from this region. These data will be helpful in short- and long-term predictions of human health risks associated with exposures to Vibrio populations within the Chesapeake Bay. Materials and Methods Sampling sites Sampling sites were selected based on their importance for human use in the Chesapeake Bay, Maryland Coastal Bays region. Two sites, Sandy Point State Park and St. Martin?s River, were characterized by frequent recreational use, and one site, the Pocomoke Sound, was characterized by heavy commercial fishing use (Figure 1). Sandy Point State Park is an artificial beach on the western shore of the Chesapeake mid-Bay region, at the base of the Chesapeake Bay Bridge. It is open year round and frequented by approximately 768,000 visitors annually, with many users frequenting park beaches during the summer (Sandy Point Park staff, Maryland Department of Natural Resources, personal communication). St. Martin?s River is a tributary of the Maryland Coastal Bays with approximately 10,000 residents. Land-use in the St. Martin?s River watershed is ~10% residential, ~48% agricultural, and ~34% forested (Thomas et al. 2009). The Pocomoke Sound is a major embayment of the Chesapeake Bay?s Eastern Shore. It is influenced by agricultural practices, including high-density concentrated poultry feeding operations, and is a popular destination for commercial and recreational fishing. Sample collection Sampling dates were chosen to coincide with times of high recreational and/or commercial use. Surface water samples (n=9) were collected during summer 2009, once a month, for three consecutive months (July, August, September) within two hours of 77 high tide and on approximately the same date each month. Water samples were collected just below the surface in sterile wide mouth polyproylene 1 L environmental sampling bottles (Nalgene Thermo Scientific, Waltham, MA). Bottles were rinsed three times with surface water and then dipped below the surface for a final 1 L collection volume. Samples collected for Vibrio culture were kept in insulated coolers, while water samples for Enterococci were stored in an insulated container on ice (4?C) upon collection and returned to the laboratory within four hours. Physical and chemical water quality measurements Water-column depth and surface-water salinity, temperature, dissolved oxygen, conductivity, and pH were measured on every sampling date and location with a YSI 556 Multi-probe system (YSI Incorporated, Yellow Springs, OH) in accordance with the manufacturer?s instructions. Fecal indicator measurements Fecal-indicator measurements were conducted following the standard methods as described for Enterococci in Standard Methods for the Examination of Water and Wastewater (Eaton et al. 1998). Briefly, surface-water samples were filtered in triplicate onto sterile 0.45 um pore size, 47 mm diameter, gridded membrane filters, and plated onto DifcoTM m Enterococcus (BD, Franklin Lakes, NJ) agar. Plates were incubated for 48 hours at 35?C. All light to dark red colonies were recorded as presumptive Enterococci. Vibrio isolation Surface water samples (100uL) were spread plated in triplicate onto Chromagar Vibrio media (DRG International, Mountainside, NJ) and incubated for 24 hours at 37?C. 78 After incubation, each plate was observed for characteristically colored bacterial colonies associated with V. vulnificus (turquoise) or V. parahaemolyticus (mauve). As V. vulnificus and V. cholerae both appear as turquoise colonies on Chromagar Vibrio media, all turquoise colonies were replated onto cellobiose-collistin (CC) agar (FDA 2004) media to confirm V. vulnificus species. The CC agar cultures were incubated for 24 hours at 37?C and yellow-colored colonies were considered presumptive V. vulnificus. Tryptic soy broth (TSB), supplemented with 5% sodium chloride, was then inoculated with individual colonies of V. vulnificus or V. parahaemolyticus and incubated at 37?C for 24 hours and stored in 30% glycerol stock at -80?C. Vibrio species confirmation A DNA template was obtained by producing crude cell lysates by boiling 1 mL aliquots of TSB cultures in 2 mL micro-centrifuge tubes at 100?C for 10 min. A Bio-rad CFX96 Touch? Real-Time PCR Detection System (Bio-rad, Hercules, CA, USA) was used to confirm the species of isolates with primers designed to detect Vibrio vulnificus (Panicker & Bej 2005) or Vibrio parahaemolyticus (Nordstrom et al. 2007). Following initial detection, samples testing positive for either species were subjected to further PCR testing for virulence genes (V. vulnificus: virulence correlated gene (vcg) (Baker-Austin 2010); V. parahaemolyticus: thermostable direct hemolysin (tdh), and thermostable related hemolysin (trh) genes (Nordstrom et al. 2007)). Real-time PCR was performed by using 1X PCR Buffer (Qiagen, Valencia, CA), (Qiagen), 0.2mM dNTP?s solution (Qiagen), 1X Q solution (Qiagen), 2.25U TopTaq DNA polymerase (Qiagen), 75nM internal control primers (each), 150nM internal control probe, 2uL internal control DNA, target primer and probe concentrations as detailed in 79 Table 1, and 3 uL DNA template per reaction, with the exception of the Vv vcg assay, where 5uL of DNA template was used and the internal control components were absent. DNase-RNase free water was added in a quantity sufficient for a 25uL total reaction volume. Two-stage qPCR cycling parameters are presented in Table 1. A linear synthetic exogenous DNA internal control, including a primer set, probe and internal control DNA, was incorporated simultaneously into each assay, excluding V. vulnificus vcg, to test for the presence and influence of inhibitors (Nordstrom et al., 2007). The following positive controls were used in each qPCR: Vibrio parahaemolyticus USFDA TX2103 and Vibrio vulnificus ATCC 27562. A randomly chosen subset of bacterial isolates were taxonomically identified with 16S rRNA gene sequences. DNA extracted from cultures was PCR amplified with bacteria-specific primers 27f (5?-AGAGTTTGATCCTGGCTCAG-3?) and 907r (5?- CCGTCAATTCCTTTRAGTTT-3?) using the following conditions: 94?C for 2 min, followed by 25 cycles of 55?C for 30 s, 72?C for 30 s, and 94?C for 2 min, followed by 72?C for 5 min. The PCR products were sequenced bi-directionally using the same primers on an ABI 3730 XL Genetic Analyzer in the BioAnalytical Services Laboratory at the University of Maryland Center for Environmental Science. Paired reads for each organism were analyzed and assembled with Phred and Phrap (Ewing & Green 1998, Ewing et al. 1998), manually edited with Consed (Gordon et al. 1998), and aligned and analyzed with the ARB sequence alignment program (Ludwig et al. 2004). 80 Clinical isolates Vibrio parahaemolyticus (n=8) were graciously provided by the State of Maryland?s Department of Health and Mental Hygiene for comparison purposes with our environmental isolates. Sample type and source of infection are presented in Table 2. Antimicrobial susceptibility testing Antimicrobial susceptibility testing was performed using the Sensititre? microbroth dilution system (Trek Diagnostic Systems, Westlake, Ohio) in accordance with the manufacturer?s instructions on all PCR-confirmed V. vulnificus (n=120 (3 vcg+)) and V. parahaemolyticus (n=77 (1 tdh+/ 1 trh+)). Cultures were grown overnight on tryptic soy agar (TSA) + 2.5% NaCl plates at 37?C. Vibrio cultures were transferred to sterile demineralized 2.5% saline solution to achieve a 0.5 McFarland standard. Then, 100 ?L of each suspension was transferred to sterile cation-adjusted Mueller Hinton broth (Trek Diagnostic Systems, Westlake, Ohio), and 50 ?L of the broth solution was dispensed into CML1FMAR custom minimal inhibitory concentration (MIC) plates (Trek Diagnostic Systems Inc.) with the following twenty-six antibiotics (range of concentrations in ?g/mL): amikacin (AMI; 8-64), ampicillin (AMP; 4-32), ampicillin- sulbactam 2:1 (A/S2; 8/4-32/16), apramycin (APR; 8-32), cefoxitin (FOX; 8-32), ceftriaxone (AXO; 8-64), cephalothin (CEP; 8-128), chloramphenicol (CHL; 8-32), ciprofloxacin (CIP; 1-4), oflaxacin (OFL; 1-8), ceftazidime (TAZ; 8-32), cefepime (FEP; 8-32), cefotaxime (FOT; 8-64), meropenem (MERO; 2-16), doxycycline (DOX; 2-16), imipenem (IMI; 2-16), levofloxacin (LEVO; 2-8), cefuroxime (FUR; 8-32), trimethoprim-sulfamethoxazole (SXT; 2/38-4/76), penicillin (PEN; 16-128), piperacillin (PIP; 16-128); piperacillin-tazobactam (P/T4; 16/4-128/4), streptomycin (STR; 8-128), 81 tetracycline (TET; 4-32), gentamicin (GEN; 2-16), amox/clav 2:1(AUG2; 8/4-32/16). Escherichia coli ATCC 25922 and E.coli ATCC 35218 were used as quality control strains. Next, MICs were recorded as the lowest concentration of an antimicrobial that completely inhibited bacterial growth (CLSI 2010). Resistance breakpoints published by the Clinical and Laboratory Standards Institute were used (CLSI 2010a,b). Breakpoints not available from CLSI (streptomycin, apramycin, penicillin) were derived from ranges used in similar studies (Chiew et al. 1998, Baker-Austin et al. 2008, Baker-Austin et al. 2009, Vizcaino et al. 2010). Multidrug resistance (MDR) was defined as resistance to two or more antibiotics. Statistical analyses Descriptive and inferential statistics were used to compare the percentage of isolates demonstrating intermediate resistance or resistance to tested antibiotics at each sampling site and sampled month, as well as the average number of antibiotics that V. vulnificus and V. parahaemolyticus isolates were resistant to at each sampling location and each month. P-values of ? 