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MICROBIAL COMMUNITY AND PARAHAEMOLYTICUS POPULATION DYNAMICS IN RELAYED

Michael Anthony Taylor University of New Hampshire, Durham

Follow this and additional works at: https://scholars.unh.edu/dissertation

Recommended Citation Taylor, Michael Anthony, "MICROBIAL COMMUNITY AND VIBRIO PARAHAEMOLYTICUS POPULATION DYNAMICS IN RELAYED OYSTERS" (2017). Doctoral Dissertations. 2288. https://scholars.unh.edu/dissertation/2288

This Dissertation is brought to you for free and open access by the Student Scholarship at University of New Hampshire Scholars' Repository. It has been accepted for inclusion in Doctoral Dissertations by an authorized administrator of University of New Hampshire Scholars' Repository. For more information, please contact [email protected]. MICROBIAL COMMUNITY AND VIBRIO PARAHAEMOLYTICUS POPULATION DYNAMICS IN RELAYED OYSTERS

BY

MICHAEL ANTHONY TAYLOR

BS, University of New Hampshire, 2002 Master’s Degree, University of New Hampshire, 2005

DISSERTATION

Submitted to the University of New Hampshire in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy in Microbiology

September, 2017

This dissertation has been examined and approved in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Microbiology by:

Dissertation Director, Stephen H. Jones Research Associate Professor, Natural Resources and the Environment

Cheryl A. Whistler, Associate Professor, Molecular, Cellular & Biomedical Sciences

Vaughn S. Cooper, Associate Professor, Microbiology & Molecular Genetics, University of Pittsburg School of Medicine

Kirk Broders, Assistant Professor, Plant Pathology, Colorado State University College of Agricultural Sciences

Thomas Howell, President / Owner, Spinney Creek Shellfish, Inc., Eliot, Maine

On March 24, 2017

Original approval signatures are on file with the University of New Hampshire Graduate School.

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ACKNOWLEDGEMENTS

I would like to thank the following individuals for helping me with this endeavor:

Dr. Steve Jones: Steve, thank you for taking me on as a graduate student six years ago and providing me the flexibility to be an unorthodox student with other priorities as important as school. Thank you for all of the advice, and the time you have dedicated to helping me.

Graduate Committee: Thank you to all 5 of my committee members. Each one of you helped me grow professionally. Thank you for your time and willingness to provide guidance and support me as a student.

Tom and Laurie Howell: Special thanks to the Howell’s and Spinney Creek Shellfish, Inc. for their generosity with their time and their facility which made this research novel.

Co-workers in the lab: Thank you to all of my fellow students and the students and technicians in the Jones and Whistler labs who assisted in this project.

Bob Mooney: Bob, thank you for showing me how to be a Microbiologist.

Dr. Robert Zsigray: I want to thank the late Bob Zsigray for teaching me about integrity in research and life.

Dr. Stephen Torosian: Thank you Steve for my start as a microbiologist and showing me into the lab.

Jong Yu: Thank you Jong for teaching me how to process oysters and all of your hard work and support during this project.

Mike Hall Mike thank you for the computer science support.

Cristiane Taylor: To my wife – I didn’t realize how tough getting a PhD is until I was at the point of no return. Thank you for making sure I did not give up and thank you for being a mother, friend, wife and an unwavering partner through the last 6 years.

Funding Sources: NH Agricultural Experiment Station NH Sea Grant College Program NH EPSCoR Program-Track 2 project

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TABLE OF CONTENTS

ACKNOWLEDGEMENTS...... iii

LIST OF TABLES...... v

LIST OF FIGURES ...... vi

ABSTRACT ...... vii

CHAPTER PAGE

I. INTRODUCTION...... 1

II. VARYING SUCCESS OF RELAYING TO REDUCE VIBRIO PARAHAEMOLYTICUS LEVELS IN OYSTERS (CRASSOSTREA VIRGINICA) …………………………………………… 12

INTRODUCTION……………………………………………………. 13

RESULTS…………………………………………………………….. 17

DISCUSSION………………………………………………………… 26

MATERIALS AND METHODS…………………………………….. 30

III. MICROBIAL COMMUNITY CHANGES ASSOCIATED WITH VIBRIO PARAHAEMOLYTICUS CONCENTRATIONS IN RELAYED OYSTERS………………………………………………………………… 34

INTRODUCTION……………………………………………………. 35

RESULTS…………………………………………………………….. 37

DISCUSSION………………………………………………………… 60

MATERIALS AND METHODS……………………………………... 64

REFERENCES………………………………………………………………………... 71 APPENDICES………………………………………………………………………… 88 APPENDIX A CHAPTER II SUPPLEMENTAL MATERIAL……………………… 89 APPENDIX B CHAPTER III SUPPLEMENTAL MATERIAL……………………... 96

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LIST OF TABLES

CHAPTER II

Table 1 Statistical analyses of V. parahaemolyticus (Vp) levels (MPN/g or mL) in oyster and water samples analyzed during relay experiments from 2011 to 2014……………………………………………………. 22

CHAPTER III

Table 1 V. parahaemolyticus concentrations and 16S sequence abundance for Vibrio spp. in paired Day 0 and 14 oyster samples from the four most effective relay experiments: 2011-12……………………… 41

Table 2 Identified genera that increased from Day 0 to Day 14 in oysters from the most effective relay experiments…………………………… 44

Table 3 Non-parametric Multi Response Permutation Procedure (MRPP) analysis of Taxa Differences within Sample Matrices and Years – 2011-2013 Relay Experiments……………………………………….. 49

Table 4 The diversity and relative abundance of taxa in oysters, water and both water and oysters in each study year (2011, 2012 and 2013)…... 58

Table 5 Vibrio spp. used in the competition assay with environmental strain G3654 of Vibrio parahaemolyticus from Spinney Creek……… 60

APPENDIX

Table A.1 2011-2014 Oyster and Water Sample MPN Metadata………………… 91

Table B.1 Oyster and water sample V. parahaemolyticus MPN/g concentrations and 16S count metadata for samples submitted for 16S rRNA sequencing………………………………………………………… 97

Table B.2 CASAVA Quality Data for Oyster and Water Samples……………….. 98

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LIST OF FIGURES

CHAPTER II

Fig. 1 The Great Bay estuary of Maine and New Hampshire, USA, with harvest site (PR) in the Piscataqua River and the relay site (SC) in Spinney Creek ………………………………………………… 16