0.05 were defined as statistically significant. Due to the violation of normality assumptions, nonparametric Friedman two-way Analysis of Variance (ANOVA) was used to determine effects related to sampling site and month sampled. For samples in which there was a significant month effect, stratified Kruskal- Wallis one-way ANOVA and pairwise post-hoc tests were conducted for each month separately. Kruskal-Wallis analysis was conducted to evaluate differences in the occurrence of antimicrobial susceptibility in non-virulent and virulent bacteria. All statistical analyses were performed using StataIC 12 (StatCorp LP, College Station, TX). Results 82 Physical, chemical and bacterial water quality Water temperature, pH, and dissolved oxygen (DO) were uniform across the three sampling locations (Table 3). Average salinity in St. Martin?s River (24.5 ppt) was approximately double that of the Pocomoke Sound (10.5 ppt) and Sandy Point State Park (9.4 ppt) sampling site. Water depth at the Pocomoke Sound was approximately double that of Sandy Point State Park and three to four-fold deeper than St. Martin?s River. Enterococci counts (colony forming units (CFU)) per 100 mL-1 were uniformly low at Sandy Point during each sampling time point and below the single sample regulatory closure level of 104 CFU per 100 mL-1 (COMAR 2013). On one sampling occasion (Table 3), St. Martin?s River (August) Enterococci counts exceeded closure levels. Presumptive Vibrio colonies isolated during the culture portion of this study indicated that Vibrio vulnificus and V. parahaemolyticus were present in all tested water samples (Table 3). 120 V. vulnificus and 77 V. parahaemolyticus were purified, confirmed via PCR and tested for antimicrobial susceptibility. Species and virulence identification Sequence analysis (16S rRNA) of a selected subset of tested Vibrio isolates confirmed all isolates (Figure 2), except in the instance of two isolates where sequences were more similar to Photobacterium damselae. Due to the fact that P. damselae (Daniels & Shafaie 2000) has been indicated in human illness much akin to the infections caused by V. vulnificus and V. parahaemolyticus (Daniels & Shafaie 2000), these isolates were kept in the study. Virulence testing of all isolates identified three V. vulnificus isolates as positive for vcg, one V. parahaemolyticus isolate for tdh, and one isolate for 83 trh. Prevalence of antimicrobial resistance in V. vulnificus. All tested Vibrio vulnificus isolates (n=120) were susceptible to 14 of the 26 antibiotics tested, including the following drug classes that are important for the treatment of Vibrio infections and antimicrobials recommended by CDC (Centers for Disease Control and Prevention ) for the treatment of V. vulnificus infections (denoted in Table 4, Figure 3): tetracyclines, quinolones, and folate pathway inhibitors. In regard to CDC recommended antimicrobial agents, 2% of the tested isolates exhibited intermediate resistance against ceftazidime, a 3rd generation cephalosporin. Within the aminoglycoside class of antibiotics, isolates exhibited resistance to apramycin (1%) and streptomycin (4%). Intermediate resistance was expressed against amikacin (1%), apramycin (5%) and streptomycin (8%). Gentamicin was the only tested aminoglycoside to which all V. vulnificus isolates were completely susceptible. The aminoglycoside, streptomycin, was associated with the highest percentage of resistance (7% of all tested isolates) and second highest percentage of intermediate resistance (17% of all tested isolates) out of all of the antimicrobials tested. Isolates displayed the highest percentage of intermediate resistance (78% of all isolates) to chloramphenicol. Antimicrobial resistance in vcg+ V. vulnificus Of the three isolates positive for the virulence correlated gene (vcg), none displayed resistance to any of the tested antibiotics, but all three expressed intermediate resistance (100%) to chloramphenicol. Prevalence of antimicrobial resistance in V. parahaemolyticus All tested Vibrio parahaemolyticus isolates were susceptible to 11 of the 26 tested 84 antibiotics and four (carbapenems, tetracyclines, quinolones folate pathway inhibitors) of the eight tested antimicrobial classes (Table 4, Figure 3). Conversely, 96% of isolates had intermediate resistance to chloramphenicol, followed by ampicillin (25%), cephalothin (17%), penicillin (16%) and cefuroxime sodium (14%). A high percentage of isolate resistance was seen within the penicillin class (penicillin (68%); ampicillin (53%)), while a low percentage of resistance was detected in piperacillin (4%) and streptomycin (4%). Antimicrobial resistance in tdh/trh+ V. parahaemolyticus One isolate was tdh+ and one isolate was trh+. Both virulent isolates exhibited multiple resistance and intermediate resistance patterns. The trh+ V. parahaemolyticus isolate was resistant to ampicillin and penicillin and expressed intermediate resistance to chloramphenicol. The tdh+ V. parahaemolyticus was resistant to ampicillin, ampicillin- sulbactam, penicillin, piperacillin-tazobactam, and amoxicillin-clavulanic acid and expressed intermediate resistance to chloramphenicol. Impact of sampling site and month on antimicrobial resistance Friedman two-way ANOVA Based on the Friedman two-way ANOVA, there were significant month effects for V. parahaemolyticus expressing antibiotic resistance (P<0.0001) and intermediate resistance (P<0.0001); and for V. vulnificus expressing resistance (P=0.0008) and intermediate resistance (P=0.0098). After adjusting for the repeated measures over time (month), site effects were significant for V. vulnificus expressing antibiotic resistance (P=0.0321) and intermediate resistance (P=0.0029), but not significant for V. parahaemolyticus expressing resistance (P=0.6133) and intermediate resistance 85 (P=0.7660). Kruskal-Wallis one-way ANOVA As there was a significant month effect in the Friedman two-way ANOVA for both V. vulnificus and V. parahaemolyticus isolates expressing antibiotic resistance and intermediate resistance, stratified Kruskal-Wallis one-way ANOVA and pairwise post- hoc tests were conducted on the site difference for each month separately. Analysis with Kruskal-Wallis showed no significant difference between sites by month for V. vulnificus or V. parahaemolyticus expressing resistance (July, August, September; (P=0.5340, 0.2801, 0.4966); (P=0.7246, 0.9448, 0.6809), respectively) or V. parahaemolyticus expressing intermediate resistance (P=0.5959, 0.8046, 0.2135). After testing V. vulnificus expressing intermediate resistance for site differences by each month separately, it was determined that there was a significant site effect only in July (P=0.035). Post-hoc testing clarified that the site, St. Martin?s River, was different from Sandy Point during the month of July, with reduced intermediate resistance for each V. vulnificus isolate recovered from St. Martin?s River (P=0.0166). Kruskal-Wallis one-way ANOVA further elucidated that there was no significant difference in the median intermediate resistance or resistance patterns during the sampling period when St. Martin?s River (August) (P=0.44) had higher levels of bacterial-indicator species. Clinical V. parahaemolyticus Clinical isolates tested displayed comparable resistance profiles to environmental isolates tested (Table 4). Environmental isolates demonstrated intermediate resistance and resistance to a greater range of antibiotics (15 antibiotics in 4 classes) when 86 compared to clinical isolates (5 antibiotics in 3 classes). However, based on analyses with two-sample proportion tests, the overall percentage of resistance and intermediate resistance (%= number of antimicrobials demonstrating resistance/ number of total antimicrobials tested) in clinical isolates was not statistically different (P=0.511, 0.430; respectively) from environmental isolates. 87 Discussion Treatability of Chesapeake Bay related Vibrio illness in Maryland Vibrio vulnificus and V. parahaemolyticus are the causative agents for wound infections, primary septicemia, and gastroenteritis related to seafood and seawater exposure (CDC 2012a). While gastroenteritis does not typically necessitate antibiotic treatment, it is required for wound infection and primary septicemia caused by both Vibrio species tested in this study. Most isolates tested in this study were susceptible to the antimicrobial agents recommended by the CDC. Treatment recommendations for such infections include tetracyclines (doxycycline, tetracycline), flouroquinolones (ciprofloxacin, levofloxacin), third-generation cephalosporins (cefotaxime, ceftazidime, ceftriaxone), aminoglycosides (amikacin, apramycin, gentamicin, streptomycin) and folate pathway inhibitors (trimethoprim-sulfamethoxazole) (Daniels & Shafaie 2000, CDC 2009). The CDC recommends a treatment course of doxycycline (100 mg PO/IV twice a day for 7-14 days) and a third-generation cephalosporin (e.g.,ceftazidime 1-2 g IV/IM every eight hours), although they state that single agent regimens employing a fluoroquinolone has been reported to be at least as effective in an animal model as combination drug regimens with doxycycline and a cephalosporin (CDC 2009). Most isolates tested in this study were susceptible to the antimicrobial agents recommended by the CDC. All tested V. vulnificus isolates were susceptible to third and fourth generation cephalosporins, although two V. parahaemolyticus isolates (3%) demonstrated intermediate resistance to cefotaxime, a 3rd generation cephalosporin and two isolates demonstrated a degree of resistance to cefepime, a 4th generation cephalosporin. While isolate intermediate resistance and resistance was relatively low for 88 the newer generation cephalosporins, these antibiotics are considered to be some of the best defenses against the dangerous infections that these organisms can elicit (CDC 2009). Due to the contraindication of doxycycline and fluoroquinolones in children, a combination of trimethoprim-sulfamethoxazole and an aminoglycoside antibiotic is recommended (CDC 2009). Given that three of the four tested aminoglycosides (amikacin, apramycin, streptomycin) were associated with intermediate resistance or resistance (e.g., streptomycin intermediate resistance and resistance in V. vulnificus: 17%, 7%, respectively; V. parahaemolyticus: 8%, 4%, respectively) in a subset of isolates, this may be a resistance pattern of concern. Conversely, for the aminoglycoside, gentamicin, all tested isolates were fully susceptible. Based on these data, it is feasible that only one aminoglycoside, gentamicin, could be administered with full confidence in its ability to fight Vibrio infections contracted by children recreating in the Chesapeake Bay. While the detection of virulence genes was very low, each gene was present in at least one tested isolate (Table 6A). Resistance patterns among these virulent isolates were similar to the patterns seen in non-virulent isolates. Due to the very limited number of environmental isolates possessing virulence markers, it is not possible to say that most virulent strains would behave as the non-virulent environmental isolates, but the susceptibilities of the clinical and virulent isolates was similar enough to the non-virulent isolates in this study that expecting similar patterns would not be unfounded. Antimicrobial susceptibility as compared to fecal indicator measurements A range of Enterococci counts were observed over the course of this study, although most studied locations were within the range of acceptable water quality for 89 recreation on each sampling date. For the sake of this study, it can be gleaned that indicator bacterial water quality was not a major determinant of levels of antibiotic resistance in the studied environments. During the one instance that the geometric mean of Enterococci was higher than regulation limits, there was no discernable difference in antimicrobial susceptibility patterns of isolates originating from that site (St. Martin?s ? August). This is counter to patterns seen in similar studies, during which antimicrobial resistance was elevated at sites contaminated with fecal waste of humans (de Oliveira & Pinhata 2008) and animals (Sapkota et al. 2007). Comparison to other U.S. studies of V. vulnificus and V. parahaemolyticus antimicrobial susceptibility Results of this study were comparable to a similar study conducted on Vibrio isolated from Gulf Coast oysters in Louisiana (Han et al. 2007). The previous study, conducted in 2005-2006, also found a higher resistance profile in V. parahaemolyticus than V. vulnificus. In addition, ampicillin was the only tested antimicrobial in the Gulf Coast study to which a large percentage of V. parahaemolyticus isolates demonstrated intermediate resistance to resistance (~81% of all tested isolates). This trend was seen as early as the 1970s in a study that tested resistance of V. parahaemolyticus to ampicillin and ?-lactamase inhibitors (Joseph et al. 1978), where over 90% of isolates were found to be resistant to ampicillin. In contrast to the present study, the Gulf Coast study (Han et al. 2007) found no resistance in either Vibrio species to chloramphenicol, cefotaxime, or ceftazidime, while we observed intermediate resistance among a subset of V. vulnificus and V. parahaemolyticus against these antimicrobial agents (78/96%, 0/3%, 2/0%, (Vv/Vp), respectively). 90 Our findings are also in partial agreement with two large studies of V. vulnificus and V. parahaemolyticus isolates originating from the Georgia and South Carolina coastline of the United States (Baker-Austin et al. 2008, Baker-Austin et al. 2009). While the Chesapeake Bay isolates did not show the same high degree of prevalence of antimicrobial resistance, the antimicrobial agents to which isolates showed resistance were similar (i.e., amoxicillin, apramycin, penicillin and streptomycin for V. parahaemolyticus). Vibrio vulnificus isolates demonstrated similar resistance profiles, especially in regard to percent intermediate resistant and resistant to the penicillin class and cefoxitin. Baker-Austin et al. (2009) reported much higher percent intermediate resistance and resistance in V. vulnificus to apramycin and streptomycin as compared to this study. In addition, key antimicrobials to which V. parahaemolyticus isolates from Georgia and South Carolina displayed full susceptibility were also found to have identical susceptibilities in this study (i.e., ceftriaxone, ciprofloxacin, imipenem, ofloxacin, meropenem, tetracycline), except in the case of chloramphenicol, for which no or low (V. vulnificus) resistance was observed in the Georgia and South Carolina study. In contrast to this study, Baker-Austin et al. (2009) found only one V. vulnificus isolate to be completely susceptible to all antimicrobials tested, while this study found 15 (12.5%) isolates fully susceptible. Study sites and influences of pollution Each studied site has a history of water pollution. Sandy Point State Park has historically been the site of low bacteriological water quality and is adjacent to the Magothy River, a site where there have been numerous wastewater treatment overflows. The Pocomoke River is located adjacent to many farming operations, including poultry 91 concentrated animal feeding operations (CAFOs), which may increase the introduction of antimicrobial residues to the waterway due to runoff of fecal mater contaminated with antimicrobials used in animal husbandry (Campagnolo et al. 2002). Finally, St. Martin?s River is adjacent to many homes on septic systems, notorious for leakage (Jones et al. 2004). While each of the studied sites has a history of contamination that may increase the incidence of antimicrobial residues and associated changes in resident bacteria in the estuarine environment, this study only detected a small difference with regard to levels of antibiotic resistance between St. Martin?s River and Sandy Point during the month of July in V. vulnificus expressing intermediate resistance. These results are unlike the high levels of resistance seen in other studies (Baker-Austin et al. 2008, Baker-Austin et al. 2009), perhaps due to differences in contamination sources (i.e., heavy metal pollution in Baker-Austin et al. (2009)), although a significant