Fig. 2 Water temperature (°C) and salinity (PPT-parts per thousand) at the relay (SC) and harvest (PR) sites………………………………… 19

Fig. 3 V. parahaemolyticus levels (MPN/g) on Day 0 of relay experiments in oysters from PR and water samples from PR and SC……………… 21

Fig. 4 V. parahaemolyticus levels (MPN/g) in oyster samples analyzed during each sampling day of relay experiments from 2011 to 2014….. 24

CHAPTER III

Fig. 1 Alpha rarefaction curves for sequences within the observed OTUs….. 39

Fig. 2 Linear regression analysis between V. parahaemolyticus MPN concentrations and Vibrio spp. rRNA gene abundances in all samples………………………………………………………… 42

Fig. 3 The relationship between V. parahaemolyticus concentrations and genera abundance in relayed oyster samples…………………….. 46

Fig. 4 Principal coordinates analysis of all oyster and water samples based on pair-wise sample Bray-Curtis dissimilarity measures……… 48

Fig. 5 The relative abundance and relatedness between OTUs associated with fresh and relayed oyster and water samples during 2011-13…….. 55

Fig. 6 Venn diagram of the number of identified taxa found in 100% of the oyster and water samples by year and Day 14 of relay…. 57

APPENDIX

Fig. A.1 Lab procedure for processing oysters after field collection..…………. 90

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ABSTRACT

MICROBIAL COMMUNITY AND VIBRIO PARAHAEMOLYTICUS POPULATION DYNAMICS IN RELAYED OYSTERS

by

Michael Anthony Taylor

University of New Hampshire, September, 2017

The CDC estimates that 45,000 people are sickened each year by foodborne Vibrio parahaemolyticus in the United States. Filter-feeding bivalve shellfish, such as oysters, are routinely inhabited by human pathogenic V. parahaemolyticus and there currently is not a contaminant management process that effectively reduces concentrations of V. parahaemolyticus in oysters. The transplanting of V. parahaemolyticus -laden oysters to an area with low concentrations or no V. parahaemolyticus, called oyster relay, is one reduction strategy that holds promise for treating live oysters. A key consideration for effective strategies to reduce Vibrio spp. in shellfish is the influence of in natural seawater. Our aim for this study was to identify taxa shifts in the microbial composition of oyster and water samples during relay that correlated with the reduction of V. parahaemolyticus. We hypothesize that changes in bacterial taxa within the oyster microbiome occur during relay and influence the reduction of V. parahaemolyticus in relayed oysters. Oysters with varying concentrations of V. parahaemolyticus were evident from relay experiments carried out in a local body of water with elevated salinity and low concentrations of V. parahaemolyticus over 4 consecutive years.

Overall, V. parahaemolyticus levels were reduced during 14-day relay in 9 of the 14 monthly trials to target reduction levels (<100 V. parahaemolyticus/g). Sequence analysis of the 16S rRNA gene from relayed oyster and the associated water samples unveiled that the

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composition of oyster microbial communities shifts during relay. We determined that oyster and water taxa profiles are dissimilar while harvest and relay waters were similar in overall taxa composition, even though there was also evidence of some taxa differences that may have been influential in reducing V. parahaemolyticus concentrations during the 14-day relay. Oyster samples from relay experiments in years that successfully reduced V. parahaemolyticus concentrations had consistent, similar taxa patterns compared to different and inconsistent patterns for 2013 when relay was not successful. Interspecies competition experiments informed by these analyses suggested one potential mechanistic explanation (competition) for why relaying to higher salinity water reduces V. parahaemolyticus concentrations. Oyster microbiome competition may aid in developing a consistent approach for reducing V. parahaemolyticus in oysters to reduce the risk of illness for oyster consumers.

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CHAPTER I

INTRODUCTION

The avid interest and appeal of the American population to consume oysters (1), and the public’s interest in the safety of oysters are both on the rise. People often eat oysters and other shellfish raw or with minimal cooking. From the consumption of these delicacies comes the risk of getting sick with an illness called vibriosis. An estimated 52,000 (but only 1,252 actually reported in 2014) (2) annual cases of vibriosis in the United States are associated with the consumption of food (3). 87% of these illnesses are from Vibrio parahaemolyticus (4) and the primary food matrix associated with vibriosis is oysters (2).

The majority of Vibrio are not pathogenic nor are they harmful to humans, plants or (5) but a select few are known to be the cause of human illness. The majority of human illnesses from are associated with Vibrio parahaemolyticus, and

Vibrio cholerae. Common factors with the pathogenic strains of these species are associations with colonization and attachment to surface areas, , nutrient acquisition, competition and adaptation in different environments (5).

The large degree of fluctuation within the estuarine ecosystems influences the predominance of different Vibrio species at different points in time. Survival mechanisms associated with fitness factors, influenced by the environment, are driven by the need to consume nutrients (6) and the ability to compete with exogenous factors via adaptation (7). A mechanism consistently involved in these adaptations is (5), which in addition to environmental changes can create advantages to survival and the increase or decrease in pathogenicity (8, 9).

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Vibrio parahaemolyticus is a halophilic bacterium very commonly found in brackish waters, grows optimally between 35-37 oC and was first discovered as a in 1950 (10).

With a short (4-96 h) and one of the fastest doubling times of all

(under ~ 10 min) (11) the effect of the two , thermostable direct (TDH) and TDH-related hemolysin (TRH), along with other virulence factors (12), can generate the rapid onset of vomiting, watery and bloody and .

One of the most significant environmental factors that have attributed to the rise of V. parahaemolyticus cases over the past decades is sea surface temperature (SST) (13-15). As the global water temperatures rise, pathogenic Vibrio concentrations are higher and these species remain more prevalent which directly correlate to the increase in disease (16). Not only does the

SST affect the proliferation of Vibrios but it also affects biotic and abiotic factors that influence

Vibrios (17-19).

The Great Bay Estuary (GBE) in New Hampshire, like other coastal waters, has been affected by warming sea surface temperatures. The diverse bacterial populations within the estuary (20) in conjunction with increase in SST promote shifts in bacterial communities and give rise to recombination events (21) that have the potential to generate pathogenic strains (22) to increase incidence of illness and outbreaks.

The incidence of from oysters in the United States continues to rise year over year (23) along with the need to mitigate these illnesses. The Interstate Sanitation

Shellfish Conference (ISSC) in conjunction with the FDA and State shellfish programs, establish policy to safeguard consumers from contaminated oysters by requiring routine monitoring and classification of harvest waters. Additionally, safeguards for sanitary transportation of oysters are included in the Food Safety Modernization Act (FSMA) regulations (24).

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The Food Safety Modernization Act (FSMA), which gives the FDA more extensive overarching authority over the regulation of food producers and manufacturers, went into effect

September of 2016. Under FSMA, oysters are regulated differently than other commodities. New legislation for mandatory oyster treatments cannot be implemented without the involvement of industry (25), however industry and regulators continue to collaborate in an attempt to expand the model ordinances with feasible contaminant reduction strategies to prevent illness.

Legislation around validated reduction strategies for that cause consumers to get sick is lacking. The average consumer is not informed about the advancement of food regulations

(26) or the legislation gaps in the oyster industry. These gaps in legislation and public knowledge make the development of processes to reduce oyster contamination in the food supply chain more important than ever.

‘Effective’ processing of oysters

Reducing the incidence of Vibrio-borne illness is important for all involved from cultivating and harvesting, to selling and consuming oysters. Even though there are innumerable ways that oyster quality may be compromised along the way from the harvest area to the consumer, one option for reducing public health threats is to be able to process oysters after harvest in a manner to reduce concentrations of potentially pathogenic Vibrio spp.

Existing post-harvest processing (PHP) strategies can have different effects on the oyster quality and reduction of pathogens (27-29) so the perspective on whether or not a PHP is effective depends on the stakeholder. In general, consumers, harvesters, sellers and regulators are all keenly interested in and responsible for preventing illness but have different agendas when it comes to how to achieve this.

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Consumers. Today’s consumer expects and assumes that the food they eat is safe (26) and can be lax with properly educating themselves about all potential risks. Education campaigns about at risk populations can potentially be effective (30), but some consumers who are at risk, whether knowingly or not, continue to eat oysters raw. The number of consumers of raw oysters in the mainstream population is small, but those who do partake expect that they are not paying for food that will make them sick. From the consumer’s perspective, effective oyster harvest management involves processes that allow for oysters to be consumed how and when consumers choose.

Harvesters and Sellers. Oyster harvesters abide by guidelines from the National Shellfish

Sanitation Program (NSSP) (31) that must be followed to safely market harvested oysters for sale. Following the mandated guidelines ensures that harvesters use ‘approved’ classified waters and best management practices (BMPs) to handle and harvest the oysters, but the guidelines do not prescribe any specific PHP methods to mitigate Vibrios in oysters. Current practices used by harvesters, mandated by the regulators, affect the financial bottom line by adding time and money to the process, but this approach has been effective in reducing illnesses in the Northeast

(32) thus preventing costly closures, recalls and bad publicity. No pre-or post-harvest process can prevent all illness, as that is probably an impossible goal, so an acceptable incidence of illness is a more practical goal. Effective management approaches for harvesters and sellers are processes that are cost effective and that substantially reduce illnesses.

Regulators. Out of all the groups of stakeholders, the FDA should be the most informed and influential in mitigating illness from the consumption of oysters. In 2009 the FDA attempted to address the issues of shellfish-borne Vibrio illness with measures that did not take into account all stakeholders. Because of the lack of industry involvement and support, the legislation was

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deemed not feasible and was revoked by the FDA (33). Mitigation strategies that have been effective in the State of California were proposed for trial in the Gulf Coast region but local officials and industry leaders blocked the attempt by the government to prevent illness for an array of reasons. Effective processing for regulators needs to be methods that effectively reduce cases of illness and are accepted by the industry.

Scientists. The discovery of effective, validated methods to reduce illness from raw shellfish consumption will come from research involving all the above stakeholders. The scientific community is the primary resource to regulators and industry for scientific information about how illness can be reduced. The scientific community carries out applied and basic research to advance scientific knowledge with the hope of contributing discoveries that can lead to effective contaminant reduction strategies that meet expectations of the oyster industry, regulators and consumers.

Current Contaminant Reduction Strategies for Bivalve Shellfish

There are a variety of ways to process oysters to reduce pathogens but the majority of these processes have not been fully validated, do not meet consumer preferences and/or have a significant negative effect on the organoleptic attributes of the oyster (27-29, 34-38). A common variable with these processes is the displacement of the oysters away from their natural environment to a facility with artificial conditions where oyster function can be sub-optimal.

Vibrio reduction strategies located in coastal waters near harvest areas are desirable for many reasons, yet there are also many challenges. Oysters filter feed high volumes of estuarine waters and have an influential effect on the estuarine environment (39). The filter feeding process generates different interplays between the oyster and the microbiota in the surrounding

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water (40, 41). Fecal-borne and indigenous pathogenic microflora are both concerns when it comes to illness from oyster consumption. Nowadays, locating unpolluted coastal areas for contaminant reduction strategies can be difficult and oysters located in areas that have not been adulterated (e.g. away from sewage polluted areas) can contain resident microflora that are pathogenic (e.g. Vibrio spp.).

One effective way to control the microbiological contamination of oysters is to harvest the oysters from areas with ‘clean’ water. Unfortunately, the NSSP model ordinances currently in place do not include any contaminant reduction strategies that have been fully evaluated for reducing naturally occurring (pathogenic) Vibrio spp. in oysters to safe concentrations, and preserve the appealing organoleptic attributes of the oysters. Instead, pre- and post-harvest handling practices, classification of harvest waters and product testing are the control measures used to mitigate the Vibrio-related public health risks of marketed oysters. (42). Time and the degree of reduction of Vibrio concentrations are the variables that are most important for any treatment process for oysters. The most effective process would incorporate factors that would prevent the persistence of viral, bacterial and chemical agents, and work in a timely manner for the process to be feasible to industry. For this dissertation research, the focus is on one potentially pathogenic Vibrio species, V. parahaemolyticus.

Depuration. Depuration is defined as the action or process of freeing something from impurities.

Shellfish depuration, which has been around in the United States since the beginning of the

1900’s (43), was established in response to outbreaks of typhoid . It is one of the most effective post-harvest processes for reducing fecal coliforms in oysters, but it is not as effective with viruses and resident oyster microbiota like Vibrio species that can cause widespread disease across the world (44-46). The premise of depuration in the oyster industry is to place oysters into

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tanks with controlled parameters in regards to tank water conditions, tank cleanliness and contaminant reduction methods to free the oysters of impurities (42) by letting them filter feed and purge contaminants in this controlled environment. The depuration process is relatively expensive to set up and maintain but effective for the removal of fecal coliforms and variably effective for Vibrio spp. reduction (44, 47). Thus, standard depuration can cleanse shellfish of fecal coliforms while still having unacceptable concentrations of other bacteria (Vibrio spp.) and viruses (Norovirus) that can make consumers sick (48-50).