difference in antibiotic resistance was not detected between pristine sites and those with heavy metal contamination in that study. Conclusions This study represents the first investigation of antimicrobial susceptibility of Vibrio species recovered from the Chesapeake Bay and provides a baseline against which future studies can be compared to determine whether susceptibilities change over time. Isolates tested in this study displayed high intermediate resistance to chloramphenicol, when compared to similar studies. Isolate intermediate resistance and resistance to aminoglycosides is of note concerning the treatment of pediatric Vibrio illness originating from the Chesapeake waters or seafood. Low-level intermediate resistance and resistance to third and fourth generation cephalosporins is also of interest with regard to treatment 92 effectiveness and should be monitored. Consensus with previous studies was reached in terms of the prevalence of intermediate resistance and resistance to the penicillin class of antimicrobials. As most of the antimicrobial agents recommended for treatment of Vibrio illness by CDC were fully effective against V. vulnificus and V. parahaemolyticus isolated from the Chesapeake Bay, treating infections contracted from the Bay, at least in adults, should not be problematic. Treatment of pediatric illnesses should gravitate towards the use of trimethoprim-sulfamethoxazole and the aminoglycoside, gentamicin, which was the only aminoglycoside 100% effective against Vibrios recovered in this study. 93 Table 1. PCR conditions for the detection of V. vulnificus and V. parahaemolyticus virulence genes. Primer Primer (forward & reverse)/Probe Concentrations (nM) PCR conditions Vibrio vulnificus/vvh 400/240 1x: 95 ?C for 60 s; 41x: 95 ?C for 5 s, 59 C for 45 s Vibrio vulnficus/vcg 250/180 1x: 95?C for 10 m; 40x: 95?C for 15 s, 60?C for 90 s Vibrio parahaemolyticus/tlh 200/150 1x: 95?C for 10 m; 45x: 95?C for 5 s, 66?C for 45 s Vibrio parahaemoylticus/tdh trh 200/75 1x: 95?C for 60 s; 50x: 95?C for 5 s, 59?C for 45 s 94 Table 2. Clinical isolates provided by Maryland Department of Health and Mental Hygiene. Sample type, infection source and associated, if any, antimicrobial resistance. Clinical isolate Sample source Infection source AST results (# antibiotics) Intermediate resistance; Resistance 1 Stool Undercooked seafood ampicillin,penicillin; 0 2 Stool Undercooked seafood chloramphenicol; ampicillin, penicillin 3 Stool No data available Chloramphenicol, penicillin; 0 4 Stool Undercooked seafood Chloramphenicol, apramycin, streptomycin; ampicillin, penicillin 5 Stool Undercooked seafood chloramphenicol ; 0 6 Stool No data available Chloramphenicol, ampicillin; penicillin 7 No data available Beach, unknown location Chloramphenicol, ampicillin, penicillin; 0 8 Wound No data available chloramphenicol ; ampicillin, penicillin 95 Table 3. Physical, chemical and bacterial water quality. Site Date Salinity Temp. (?C) pH Dissolved oxygen (mg L-1) Depth (feet) Average Enterococcus geometric mean CFU 100mL-1 (+/- std. dev.) Average Vibrio vulnificus CFU mL-1 (+/- std. dev.) Average Vibrio parahaemolyti cus CFU mL-1 (+/- std. dev.) Pocomoke 16-Jul 09 10.5 26.1 7.6 n/a 15.6 24 (8) 51 (41) 13 (9) Pocomoke 18-Aug-09 10.0 28.8 7.4 4.9 14.4 15 (10) 35 (29) 8 (9) Pocomoke 21-Sep-09 11.1 22.6 7.3 6.3 13.8 38 (6) 52 (40) 9 (10) Sandy Point 9-Jul-09 8.6 24.5 8.3 7.4 7.6 2 (3) 204 (137) 11 (23) Sandy Point 3-Aug-09 10.0 26.5 8.0 7.0 7.6 5 (4) 234 (76) 19 (15) Sandy Point 3-Sep-09 9.6 24.6 7.8 7.1 7.6 2 (3) 294 (71) 18 (11) St. Martin?s 6-Jul-09 24.5 25.9 7.9 6.6 4.4 3 (7) 28 (46) 17 (20) St. Martin?s 9-Aug-09 23.4 26.5 7.8 5.6 4.8 365 (6) 122 (47) 48 (40) St. Martin?s 6-Sep-09 25.5 23.1 7.5 2.9 4.9 3 (5) 32 (24) 12 (12) 96 Table 4. Antimicrobial intermediate resistance and resistance, for respective number and percent, denoted for antibiotic class and specific antibiotic. 97 Table 5. Comparison of environmental and clinical isolates and their respective associated antimicrobial resistance to a subset of antibiotics to which highest resistance within tested isolates was displayed. Environmental Isolates Clinical Isolates Antibiotic Intermediate n (%) Resistant n (%) Intermediate n (%) Resistant n (%) Ampicillin 19 (25) 40 (53) 1 (12.5) 2 (25) Apramycin 4 (5) 1 (1) 1 (12.5) 0 (0) Streptomycin 6 (8) 3 (4) 1 (12.5) 0 (0) Chloramphenicol 74 (96) 0 (0) 7 (87.5) 0 (0) Penicillin 12 (16) 52 (68) 3 (37.5) 4 (50) 98 Table 6. Antibiotic resistance (AR) and multiple antibiotic resistance (MAR) by virulence factors (6A), site (6B) and month (6C). 6A. Vv vcg+ (n=3) Vv vcg ? (n=117) Vp tdh+ (n=1) Vp trh+ (n=1) Vp tdh/trh- (n=75) AR MAR AR MAR AR MAR AR MAR AR MAR Resistant 0 (0%) 0 (0%) 21 (18%) 5 (4%) 1 (100%) 1 (100%) 1 (100%) 1 (100%) 51 (68%) 39 (52%) Intermediate resistance 3 (100%) 0 (0%) 101 (86%) 29 (25%) 1 (100%) 1 (100%) 1 (100%) 1 (100%) 74 (99%) 44 (59%) 6B. Pocomoke St. Martin?s Sandy Point (n=44) (n=11) (n=65) V. vulnificus AR MAR AR MAR AR MAR Resistant 10 (23%) 3 (7%) 0 (0%) 0 (0%) 12 (18%) 2 (3%) Intermediate resistance 42 (95%) 11 (25%) 6 (55%) 0 (0%) 58 (89%) 18 (28%) (n=14) (n=29) (n=34) V. parahaemolyticus AR MAR AR MAR AR MAR Resistant 10 (71%) 7 (50%) 22 (76%) 17 (59%) 22 (65%) 17 (50%) Intermediate resistance 14 (100%) 8 (57%) 28 (97%) 15 (52%) 34 (100%) 23 (68%) 6C. July August September (n=40) (n=47) (n=33) V. vulnificus AR MAR AR MAR AR MAR Resistant 3 (8%) 0 (0%) 13 (28%) 4 (9%) 6 (18%) 1 (3%) Intermediate resistance 32 (80%) 4 (10%) 42 (89%) 16 (34%) 30 (91%) 9 (27%) (n=11) (n=40) (n=26) V. parahaemolyticus AR MAR AR MAR AR MAR Resistant 9 (82%) 8 (73%) 31 (78%) 22 (55%) 14 (54%) 10 (38%) Intermediate resistance 11 (100%) 7 (64%) 39 (98%) 24 (60%) 26 (100%) 14 (54%) 99 Figure 1. Sampling sites. (Tracey Saxby, Kate Boicourt, Integration and Application Network, University of Maryland Center for Environmental Science (ian.umces.edu/imagelibrary/ displayimage-127-5815.html) 100 Figure 2. 16S rRNA sequencing analysis of a subset of Vibrio isolates tested. 101 Figure 3. Number of antibiotics against which Vibrio isolates expressed resistance or intermediate resistance. 102 CHAPTER 5: CONCLUSIONS 103 This dissertation work was designed to answer pressing questions concerning potentially pathogenic Vibrio species of bacteria in the Chesapeake Bay. Recently, public health managers in the Chesapeake Bay region have been armed with models for predicting surface water concentrations of V. vulnificus and V. parahaemolyticus, but it is difficult to know how, or if, that information should translate to policy changes in closures of beaches or shellfish harvest areas in the Chesapeake Bay. Additionally, it has been proposed to make these predictions available to the general public, to interpret the data based on their own depth of knowledge. These interpretations carry great risk not only for the public?s perception of the safety of their environment but also for the associated economic risk if people improperly interpret predictions to mean that water and seafood products are unsafe. It would be irresponsible to share these data without the appropriate framework in which to approximate relative risk of infection. As similar predictive models are developed for other estuaries and coastal regions, it will not be surprising to find regional managers and scientists faced with this same problem regarding translation of predictive models. Portions of this dissertation research were undertaken to begin to answer the basic question: ?What do the predictions of Vibrio abundance mean for human health?? The first goal of this dissertation was to quantify exposure of humans to potentially pathogenic Vibrio cells when swimming. Exposure assessment swim studies were undertaken to calculate the first estimates of how many Vibrio bacterial cells a person may come into contact with and subsequently contract while recreating in the Chesapeake. Results were clear that during the months when surface water is host to an elevated abundance of Vibrio cells, a person recreating or working in those waters could 104 expect a significant dermal exposure. If that user has any abrasions or wounds, it is more possible that they are exposed to a number of potentially pathogenic cells. The dose of V. vulnificus or V. parahaemolyticus needed to cause an infection in a healthy, moreover an immunocompromised, individual is poorly understood, particularly for non-consumption exposure. Future research will aim to take these first exposure estimates and interpret them in terms of a quantitative microbial risk assessment. Additionally, these