For reducing concentrations of pathogenic Vibrio species, the use of depuration requires consideration of several factors. Depuration relies on the proper set up of expensive tank systems and intensive maintenance of a controlled environment to purify oysters of impurities, so it requires training and skilled oversight. Additionally, oysters displaced from their natural environment can be prone to die off if they are not properly handled (51). Other post-harvest processes, such as relay (see next section), are not as time efficient as depuration but can effectively reduce both fecal coliform concentrations in oysters to safe concentrations (52) and reduce Vibrio spp. and viral loads in oysters (53-55).

Relay. Relay is basically defined as the act of passing something along from one group to another. The premise of relay in the shellfish industry is to transfer contaminated shellfish to an area where the contaminants are absent or at lower, safe levels and where the appropriate environmental conditions to cause reduction of the contaminants in the relayed shellfish are present. During relay, shellfish filter feed in a natural uncontrolled setting. The environmental parameters and the natural flora of the oyster and water influence the effectiveness of relay (41,

56-58). This less costly approach can be more effective than depuration in addressing viral particles, natural oyster flora, and fecal coliforms but is more time consuming than depuration

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(53, 54, 59) and environmental conditions can vary and still negatively affect outcomes. We do not know in full what environmental factors are key to successful relaying. As the level of consumer knowledge increases along with incidence of Vibrio-related illnesses, effective post- harvest treatments such as relay and depuration need further investigation using current technologies in order to address all facets of oyster contamination that make consumption risky.

Why is Vibrio illness from food consumption such a significant issue?

Death and sickness associated with food consumption are two factors that get the attention of regulators, consumers and the shellfish industry. In total, many more consumers become ill from food contaminated with other pathogens besides Vibrio spp. (60). Microbes such as Salmonella, norovirus, Campylobacter, Shigella and cause more hospitalizations than Vibrio spp. (61). These pathogens are more widespread via humans and contaminated foods and are ingested by a larger part of the total population. Oysters in particular are consumed by a much smaller portion of the total population than meat, poultry, produce, and other food stuffs where the aforementioned pathogens are typically found. What differentiates pathogenic Vibrio spp. epidemiology is that the large majority of foodborne come from a limited amount of food sources, so the approach for combatting illnesses is less complex than for other pathogens and foods, and thus theoretically achievable by industry and regulators.

Preventing the consumption of raw untreated oysters would address the large majority of illness that comes from consuming oysters (2). Since current legislation is heavily influenced by the financial motives of industry (33) and the desires of consumers, and it has not mandated measures to limit death and sickness from the consumption of raw untreated oysters, the scientific community is a logical source to provide information that will lead to acceptable

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answers. Research is needed that will help make known contaminant reduction strategies as effective and feasible as possible for the shellfish industry. Continued research towards verifying the effectiveness of novel and improved contaminant reduction and PHP strategies is needed to establish effective strategies that will be acceptable to all stakeholders. Here we look to understand some of the mechanism(s) within the relay process that reduce concentrations of

Vibrio spp., specifically Vibrio parahaemolyticus, from oysters.

Novel research to better understand effective relay

The microbiota in the oyster microbiome are diverse (62-65). The water where oysters reside can influence the oyster microbiome but the two tend to contain different microbial profiles (66). Estuarine waters are influenced by the ever changing environment (67), as are the oysters, and the fauna in the water (68-70) which are also affected by the myriad of microbiota associated with the water column (71). The water that flows into niches within the oyster introduces pathogenic and non-pathogenic microorganisms that colonize or pass through the oyster tissue (41, 72-75). Sediment, vegetation and food sources (plankton) found within the water harbor their own source of microbiota that have the potential to colonize the oyster (76) or interact transiently (75). Vibrios typically have strong adhesion capabilities to sediment and other marine animals (77). The ability to form biofilms (78) and affix to chitin particles (79) are some of the attributes that allow Vibrios to colonize the oyster better than other microbiota (41,

80). The natural association of Vibrios in oysters, the dynamics of the interaction between

Vibrios and other materials (planktonic bacteria, phytoplankton and zooplankton, etc.) (81), and the effect the environment has on all of this, are why Vibrio spp. have a strong affiliation in the niches of the oyster (56). Current efforts in the research community have focused on the addition

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of materials into post-harvest processes (82-85) to affect the affiliation between Vibrios and the oyster. Introduction of probiotics, phages and other materials have been a part of post-harvest treatment experimentation for many years (38, 86-90). The aforementioned experiments focus on specific antagonistic and competitive attributes of materials and microbes in vitro.

Our efforts differ from others in that we have the privilege of utilizing a salt pond located in Eliot, Maine that provides environmental conditions (elevated salinity and low or non-existent concentrations of V. parahaemolyticus) that can promote effective relay. We also are privileged to be able to collaborate with an established shellfish industry partner, Spinney Creek Shellfish

Company Inc., located on Spinney Creek, with this research. We were able to utilize this relay site to carry out experiments for four years during the warmer summer months to relay oysters contaminated with V. parahaemolyticus from the nearby Piscataqua River in New Hampshire to

Spinney Creek. Having the ability to perform relay year over year allowed us to reconfirm that relay can be effective (53, 54, 91) and obtain oyster and water samples from relay experiments for further analyses.

We hypothesize that changes in bacterial taxa within the oyster microbiome occurs during relay and influences the reduction of V. parahaemolyticus in relayed oysters. For the first time, that we are aware of, relayed oyster samples that resulted in the reduction of V. parahaemolyticus concentrations to safe concentrations (42) are being analyzed for microbial community dynamics to better understand how community profile changes in the oyster may assist in effective treatment processes for V. parahaemolyticus.

This dissertation is separated into two parts. Chapter Two captures four years of relay experiments in which we measured the temperature and salinity of harvest and relay waters along with the concentrations of V. parahaemolyticus in oysters at different time points during 14 days

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of relay. Chapter Three presents our findings on oyster and production area water microbiome dynamics and explains our approach to gain an understanding of the mechanism by which relay works. We used the oyster and water samples from the relay experiments to analyze microbial community shifts that take place during relay. Microbial community patterns were analyzed using 16S rRNA gene sequences to determine which genera were evident at day 0 and day 14 in the oyster and water samples from relay experiments. These community profile analyses along with other assays were used to inform some initial competition assays that serve as a step towards understanding the hypothesized competitive interaction of oyster microbiota during relay. We analyzed oyster samples directly from relay experiments to generate an understanding of which bacteria could be ideal candidates for potential use to outcompete naturally occurring V. parahaemolyticus in oysters. These findings can inform future work with microbes to reduce V. parahaemolyticus concentrations in oysters and help prevent human sickness.

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CHAPTER II

VARYING SUCCESS OF RELAYING TO REDUCE VIBRIO PARAHAEMOLYTICUS LEVELS IN OYSTERS (CRASSOSTREA VIRGINICA)

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INTRODUCTION

The shellfish aquaculture industry in the Northeast US has suffered increasingly more frequent Vibrio-associated disease illnesses and outbreaks over the past six years (4, 8, 92, 93).

Oysters are often eaten raw, thus exposing consumers to live microorganisms, including pathogenic Vibrio species that associate with oysters and reflect climate, harvest area and post- harvest environmental conditions and factors. There are a number of Vibrio spp. that are responsible for human illness and of these, Vibrio parahaemolyticus, Vibrio vulnificus and are responsible for most of the illnesses in the United States (4). V. parahaemolyticus is a leading cause of seafood-borne bacterial infections worldwide (94-97). In the U.S. there were 605 reported cases of illness from V. parahaemolyticus in 2014 (2) although due to underreporting (60) the number of cases of illness caused by V. parahaemolyticus each year is estimated to be ~45,000 (4).

The recent outbreaks of illness caused by V. parahaemolyticus in the Northeast US have coincided with changing climate conditions (8, 19, 98), resulting in economic and public health impacts, as well as instigating management control measures for shellfish production (93, 99-

101). Cost effective treatments that reduce levels of potentially pathogenic Vibrios in live shellfish would greatly benefit the shellfish industry. A variety of treatment strategies have been utilized in an effort to mitigate the levels of Vibrios in shellfish (34, 35, 102-104), including depuration and relay which are accepted strategies for reducing fecal-borne bacteria from shellfish (42). Previous studies have explored the use of both depuration and relay for Vibrio spp. reduction and several reported that depuration is ineffective in reducing Vibrio levels (45, 49,

105-108) yet more recent approaches have shown evidence that depuration significantly reduced levels of V. parahaemolyticus and V. vulnificus (47, 53, 54, 109).

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Relaying is a pre-harvest strategy of trans-locating oysters from contaminated growing waters to less contaminated waters to reduce microbial contaminant levels (42) and has shown promise for reducing Vibrios in several studies in different areas of the US (53, 91, 107, 110).

The effectiveness of V. parahaemolyticus reduction in oysters can be considered in regard to reference levels that imply risk thresholds. The V. parahaemolyticus level that shellfish post- harvest process treatments in the U.S. must reach to be considered effective is 30 MPN/g (42,

53). However, relaying is a different type of treatment because it depends on natural conditions and V. parahaemolyticus levels at the relay site, thus, the current Health Canada limit for minimal risk conditions, 100 MPN/g (111) that applies now to all imported and domestically produced raw oysters is probably more appropriate. These guidelines also take into account adequate pre- and post-harvest management measures to eliminate or reduce V. parahaemolyticus illness risk, including documentation that oyster meat temperature is below

15°C. The U.S. also accounts for pre- and post-harvest handling and temperature requirements; however, in the U.S. there is no upper limit for V. parahaemolyticus risk levels.

Coastal areas that support consistent and significant oyster production are also often areas with elevated levels of pathogenic Vibrios during seasonally warm conditions. Conversely, pathogenic Vibrios in non-production areas, typically offshore and characterized, in part, by higher salinity / lower temperature water are often absent or present at low levels (91, 110).

Location and cost of transport along with the need to obtain and maintain permits for harvest sites are factors that may be important to consider for selecting the best strategy to use to reduce the risk of V. parahaemolyticus illness. Thus, if suitable sites are near harvest sites, relaying may be cost effective if it is shown to be an effective strategy.

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In addition to environmental parameters such as salinity and temperature, the uptake and clearance activities of resident and transient microbiota by the oysters affect the levels of Vibrio spp. found in oysters (74, 75, 112-114). The oyster bacterial community is influenced by environmental conditions (57) yet, like other marine animals (115), the composition of the bacterial community within the oyster is not as influenced by dominant taxa in the surrounding water as much as it is by the competitive interactions within the oyster microbial community

(116). Thus, although Vibrio spp. are resident flora of oysters and other marine shellfish (94), they and other oyster microbiome taxa are influenced by a complex interaction of environmental conditions, especially temperature, salinity, as well as biological factors (113, 114) within the oysters and in the oyster bed habitat.

Translocation of oysters generates shock to the oyster which in turn necessitates an acclimation period for the oyster to stabilize (117). Other studies have shown that a minimum of

7 days are needed for successful relay (53, 91, 118) which in part probably reflects the time needed for the oyster to stabilize after transfer to a new body of water. Taking into account previous work, we hypothesize that relay of oysters to a body of water with elevated salinity and low V. parahaemolyticus levels will result in oysters with acceptable levels of V. parahaemolyticus.

The Great Bay Estuary (GBE) of New Hampshire and Maine, USA (Fig 1.) is an area where there has for many years been extremely rare reported Vibrio disease from local shellfish, despite long-term detection of pathogenic Vibrio spp. (119-121). Environmental conditions in the

GBE are spatially variable and climatic conditions (i.e., temperature, rainfall) vary seasonally with water temperature extremes of <0°C to ~30°C (19), making this estuary an ideal site for studying conditions associated with variation in Vibrio populations (19, 122, 123). Long cold

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winters cause most pathogenic Vibrio spp. to become undetectable in shellfish (70, 124), and they re-emerge in increasing levels in shellfish as conditions warm. Vibrio spp. are consistently detected in many areas of the estuary during June to September when water temperatures are warmer (19) with some exceptions, like Spinney Creek , a small tidal pond that has very little freshwater inflow and a tidal dam at the mouth that allows high tide salt water to enter, creating elevated salinity conditions. Historically, at Spinney Creek, Vibrio levels have consistently been extremely low or absent (110) even though water temperatures can be elevated relative to surrounding waters. This makes Spinney Creek a uniquely suitable site for this study because it is not only on the border between cold saline waters with few Vibrios and warm, lower salinity estuarine waters where Vibrios are prominent, it is also in close proximity to local oyster farms and is itself a shellfish production area.