estimations will be paired with calculations of climate change related increases in surface water concentrations of Vibrio. The second goal of this dissertation was to determine if storm events, especially those causing wind-driven resuspension of sediments, are important mechanisms of Vibrio introduction to the water column and to aquacultured oysters. Because the increase in aquacultured oyster operations will likely result in a larger summer harvest of oysters, which are typically consumed raw on the half shell, understanding how oyster concentrations of Vibrio might be impacted is vital. One relevant question posed by the aquaculture community is whether Vibrio concentrations in oysters differ based on their position in the water column. This study found no difference in Vibrio concentrations between oysters collected from the bottom and the top of the water column. The sampling location may have been too shallow to see a true difference in surface versus bottom culture, but since much of the Chesapeake is shallow, and most aquaculture operations will likely be near-shore, this was probably a good indicator site. Our study identified two storm-induced events ? wind-driven resuspension and flushing due to heavy precipitation. While the wind likely drove Vibrio into the water column from the sediment, and into contact with oysters, the subsequent rain event likely 105 caused a flushing effect, which may have diluted Vibrio cell concentrations at the aquaculture site. An inverse relationship between in-oyster concentrations and surface- water concentrations of Vibrio was observed, suggesting a dynamic relationship between oysters and Vibrio in the water column of an aquaculture facility in which rates of oyster filtration alter the water column concentration of Vibrio. The pattern observed was potentially caused by changes in the rate of oyster filtration during and directly after the storm, when suspended solids were likely high and oysters likely slowed or ceased filtration for a period of time. This reaction to high concentrations of suspended solids may prevent increases of in-oyster Vibrio, but this research should be repeated during more storm events to verify that prediction. Although not statistically significant due to the Most Probable Number variability between samples, an increase was detected in in-oyster concentrations of V. parahaemolyticus directly after the storm, but not of V. vulnificus. At the end of the study, in-oyster concentrations were approximately similar to the pre-storm concentrations, suggesting a possible ?ambient? concentration of summer Vibrio density. Further testing should be conducted to determine if these levels vary based on site, position in the water column, and after storm impacts. Ideally, further research would have the opportunity to sample before and after separate wind and precipitation events. Initially, this work was proposed for a full summer storm-period, with multiple events, but 2011 was a very quiet year in term of summer squalls. On the spectrum of storm intensity, the study fell on the high end of impacts with Hurricane Irene. Further inquiries should be repeated with measurements along a spectrum of storm intensity, not just the most intense. 106 At this time, it is inconclusive whether a storm event should trigger closure of a shellfish fishery. However, concentrations of Vibrio in oysters were very high throughout the study and the percentage of samples that were positive for virulent V. vulnificus and V. parahaemolyticus increased after the storm. Changes of in-oyster virulence is another area where further research would be beneficial in determining the public health risk of oyster consumption after a storm event. From the results of this study, it can be concluded that sampling sediment and surface water for general concentrations of Vibrio, or pathogenic subspecies, may not be enough to predict the concentrations in oysters. Additionally, climate change estimates of increases in surface water temperature, changes in salinity and intensity or frequency of storm events may also drive changes to shellfish management practices. Finally, this research addressed the ability to treat Vibrio infections contracted from exposure with Chesapeake Bay waters or seafood products. Environmental isolates from three areas known for use by recreationists and commercial fishermen were tested for their susceptibility to a wide range of antibiotic agents. Antibiotics were chosen not just for their clinical importance (i.e., CDC recommended antibiotics), but also to compare resistance patterns in relation to other studies conducted on Vibrio in other geographical areas of the United States. Overall, V. parahaemolyticus isolates from the Chesapeake Bay displayed more intermediate resistance and resistance to tested antimicrobial agents than V. vulnificus. Since approximately 34% of V. parahaemolyticus are wound infections (Daniels et al. 2000), this may be more troubling than if cases simply resulted in self-resolving gastroenteritis, due to the deleterious nature of such infections. Luckily, most CDC recommended antibiotic treatments for Vibrio 107 illness were effective in controlling growth of these bacteria, although there was some low-level intermediate resistance to 3rd and 4th generation cephems and moderate intermediate resistance and resistance to three of the four tested aminoglycosides. Chesapeake Bay isolates expressed high-level intermediate resistance to chloramphenicol, unlike the findings from other sites within the United States. Resistance patterns were not related to site contamination, as measured by fecal indicator bacteria, and only one site, St. Martin?s, had lower V. vulnificus intermediate resistance, as compared to Pocomoke Sound and Sandy Point, during the month of July. Overall, most antibiotics recommended by CDC would be expected to control Vibrio infection, but clinicians may need to consider gentamicin as the only aminoglycoside that was 100% effective against controlling Vibrio growth. Such information should be taken into consideration when treating pediatric patients, for whom a combination treatment of trimethoprim-sulfamethoxzaole and an aminoglycoside is recommended. In summary, the culmination of this work begins to answer some of the questions that have recently been asked by clinicians, research scientists and public health managers in the Chesapeake Bay region concerning pathogenic Vibrio. When exposed to typical summer season surface water concentrations of V. vulnificus and V. parahaemolyticus, an exposure should be expected. Storm-related changes in the aquatic environment will change the density of Vibrio in surface waters, but possibly not in simple increases and decreases of Vibrio concentrations. Finally, if a patient contracts a Vibrio illness from the Chesapeake Bay, it is likely that it can be controlled with recommended antimicrobial treatment regimes, if the clinician is properly informed and diagnosis is accurate. Clinicians treating patients in the Chesapeake Bay region should 108 be well-informed about the symptoms, proper diagnosis and treatment of Vibrio infections. Although these results are not the end-all conclusions needed to inform managers about decisions to take preventative action to control Vibrio illness, these data sets serve as useful starting points to direct the fine-tuning of questions and future research projects. 109 APPENDIX 110 Appendix: Microcosm Sediment Resuspension Experiment Specific Aims Laboratory based microcosm experiments were conducted to mimic a resuspension of sediment event, with two levels of resuspension (low, high) used to determine the approximate increase of V. vulnificus and V. parahaemolyticus in oysters following exposure to suspended sediment. Methods Three treatment levels of control (no sediment resuspension), low, and high resuspension were used to mimic the field environment. Three tanks (10 gallon) were used for each treatment. On October 10, 2011, a surface layer sediment sample was retrieved from the same area of the aquaculture site that was used for the environmental resuspension experiment. Sediment (550 g) was spread evenly on the bottom of each tank and ambient Choptank River water was slowly poured over sediment to minimize resuspension into water. Tanks were allowed to settle for 24 h in a climate controlled chamber heated to 32?C. Oxygen air lines were run to each tank, with an air stone anchoring the lines in the tank, above the sediment layer, and air was not turned on until the experiments were commenced to allow for full particle settlement in tanks prior to experiments. Chamber lights were used only during sample collection. On the morning of October 11, 2011 oysters were collected from the aquaculture site and placed on the bottom of the tank (36 oysters per tank) after 24 h. An initial representative oyster sample (homogenate of 6 oysters, as described in Chapter 3 Methods) was taken at this time to determine background Vibrio level in oysters. 