Fig. 1. The Great Bay estuary of Maine and New Hampshire, USA, with harvest site (PR) in the Piscataqua River and the relay site (SC) in Spinney Creek.

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The objective for this study was to determine the time and environmental conditions that are needed for successful reduction of V. parahaemolyticus levels in oysters relayed from the

Piscataqua River to Spinney Creek. Our findings reflect both successful and ineffective relaying to reduce V. parahaemolyticus levels in oysters. Testing the effectiveness of relaying oysters containing resident microbial communities and V. parahaemolyticus populations in a body of water with different resident microbial communities, consistently lower V. parahaemolyticus levels and higher salinity over multiple sampling seasons is, in addition to informing management of V. parahaemolyticus risk levels in shellfish, a step towards understanding the ecosystem conditions that cause variation in oyster V. parahaemolyticus levels. This study is foundational work to gain an understanding of interactions between V. parahaemolyticus and the total oyster microbiota.

RESULTS

Seasonally variable environmental conditions. Environmental conditions in the relay and harvest waters were measured at the start of relay experiments from 2011 to 2014 (Fig. 2 A+

B). Water temperatures exhibited typical seasonal patterns, trending in parallel for both sites and consistently conducive to V. parahaemolyticus presence (>15°C) during June-September each year. Salinity did not exhibit any seasonal trends and trends for each site were different, with relatively consistent and higher salinity at Spinney Creek and more variable and lower salinity at the Piscataqua River. Water at Spinney Creek was consistently, though not significantly (p>0.05) warmer with an average difference of 1.9oC between the sites. Spinney Creek also had significantly higher salinity levels (p < 0.001) with an average difference of 12 parts per thousand (ppt). Water temperature differences on relay sampling dates ranged from 0.5 oC (June

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2013) to 4.7 oC (June 2014) between the sites, and differences in salinity ranged from 3.4 ppt

(July 2012) to 19.1 ppt (September 2013). Levels of rainfall from three days prior to the day of sampling in the area near the sites did not significantly correlate to salinity levels at Spinney

Creek (Fig. 2A+B).

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Fig 2. Water temperature (°C) and salinity (PPT-parts per thousand) at the relay (SC) and harvest (PR) sites. A) Water temperatures at SC and PR on Day 0 of relay. B) Salinity levels at the sample and relay sites on Day 0.

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Background V. parahaemolyticus levels at Spinney Creek and Piscataqua River.

Oyster and water samples collected from 2011 to 2014 during June to November were analyzed for V. parahaemolyticus levels (Fig. 3). The geometric mean V. parahaemolyticus levels in the

Piscataqua River oysters from June to September was 121 V. parahaemolyticus/g, with a range from 13 to 10,100 V. parahaemolyticus/g for all samples and from 374 to 10,100 V. parahaemolyticus/g for July and August samples. V. parahaemolyticus levels were consistently greater than the target reduction level (100 MPN V. parahaemolyticus/g oysters) during July and

August, providing a source of oysters with elevated levels of V. parahaemolyticus that were always conducive to relaying experiments. In some years V. parahaemolyticus levels were also adequate for relay experiments during earlier and later months, and relay experiments were conducted in most of the warmer months when V. parahaemolyticus levels were most likely to be elevated in Day 0 samples (Fig. 3A). Water from Spinney Creek and Piscataqua River had consistently low V. parahaemolyticus levels (Fig. 3B), with the exception of some samples from

2013. V. parahaemolyticus levels in relay water samples from Spinney Creek did not rise above

120 MPN V. parahaemolyticus/mL except during July 2013. The consistently high salinity levels in conjunction with historically low levels of Vibrio spp. at Spinney Creek supported its selection as an appropriate test site for relay experiments.

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Fig 3. V. parahaemolyticus levels (MPN/g) on Day 0 of relay experiments in oysters from PR and water samples from PR and SC. A.) V. parahaemolyticus levels (MPN/g) in oysters collected at Day 0 of relay at PR. The median and the 25th and 75th percentiles are shown with error bars. Dashes mean that only one oyster sample was analyzed at Day 0. B.) V. parahaemolyticus levels (MPN/mL) in SC and PR water on Day 0 of relay.

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The effect of relay on V. parahaemolyticus levels. Relaying experiments were conducted during 2011-14 using oysters from the Piscataqua River that were naturally contaminated with V. parahaemolyticus. Levels of V. parahaemolyticus in the Piscataqua River and Spinney Creek water were not statistically different (p = 0.42), even though the sites were different in regards to temperature and salinity. Overall V. parahaemolyticus levels in the

Piscataqua River and Spinney Creek water were significantly lower (p <0.001) than V. parahaemolyticus levels in the oysters from the Piscataqua River (Table 1).

Table 1. Statistical analyses of V. parahaemolyticus (Vp) levels (MPN/g or mL) in oyster and water samples analyzed during relay experiments from 2011 to 2014. *Vp Standard Year Source p Value MPN Deviation PR Water 23.3 5.9 All 0.42 SCS Water 17.8 8.1 Oysters 121.3 25.2 All <0.001 Water 18.9 7.5 All Oyster Day 0 181.2 12.7 All All Oyster Day <0.05 51.7 14.9 14

Day 0 Oysters 83.4 5.6 2011 0.12 Day 14 Oysters 29.7 3.4 Day 0 Oysters 130.2 16.6 2012 <0.01 Day 14 Oysters 7.4 5.2 Day 0 Oysters 436.4 12.6 2013 0.28 Day 14 Oysters 2258.8 11.2 Day 0 Oysters 483.9 21.5 2014 0.94 Day 14 Oysters 132.0 8.9 *Geometric mean of all samples from the respective source

The overall Day 0 and Day 14 geometric mean V. parahaemolyticus levels in oysters were significantly different (Table 1). Comparison of relay success for all samples from each

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year show that there were overall reductions in V. parahaemolyticus levels from Day 0 to Day 14 during 2011, 2012 and 2014, but not during 2013, when the geometric mean V. parahaemolyticus levels increased from Day 0 to Day 14. For 2011 there was a lower overall level of V. parahaemolyticus on Day 14 compared to Day 0 but the difference was not statistically significant, whereas in 2012, Day 14 levels were significantly less than Day 0 levels

(Table 1). July and August 2012 water temperatures and salinity levels during August and

September 2012 were the highest recorded for this study (Fig. 2 A + B). There was a non- significant increase in the overall geometric mean V. parahaemolyticus levels from Day 0 to Day

14 during 2013, and in 2014 there was a non-significant decrease in V. parahaemolyticus levels from Day 0 to Day 14. Thus, the consistently successful relaying experiments in the first two years set up the third and fourth years where the results were unexpectedly unsuccessful. The reason for this change in relay success, however, was not obvious from environmental conditions, i.e., water temperature and salinity trends.