111 Tank sediment was resuspended for 2 hours per treatment, excluding control tanks. Resuspension was achieved by placing tanks on wooden frames custom built to support tanks above stir plates. A large magnetic stir bar was added to each tank and stirring activity was set to achieve the desired level of relative resuspension (low and high). Water, total suspended solids (TSS), and oyster samples were taken at each sampling time point (before sediment suspension (T0), directly after suspension (T1), 24h (T2), 72h (T3), 7d (T4)) from each tank. Water samples were collected by filtering 180 mL of tank water and analyzed using qPCR as described in Chapter 3. Water for TSS (50 mL) was collected. Oysters were sampled (6 oysters per sample, per tank) and analyzed using MPN-PCR methods as described in Chapter 3. Results Average (? standard error) TSS for treatments were 227 (?5.43) mg L-1 (high) and 57 (? 10.6) mg L-1 (low) directly following resuspension treatments (Table 1). Treatments, including controls, had appreciably the same TSS averages at subsequent sampling time points. Oyster MPN g-1 V. parahaemolyticus increased from pre-treatment average of 46,798 to 466,140 (high),105,533 (low) and 496,597 (control) at T1 (Table 2). At T2, V. parahaemolyticus decreased to 77,373 (high), 22, 766 (low) and 147,588 (control) MPN g-1. High and control treatments reduced further at T3 (6,533 and 72,333, respectively), while the low treatment increased to 47,733 MPN g-1. At T4, all treatments had reduced to 3,000-5,000 MPN g-1. Water concentrations of V. parahaemolyticus were highest in control tanks (160 CFU mL-1) at T1, while the resuspension treatments averaged 111 (high) and 47 (low) CFU mL-1. All tanks had approximately the same CFU mL-1 V. 112 parahaemolyticus at T2-T4. Due to control V. parahaemolyticus MPN g-1 and CFU mL-1 being appreciably higher in concentration than the ?high? treatment, this experiment was deemed unsuccessful. Oyster MPN g-1 V. vulnificus increased from pre-treatment average of 459,052 to 1,002,621 (high),1,148,601 (low) and 834,673 (control) at T1 (Table 3). At T2, V. vulnificus decreased to 834,674 (high and low) and increased in the control treatment to 1,148,600 MPN g-1. High treatments reduced further at T3 (459,053), while the low and control treatments increased (low) or plateaued (control)to 1,148,600 MPN g-1. At T4, high treatments had increased to 1,148,600 MPN g-1, but decreased in low treatments (834,674 MPN g-1) . Water concentrations of V. vulnificus were highest in low treatment tanks (32,210 CFU mL-1) at T1, followed by the control and high resuspension treatments (6,843 (control), 1,310 (high) CFU mL-1). All tanks had approximately the same CFU mL-1 V. vulnificus at T2-T3, with a slight increase at T4 (5,000-8,000 CFU mL-1. Due to control V. vulnificus MPN g-1 and CFU mL-1 being appreciably the same concentration as high and low treatments, this experiment was deemed unsuccessful. 113 Table 1. Average Total Suspended Solids (mg L-1) Sample Date Treatment 1 - High Treatment 2 - Low Control T0 (11 OCT 11) 10 10 10 T1 (11 OCT 11) 227 57 19 T2 (12 OCT 11) 10 6 6 T3 (14 OCT 11) 6 5 4 T4 (17 OCT 11) 4 4 5 114 Table 2. Oyster MPN and water CFU mL-1 V. parahaemolyticus. Oysters Water Sample Date Treatment 1 - High (MPN g-1) Treatment 2 - Low (MPN g-1) Control (MPN g-1) Treatment 1 - High (CFU mL-1) Treatment 2 - Low (CFU mL-1) Control (CFU mL-1) T0 (11 OCT 11) - - 46,798 - - - T1 (11 OCT 11) 466,140 105,533 496,597 111 47 160 T2 (12 OCT 11) 77,373 22,766 147,588 35 39 17 T3 (14 OCT 11) 6,533 47,733 72,333 6 5 7 T4 (17 OCT 11) 3,000 5,066 5,066 5 4 2 115 Table 3. Oyster MPN and water CFU mL-1 V. vulnificus. Oysters Water Sample Date Treatment 1 - High (MPN g-1) Treatment 2 - Low (MPN g-1) Control (MPN g-1) Treatment 1 - High (CFU mL-1) Treatment 2 - Low (CFU mL-1) Control (CFU mL-1) T0 (11 OCT 11) - - 459,052 - - - T1 (11 OCT 11) 1,002,621 1,148,601 834,673 1,310 32,210 6,843 T2 (12 OCT 11) 834,674 834,674 1,148,600 1,774 2,474 1,488 T3 (14 OCT 11) 459,053 1,148,600 1,148,600 2,082 1,401 704 T4 (17 OCT 11) 1,148,600 834,674 1,148,600 5,164 8,473 7,042 116 References Akinbami LJ, Moorman JE, Bailey C, Zahran HS, King M, Johnson CA, Liu X (2012) Trends in asthma prevalence, health care use, and mortality in the United States, 2001-2010. NCHS Data Brief:1-8 Amirsheybani HR, Crecelius GM, Timothy NH, Pfeiffer M, Saggers GC, Manders EK (2001) The natural history of the growth of the hand: I. Hand area as a percentage of body surface area. Plast Reconstr Surg 107:726-733 Asplund ME, Rehnstam-Holm AS, Atnur V, Raghunath P, Saravanan V, Harnstrom K, Collin B, Karunasagar I, Godhe A (2011) Water column dynamics of Vibrio in relation to phytoplankton community composition and environmental conditions in a tropical coastal area. Environ Microbiol 13:2738-2751 Audemard C, Kator HI, Rhodes MW, Gallivan T, Erskine AJ, Leggett AT, Reece KS (2011) High salinity relay as a postharvest processing strategy to reduce vibrio vulnificus levels in Chesapeake Bay oysters (Crassostrea virginica). J Food Prot 74:1902-1907 Avila LA, Cangialosi J (2011) Tropical Cyclone Report: Hurricane Irene (AL092011), 21-28 August 2011. In. NOAA, National Hurricane Center Baker-Austin C, Anthony Gore, James D. Oliver, Rachel Rangdale, J Vaun McArthur, David N. Lees (2010) Rapid in situ detection of virulent Vibrio vulnificus strains in raw oyster matrices using real-time PCR Environmental Microbiology Reports 2:76-80 Baker-Austin C, McArthur JV, Lindell AH, Wright MS, Tuckfield RC, Gooch J, Warner L, Oliver J, Stepanauskas R (2009) Multi-site analysis reveals widespread antibiotic resistance in the marine pathogen Vibrio vulnificus. Microb Ecol 57:151-159 Baker-Austin C, McArthur JV, Tuckfield RC, Najarro M, Lindell AH, Gooch J, Stepanauskas R (2008) Antibiotic resistance in the shellfish pathogen Vibrio parahaemolyticus isolated from the coastal water and sediment of Georgia and South Carolina, USA. J Food Prot 71:2552-2558 Baker-Austin C, Stockley L, Rangdale R, Martinez-Urtaza J (2010) Environmental occurrence and clinical impact of Vibrio vulnificus and Vibrio parahaemolyticus: a European perspective. Environ Microbiol Rep 2:7-18 Baker-Austin C, Trinanes JA, Taylor NGH, Hartnell R, Siitonen A, Martinez-Urtaza J (2012) Emerging Vibrio risk at high latitudes in response to ocean warming. Nature Climate Change 117 Baquero F, Martinez JL, Canton R (2008) Antibiotics and antibiotic resistance in water environments. Curr Opin Biotechnol 19:260-265 Beardsley C, Pernthaler J, Wosniok W, Amann R (2003) Are readily culturable bacteria in coastal North Sea waters suppressed by selective grazing mortality? Appl Environ Microbiol 69:2624-2630 Bisharat N, Agmon V, Finkelstein R, Raz R, Ben-Dror G, Lerner L, Soboh S, Colodner R, Cameron DN, Wykstra DL, Swerdlow DL, Farmer JJ, 3rd (1999) Clinical, epidemiological, and microbiological features of Vibrio vulnificus biogroup 3 causing outbreaks of wound infection and bacteraemia in Israel. Israel Vibrio Study Group. Lancet 354:1421-1424 Boesch DF, Coles VJ, Kimmel DG, Miller WD (2007) Ramifications of climate change for Chesapeake Bay hypoxia, p. 54-70. In Regional Impacts of Climate Change: Four Case Studies in the United States. Pew Center on Global Climate Change, Arlington, VA. Bowdre JH, Poole MD, Oliver JD (1981) Edema and hemoconcentration in mice experimentally infected with Vibrio vulnificus. Infect Immun 32:1193-1199 Broberg CA, Calder TJ, Orth K (2011) Vibrio parahaemolyticus cell biology and pathogenicity determinants. Microbes Infect 13:992-1001 Brown C, Hood R, Long W, Jacobs J, Ramers D, Wazniak C, Wiggert J, Wood R, Xu J (2012) Ecological Forecasting in Chesapeake Bay: Using a Mechanistic- Empirical Modelling Approach. Journal of Marine Systems Campagnolo ER, Johnson KR, Karpati A, Rubin CS, Kolpin DW, Meyer MT, Esteban JE, Currier RW, Smith K, Thu KM, McGeehin M (2002) Antimicrobial residues in animal waste and water resources proximal to large-scale swine and poultry feeding operations. The Science of the total environment 299:89-95 CDC (2009) Vibrio vulnificus: General Information. Accessed January 6. http://www.cdc.gov/nczved/divisions/dfbmd/diseases/vibriov/ CDC (2012a) Cholera and Other Vibrio Illness Surveillance Overview. In. US Department of Health and Human Services, Atlanta, Georgia CDC (2012b) Increasing prevalence of diagnosed diabetes--United States and Puerto Rico, 1995-2010. MMWR Morb Mortal Wkly Rep 61:918-921 CDC (2012c) Trends in Foodborne Illness in the United States, 1996-2010. Accessed October 2, 2012. http://www.cdc.gov/foodborneburden/trends-in-foodborne- illness.html 118 CFSAN (Center for Food Safety and Applied Nutrition) USFDA (2005) Quantitative Risk Assessment on the Public Health Impact of Pathogenic Vibrio parahaemolyticus in Raw Oysters. In. U.S. Food and Drug Administration Chiew YF, Yeo SF, Hall LM, Livermore DM (1998) Can susceptibility to an antimicrobial be restored by halting its use? The case of streptomycin versus Enterobacteriaceae. J Antimicrob Chemother 41:247-251 Colwell RR (2000) Viable but nonculturable bacteria: a survival strategy. J Infect Chemother 6:121-125 COMAR (2013) Maryland Code of Regulations: 26.08.02.03-3 Water Quality Criteria Specific to Designated Uses. In: Environment Dot (ed), Annapolis, MD Criminger JD, Hazen TH, Sobecky PA, Lovell CR (2007) Nitrogen fixation by Vibrio parahaemolyticus and its implications for a new ecological niche. Applied and Environmental Microbiology 73:5959-5961 Daniels NA, MacKinnon L, Bishop R, Altekruse S, Ray B, Hammond RM, Thompson S, Wilson S, Bean NH, Griffin PM, Slutsker L (2000) Vibrio parahaemolyticus infections in the United States, 1973-1998. The Journal of infectious diseases 181:1661-1666 Daniels NA, Shafaie A (2000) A Review of Pathogenic Vibrio Infections for Clinicians. Infections in medicine 17:665-685 de Oliveira AJ, Pinhata JM (2008) Antimicrobial resistance and species composition of Enterococcus spp. isolated from waters and sands of marine recreational beaches in Southeastern Brazil. Water Res 42:2242-2250 Dechet AM, Yu PA, Koram N, Painter J (2008) Nonfoodborne Vibrio infections: an important cause of morbidity and mortality in the United States, 1997-2006. Clin Infect Dis 46:970-976 DePaola A, Kaysner CA (2004) Vibrio, chapter 9. In Bacteriological analytical manual. . In. U.S. Food and Drug Administration, Washington, DC. Dickinson G, Lim KY, Jiang SC (2013) Quantitative microbial risk assessment of pathogenic vibrios in marine recreational waters of southern california. Appl Environ Microbiol 79:294-302 Dufour AP, Evans O, Behymer TD, Cantu R (2006) Water ingestion during swimming activities in a pool: a pilot study. J Water Health 4:425-430 Eaton AD, Clesceri LS, Greenberg AE, Franson MAH, American Public Health Association., American Water Works Association., Water Environment Federation. (1998) Standard methods for the examination of water and wastewater, Vol. American Public Health Association, Washington, DC 119 EPA US (2011a) The Great Waters Program: Introduction to the Issues and Ecosystesms, Chesapeake Bay. Accessed February 10. http://www.epa.gov/oaqps001/gr8water/xbrochure/chesapea.html EPA US (2011b) Recreational Water Quality Criteria Draft Document. In. U.S. Environmental Protection Agency, Office of Water, Washington, DC Ewing B, Green P (1998) Base-calling of automated sequencer traces using phred. II. Error probabilities. Genome Research 8:186-194 Ewing B, Hillier L, Wendl MC, Green P (1998) Base-calling of automated sequencer traces using phred. I. Accuracy assessment. Genome Research 8:175-185 FDA (1998) R59: Phosphate-Buffered Saline (PBS), pH 7.4. http://www.fda.gov/Food/ScienceResearch/LaboratoryMethods/BacteriologicalA nalyticalManualBAM/ucm062268.htm FDA (2012) Bad Bug Book: Foodborne Pathogenic Microorganisms and Natural Toxins Handbook. In: Vibrio Washington, DC Ford TE, Colwell RR, Rose JB, Morse SS, Rogers DJ, Yates TL (2009) Using satellite images of environmental changes to predict infectious disease outbreaks. Emerg Infect Dis 15:1341-1346 Fries JS, Characklis GW, Noble RT (2008) Sediment???water exchange of< i> Vibrio sp. and fecal indicator bacteria: Implications for persistence and transport in the Neuse River Estuary, North Carolina, USA. Water Res 42:941- 950 Froelich BA, Williams TC, Noble RT, Oliver JD (2012) Apparent loss of Vibrio vulnificus from North Carolina oysters coincides with a drought-induced increase in salinity. Appl Environ Microbiol 78:3885-3889 Goldenberg SB, Landsea CW, Mestas-Nunez AM, Gray WM (2001) The recent increase in Atlantic hurricane activity: causes and implications. Science 293:474-479 Gordon D, Abajian C, Green P (1998) Consed: A graphical tool for sequence finishing. Genome Research 8:195-202 Gotoh K, Kodama T, Hiyoshi H, Izutsu K, Park KS, Dryselius R, Akeda Y, Honda T, Iida T (2010) Bile acid-induced virulence gene expression of Vibrio parahaemolyticus reveals a novel therapeutic potential for bile acid sequestrants. PLoS ONE 5:e13365 Groubert TN, Oliver JD (1994) Interaction of Vibrio vulnificus and the eastern oyster, Crassostrea virginica. Journal of Food Protection 57:224-228 120 Han F, Walker RD, Janes ME, Prinyawiwatkul W, Ge B (2007) Antimicrobial susceptibilities of Vibrio parahaemolyticus and Vibrio vulnificus isolates from Louisiana Gulf and retail raw oysters. Appl Environ Microbiol 73:7096-7098 Hedlund BP, Staley JT (2001) Vibrio cyclotrophicus sp. nov., a polycyclic aromatic hydrocarbon (PAH)-degrading marine bacterium. Int J Syst Evol Microbiol 51:61-66 Hsieh JL, Fries S, Noble RT (2008) Dynamics and predictive modelling of Vibrio spp. in the Neuse River Estuary, North Carolina, USA. Environ Microbiol 10:57-64 IPCC (2007) Climate Change 2007: Synthesis report. IPCC, Geneva Jacobs J, Rhodes M, Sturgis B, Wood B (2009) Influence of environmental gradients on the abundance and distribution of Mycobacterium spp. in a coastal lagoon estuary. Appl Environ Microbiol 75:7378-7384 Jacobs JM, Rhodes MR, Brown CW, Hood RR, Leight AK, Long W, Wood RJ (2010) Predicting the distribution of Vibrio vulnificus in Chesapeake Bay, Vol. Oxford Cooperative Lab Johnson CN (2013) Fitness Factors in Vibrios: a Mini-review. Microbial Ecology:1-26 Johnson CN, Bowers JC, Griffitt KJ, Molina V, Clostio RW, Pei S, Laws E, Paranjpye RN, Strom MS, Chen A, Hasan NA, Huq A, Noriea NF, 3rd, Grimes DJ, Colwell RR (2012) Ecology of Vibrio parahaemolyticus and Vibrio vulnificus in the coastal and estuarine waters of Louisiana, Maryland, Mississippi, and Washington, United States. Appl Environ Microbiol 78:7249-7257 Johnson CN, Flowers AR, Noriea NF, 3rd, Zimmerman AM, Bowers JC, DePaola A, Grimes DJ (2010) Relationships between environmental factors and pathogenic Vibrios in the Northern Gulf of Mexico. Appl Environ Microbiol 76:7076-7084 Jones A, Carruthers T, Pantus F, Thomas J, Saxby T, Dennison W (2004) A water quality assessment of the Maryland Coastal Bays including nitrogen source identification using stable isotopes. In. Integration and Application Network, University of Maryland Center for Environmental Science, Marine Ecoservices Australia Jones JL, Ludeke CH, Bowers JC, Garrett N, Fischer M, Parsons MB, Bopp CA, DePaola A (2012) Biochemical, serological, and virulence characterization of clinical and oyster Vibrio parahaemolyticus isolates. J Clin Microbiol 50:2343-2352 Joseph SW, DeBell RM, Brown WP (1978) In vitro response to chloramphenicol, tetracycline, ampicillin, gentamicin, and beta-lactamase production by halophilic Vibrios from human and environmental sources. Antimicrob Agents Chemother 13:244-248 121 Kaneko T, Colwell RR (1973) Ecology of Vibrio parahaemolyticus in Chesapeake Bay. J Bacteriol 113:24-32 Kaufman GE, Bej AK, Bowers J, DePaola A (2003) Oyster-to-oyster variability in levels of Vibrio parahaemolyticus. J Food Prot 66:125-129 Lipp EK, Huq A, Colwell RR (2002) Effects of global climate on infectious disease: the cholera model. Clinical Microbiology Reviews 15:757-770 Lipp EK, Rose JB (1997) The role of seafood in foodborne diseases in the Unites States of America. Rev Sci Tech Office Int Epizootics 16:620-640 Loosanoff VL, Tommers FD (1948) Effect of Suspended Silt and Other Substances on Rate of Feeding of Oysters. Science 107:69-70 Luckenbach MW, O?Beirn FX, Taylor J (1999) An Introduction to Culturing Oysters in Virginia, Vol. School of Marine Science, Virginia Institute of Marine Science, College of William & Mary, Gloucester Point, VA. Ludwig W, Strunk O, Westram R, Richter L, Meier H, Yadhukumar, Buchner A, Lai T, Steppi S, Jobb G, Forster W, Brettske I, Gerber S, Ginhart AW, Gross O, Grumann S, Hermann S, Jost R, Konig A, Liss T, Lussmann R, May M, Nonhoff B, Reichel B, Strehlow R, Stamatakis A, Stuckmann N, Vilbig A, Lenke