More in-depth interpretation of individual relay experiments provides more information on the relative success of relaying oysters for reducing V. parahaemolyticus levels. Overall, V. parahaemolyticus levels were reduced during 14-day relay in 9 of the 14 monthly trials, and target reduction levels (<100 V. parahaemolyticus/g) were achieved in 9 of the 14 monthly trials

(Fig. 4). The greatest reduction occurred in July 2012 and August 2014. In terms of target reduction levels, three of the experiments started with V. parahaemolyticus levels <100 V. parahaemolyticus/g on Day 0 and the V. parahaemolyticus levels in the September 2012 experiment were < 100 V. parahaemolyticus/g on both Day 0 and 14. In five of the ten trials where samples were analyzed between Days 0 and 14 (2, 7, or 10 days), V. parahaemolyticus levels in the oyster samples increased during the intermediate sampling days, suggesting that

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relaying to Spinney Creek for greater than 10 days is necessary to avoid otherwise inconsistent results and to allow for effective V. parahaemolyticus reduction.

Fig 4. V. parahaemolyticus levels (MPN/g) in oyster samples analyzed during each sampling day of relay experiments from 2011 to 2014. The median and the 25th and 75th percentiles are shown with error bars. No relay experiment took place where month-year boxes are blank. Days of relay with V. parahaemolyticus levels above 100 MPN V. parahaemolyticus/g are colored red, and below 100 MPN V. parahaemolyticus/g blue. X axis = sampling day during relay.

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V. parahaemolyticus levels were, to varying degrees, higher after 14 days in 5 of the relay trials. V. parahaemolyticus levels were higher after 14 days in 2 out of 3 of the trials that had low

(<100 V. parahaemolyticus/g) levels at Day 0 (September 2011 and June 2014), and the Day 14 levels were also relatively low (<102 V. parahaemolyticus/g). In 2013 two other relay experiments resulted in substantially increased V. parahaemolyticus levels after 14 days (Fig. 4).

V. parahaemolyticus levels increased in relayed oysters in June and July, 2013 to unusually elevated levels (>106 MPN V. parahaemolyticus/g in one sample). During September and

October 2013, however, relaying again reduced V. parahaemolyticus levels by factors of 32x and

15x, respectively, even though the final levels after 14 days in the September 2013 trial was 290

MPN/g, which is above the target reduction level. V. parahaemolyticus levels in relayed oysters increased after 14 days again in 2014 during June and July, but V. parahaemolyticus levels in relayed oysters decreased by a factor of 49x during the August 2014 relay experiment (Fig. 4).

In 2011, 2012 and 2014 no cultured V. parahaemolyticus isolates contained trh and tdh , but in 2013 four isolates, from oysters collected on Days 0 and 14 in June and Day 7 in

July, did contain the trh marker. The detection of trh and the increased V. parahaemolyticus levels collectively suggest that a shift occurred in both the V. parahaemolyticus population and the microbial community despite water conditions that were, other than much lower salinity levels at the Piscataqua River during July and September 2013, within the range of environmental conditions compared to previous years. Even though 2014 strains did not contain tdh and trh genes, the microbial community in the oyster may have retained some characteristics from what was present under 2013 conditions that interfered with relay success in June and July

2014.

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The harvest study site routinely had V. parahaemolyticus levels that were > 100 V. parahaemolyticus/g yet not high enough to demonstrate the potential for multi-log V. parahaemolyticus reductions during relay. To enable this, naturally contaminated oysters from the August 2012 trial were temperature abused to increase V. parahaemolyticus levels from 178 to 9.3 x 104 V. parahaemolyticus/g. There was a 4-log reduction of V. parahaemolyticus in the temperature-abused oysters to 3 V. parahaemolyticus/g after a 14-day relay. The non- temperature abused Day 14 oyster samples from this same date reached the same level demonstrating the potential for successful V. parahaemolyticus level reduction to target levels after 14 days of relay even with highly contaminated oysters.

DISSCUSSION

The results of this study suggest that relaying oysters to reduce V. parahaemolyticus levels in oysters may be a viable management strategy to reduce the risk of illness to oyster consumers. The first two years of the study confirmed previous study results that reported consistent success with oyster relaying for this purpose. The relative overall success of relaying, i.e., reducing V. parahaemolyticus levels and reducing them to <100 V. parahaemolyticus/g, in this study was greatly impacted by unexpected results in 2013 and again in early 2014 trials.

There is no obvious explanation for this change in relaying success at this site, including the influence of the limited array of measured environmental conditions, yet there are some plausible explanations related to microbial community dynamics (75, 96, 125).

Successful relaying of live oysters to reduce V. parahaemolyticus levels requires the right conditions. These include lower V. parahaemolyticus levels at the relay site compared to the harvest site, and conditions, often associated with elevated salinity, for the oysters to interact

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with microbiome constituents in the overlying water microbial community in a way that results in decreasing V. parahaemolyticus levels. Studies at the same sites used in this study have shown this to be possible (53, 54, 107, 126). These relay studies and some depuration studies suggest that certain environmental conditions may be important, including water temperature (36, 106,

109), elevated salinity (53, 54, 91) and biotic factors (91, 127). In fact, relay to higher salinity waters to reduce V. parahaemolyticus levels in oysters has recently been adopted as a strategy to reduce pathogenic V. parahaemolyticus strain levels in Massachusetts, and appears to be successful (32). Optimal salinity levels at relay sites for effective relaying have not been determined and may be geographically variable (128). Even though there was an overall decrease in V. parahaemolyticus levels during the consistent 14 day relay trials in this study, the range of salinity (26.5 – 31.2 ppt) at the relay site was variable, especially during 2013, and may have been a factor in the varying success of the relay compared to the more consistent salinities for other study areas where relaying was more consistently successful (53).