M, Ludwig T, Bode A, Schleifer KH (2004) ARB: a software environment for sequence data. Nucleic Acids Res 32:1363-1371 Mahoney JC, Gerding MJ, Jones SH, Whistler CA (2010) Comparison of the pathogenic potentials of environmental and clinical vibrio parahaemolyticus strains indicates a role for temperature regulation in virulence. Appl Environ Microbiol 76:7459- 7465 Martinez JL (2003) Recent Advances on antibiotic resistance genes. In: Fingerman, Nagabhushanam (eds) Recent Advances in Marine Biotechnology Molecular Genetics of Marine Organisms, Book 10 Martinez JL (2008) Antibiotics and Antibiotic Resistance Genes in Natural Environments. Science 321:365-367 Mead PS, Slutsker L, Dietz V, McCaig LF, Bresee JS, Shapiro C, Griffin PM, Tauxe RV (1999) Food-related illness and death in the United States. Emerg Infect Dis 5:607-625 Meehl GA, Stocker TF, Collins WD, Friedlingstein P, Gaye AT, Gregory JM, Kitoh A, Knutti R, Murphy JM, Noda A, Raper SCB, Watterson IG, Weaver AJ, Zhao ZC (2007) Global Climate Projections. In: Solomon S, Qin D, Manning M, Chen Z, Marquis M, Averyt KB, Tignor M, Miller HL (eds) Climate Change 2007: The Physical Science Basis Contribution of Working Group I to the Fourth 122 Assessment Report of the Intergovernmental Panel on Climate Change, Cambridge, United Kingdom and New York, New York Merkel SM, Alexander S, Zufall E, Oliver JD, Huet-Hudson YM (2001) Essential role for estrogen in protection against Vibrio vulnificus-induced endotoxic shock. Infect Immun 69:6119-6122 Molodecky NA, Soon IS, Rabi DM, Ghali WA, Ferris M, Chernoff G, Benchimol EI, Panaccione R, Ghosh S, Barkema HW (2012) Increasing incidence and prevalence of the inflammatory bowel diseases with time, based on systematic review. Gastroenterology 142:46-54. e42 Morris JG, Jr. (2003) Cholera and other types of vibriosis: a story of human pandemics and oysters on the half shell. Clin Infect Dis 37:272-280 Mosteller RD (1987) Simplified calculation of body-surface area. N Engl J Med 317:1098 Motes ML, DePaola A, Cook DW, Veazey JE, Hunsucker JC, Garthright WE, Blodgett RJ, Chirtel SJ (1998) Influence of Water Temperature and Salinity on Vibrio vulnificus in Northern Gulf and Atlantic Coast Oysters (Crassostrea virginica). Appl Environ Microbiol 64:1459-1465 Myasoedova E, Crowson CS, Kremers HM, Therneau TM, Gabriel SE (2010) Is the incidence of rheumatoid arthritis rising?: results from Olmsted County, Minnesota, 1955-2007. Arthritis and rheumatism 62:1576-1582 Nielsen LT, Hallegraeff GM, Wright SW, Hansen PJ (2012) Effects of experimental seawater acidification on an estuarine plankton community. Aquat Microb Ecol 65:271-285 NOAA NWS (2011) Post Tropical Storm Report: Hurricane Irene. Accessed February 2, 2013. http://www.erh.noaa.gov/akq/wx_events/hur/Irene/index.html Nordstrom JL, Vickery MC, Blackstone GM, Murray SL, DePaola A (2007) Development of a multiplex real-time PCR assay with an internal amplification control for the detection of total and pathogenic Vibrio parahaemolyticus bacteria in oysters. Appl Environ Microbiol 73:5840-5847 Oh MH, Lee SM, Lee DH, Choi SH (2009) Regulation of the Vibrio vulnificus hupA gene by temperature alteration and cyclic AMP receptor protein and evaluation of its role in virulence. Infect Immun 77:1208-1215 Oliver JD (2005) The viable but nonculturable state in bacteria. J Microbiol 43 Spec No:93-100 Oliver JD (2012) Vibrio vulnificus: Death on the Half Shell. A Personal Journey with the Pathogen and its Ecology. Microbial Ecology:1-7 123 Orr JC, Fabry VJ, Aumont O, Bopp L, Doney SC, Feely RA, Gnanadesikan A, Gruber N, Ishida A, Joos F (2005) Anthropogenic ocean acidification over the twenty-first century and its impact on calcifying organisms. Nature 437:681-686 Panicker G, Bej AK (2005) Real-time PCR detection of Vibrio vulnificus in oysters: comparison of oligonucleotide primers and probes targeting vvhA. Appl Environ Microbiol 71:5702-5709 Parveen S, Hettiarachchi KA, Bowers JC, Jones JL, Tamplin ML, McKay R, Beatty W, Brohawn K, DaSilva LV, DePaola A (2008) Seasonal distribution of total and pathogenic< i> Vibrio parahaemolyticus in Chesapeake Bay oysters and waters. International journal of food microbiology 128:354-361 Randa MA, Polz MF, Lim E (2004) Effects of temperature and salinity on Vibrio vulnificus population dynamics as assessed by quantitative PCR. Appl Environ Microbiol 70:5469-5476 Sanford LP (1994) Wave-Forced Resuspension of Upper Chesapeake Bay Muds. Estuaries 17:148-165 Sapkota AR, Curriero FC, Gibson KE, Schwab KJ (2007) Antibiotic-resistant enterococci and fecal indicators in surface water and groundwater impacted by a concentrated swine feeding operation. Environmental Health Perspectives 115:1040 Scallan E, Griffin PM, Angulo FJ, Tauxe RV, Hoekstra RM (2011a) Foodborne illness acquired in the United States--unspecified agents. Emerg Infect Dis 17:16-22 Scallan E, Hoekstra RM, Angulo FJ, Tauxe RV, Widdowson MA, Roy SL, Jones JL, Griffin PM (2011b) Foodborne illness acquired in the United States--major pathogens. Emerg Infect Dis 17:7-15 Shaw K, Sapkota A, Jacobs J, Crump B (2011) Recreational Swimmers? Exposure to Vibrio vulnificus and Vibrio parahaemolyticus in Chesapeake Bay, Maryland, USA. In: Vibrio 2011: The fourth conference on the biology of vibrios, Santiago de Compostela, Spain Shiah FK, Ducklow HW (1994) Temperature regulation of heterotrophic bacterioplankton abundance, production, and specific growth rate in Chesapeake Bay. Limnol Oceanogr 39:1243-1258 Staley C, Harwood VJ (2010) The use of genetic typing methods to discriminate among strains of Vibrio cholerae, V. parahaemolyticus, and V. vulnificus. J AOAC Int 93:1553-1569 Staley C, Jones MK, Wright AC, Harwood VJ (2011) Genetic and quantitative assessment of Vibrio vulnificus populations in oyster (Crassostrea virginica) tissues. Environmental Microbiology Reports 3:543-549 124 Strom MS, Paranjpye RN (2000) Epidemiology and pathogenesis of Vibrio vulnificus. Microbes and infection / Institut Pasteur 2:177-188 Thiaville PC, Bourdage KL, Wright AC, Farrell-Evans M, Garvan CW, Gulig PA (2011) Genotype is correlated with but does not predict virulence of Vibrio vulnificus biotype 1 in subcutaneously inoculated, iron dextran-treated mice. Infect Immun 79:1194-1207 Thomas J, Woerner J, Abele R, Cain C, Charland J, Jesien R, Silaphone K (2009) Ch. 5, St. Martin River. Shifting Sands: Environmental and cultural change in Maryland's Coastal Bays. In: Dennison WC TJ, Cain CJ, Carruthers TJB, Hall MR, Jesien RV, Wazniak CE, Wilson DE (ed), Cambridge, MD Vezzulli L, Colwell RR, Pruzzo C (2013) Ocean Warming and Spread of Pathogenic Vibrios in the Aquatic Environment. Microbial Ecology:1-9 Vizcaino MI, Johnson WR, Kimes NE, Williams K, Torralba M, Nelson KE, Smith GW, Weil E, Moeller PD, Morris PJ (2010) Antimicrobial resistance of the coral pathogen Vibrio coralliilyticus and Caribbean sister phylotypes isolated from a diseased octocoral. Microb Ecol 59:646-657 Warner EaJDO (2008) Population Structures of Two Genotypes of Vibrio vulnificus in Oysters (Crassostrea virginica) and Seawater. Appl Environ Microbiol 74:80-85 Wetz J, Blackwood A, Fries J, Williams Z, Noble R (2008) Trends in total Vibrio spp. and Vibrio vulnificus concentrations in the eutrophic Neuse River Estuary, North Carolina, during storm events. Aquat Microb Ecol 53:141-149 World Health Organization., Food and Agriculture Organization of the United Nations. (2005) Risk assessment of Vibrio vulnificus in raw oysters : interpretative summary and technical report, Vol. World Health Organization; Food and Agriculture Organization of the United Nations, Geneva; Rome World Health Organization., Food and Agriculture Organization of the United Nations. (2011) Risk assessment of Vibrio parahaemolyticus in seafood : interpretative summary and technical report, Vol. World Health Organization ; Food and Agriculture Organization of the United Nations, Geneva, Rome Wright AC, Hill RT, Johnson JA, Roghman MC, Colwell RR, Morris JG, Jr. (1996) Distribution of Vibrio vulnificus in the Chesapeake Bay. Applied and Environmental Microbiology 62:717-724 Wright GD (2007) The antibiotic resistome: the nexus of chemical and genetic diversity. Nat Rev Micro 5:175-186 Yamazaki K, Esiobu N (2012) Environmental Predictors of Pathogenic Vibrios in South Florida Coastal Waters. The Open Epidemiology Journal 5:1-4 125 Zimmerman AM, DePaola A, Bowers JC, Krantz JA, Nordstrom JL, Johnson CN, Grimes DJ (2007) Variability of total and pathogenic Vibrio parahaemolyticus densities in northern Gulf of Mexico water and oysters. Appl Environ Microbiol 73:7589-7596