The variable success in relaying in this study is of great interest as it suggests that only some conditions and, potentially, as yet undetermined biological conditions like the oyster microbiome taxa may be critical to successful relaying (110). Modeling Vibrio spp. population dynamics and environmental conditions in a shellfish harvesting area to identify not only high but also low risk conditions (19) that would be conducive to shellfish relay success would be a key part of this transferable strategy to reduce V. parahaemolyticus levels in live oysters. Thus, relay to generally higher salinity waters that may be in much closer proximity to production areas than the open ocean, is worth further investigation to determine how to achieve consistent reductions in V. parahaemolyticus levels with minimal requirements for controlled conditions.

This study provides findings to support the possibility that this approach could be successful,

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although the change in success during 2013 and 2014 also warrants further study. When relaying experiments were successful in reducing V. parahaemolyticus to below target levels, it appeared to require 10 to 14 days. Reductions in V. parahaemolyticus levels were at best inconsistent for shorter relay times, perhaps due to a required acclimation by the oysters to the disturbance caused by relaying them to different water conditions (129-131).

During 2013, V. parahaemolyticus levels in Spinney Creek water were abnormally high for the first time since the onset of environmental monitoring in the 1990’s (107), and were even higher than levels at the Piscataqua River oyster sampling site, while salinity levels at the relay site in 2013 were comparable to 2011 and 2012. It was not until the arrival of colder waters in

November 2013 that V. parahaemolyticus levels in the creek water, and in relayed oysters, dropped to more typical low levels. Record high regional sea surface temperatures in 2012 that carried over into 2013 (98) and coincident effects on regional coastal ecosystems (132) may have been a significant catalyst underlying the 2013 anomalous V. parahaemolyticus levels. The elevated V. parahaemolyticus levels in the creek’s water column generated concerns about a potential shift in the creek water microbiome. A shift in environmental conditions and the water microbiome could have affected the oyster microbiome (74, 133, 134). Relaying remained unsuccessful again in early (June-July) 2014 when regional sea surface temperatures returned to more average and cooler levels, though the August 2014 relay was successful, which suggests that there may have been a residual effect from 2013 remaining in 2014 that influenced relay success.

V. parahaemolyticus isolates from the GBE with detectable clinical markers were rarely evident for six years (2007-2012) of surveillance studies leading up to 2013 (8). In 2012-2013, total V. parahaemolyticus levels in Great Bay oysters were higher than in the previous 5 years

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(19), and four environmental isolates from 2013 relayed oysters were identified as trh positive, suggesting that the emergence of potentially pathogenic populations in the Great Bay coincided with increased total Vibrio levels. Previous identification of potentially pathogenic V. parahaemolyticus strains (trh+) has been rare, with one isolated from Great Bay estuary water in

2008 and one from oysters in 2009 (8). The correlation between the presence of pathogenic strains of V. parahaemolyticus and the anomalous environmental conditions in 2012-13 is aligned with results from other published work in that the increase in the incidence of disease is often correlated to shifts in environmental conditions (14, 96, 135). Shifts in microbial community taxa induced by regional climate changes could also affect relay success in historically effective relay sites.

Post-summer oyster relay experiments in cooler water were carried out each year to provide data confirming the drastic seasonal decline in V. parahaemolyticus levels in this region.

At the end of each season, even after seeing record highs in water and oyster levels in 2013, V. parahaemolyticus levels dropped to undetectable MPN/g levels, with water temperatures of 9.5,

13, and 3.9 oC for December 5, 2011, October 25, 2012, and November 21, 2013, respectively.

These decreases in V. parahaemolyticus levels in cooler months highlights the influence of environmental conditions on the persistence of Vibrio spp. in oysters (96) and supports the potential for artificially recreating these conditions to enhance reductions during warmer higher risk seasons (41, 106, 109). For this more intensive relay strategy, some production areas may have cooler water areas nearby, although many would probably require cooling and maintaining low water temperatures, an energy intensive strategy, at on-land facilities. The same would hold true for any requirement to artificially increase and maintain a specific high salinity condition.

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The mechanisms behind the effectiveness of relay remain largely unknown. One theory is that microbial interactions play a role in reducing levels of Vibrio spp. in oysters (127, 133).

Current thoughts are that condition-specific bacteria that reside within the oyster are responsible for the displacement of V. parahaemolyticus from the oyster (74). Ongoing research involves analyzing the genomic sequence data of community members in the oyster and water samples obtained from the relay experiments described in this study to test these hypotheses and inform further research.

MATERIALS AND METHODS

Oyster harvest and relay. Oysters (Crassostrea virginica) were harvested by diving or tonging from an oyster bed in the Piscataqua River (43°10’08.49”/70°49’42.54”), Dover, New

Hampshire during June to October in 2011 to 2014, and relayed to Spinney Creek , Eliot, Maine

(43-05'48''/070-45'58'') (Fig. 1). Sampling day water column conditions (temperature and salinity) were measured at the harvest and relay sites using a YSI 85 meter (YSI, Yellow

Springs, OH). Enough oysters were harvested and transported to allow for analysis of V. parahaemolyticus levels in three separate sub-samples of 12 animals (homogenate) at each sample time. Water samples (500 ml) were also collected to measure V. parahaemolyticus levels in the harvest and relay waters. Oyster samples were transported from the sampling site to the relay site within two hours of collection. Samples were kept in a cooler within small baskets, separated from ice packs, during transportation.

During June to October over a four-year period (2011-2014), samples were analyzed at set intervals (0, 2, 7, 10 and 14 days) during the relay experiments. Oysters were suspended in the water column of Spinney Creek within a basket attached to a mooring ~1 m from the water

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surface. On each sampling day, oysters were removed from the basket and transported to the lab for bacteriological analyses.

Oyster processing. Triplicate oyster meat homogenates for each sampling day were analyzed for V. parahaemolyticus using detection methods described in the US FDA

Bacteriological Analytical Manual with modifications (136). Each oyster was scrubbed using a metal brush and shucked to dispense the meat and liquor into a sterile beaker. Homogenate meats and liquors were weighed and then diluted 1:2 (w:v) with alkaline peptone water (APW) and the contents were transferred to a sterile stainless steel blender container and homogenized for a total of 90 sec. An aliquot (20 ml) of oyster homogenate was added into 80 ml of APW.

One mL of oyster homogenate was added into three separate tubes containing 9 mL of APW.

The APW tubes were then serially diluted 10-fold down to 10-6. All dilutions were incubated at

37°C overnight. Aliquots of the leftover oyster homogenate replicates (~40 mL) were transferred into a sterile 50 mL conical tube and stored at -80°C. In 2012, twelve oysters from the August relay sample collection were incubated for 24 h at 28oC to generate higher V. parahaemolyticus levels in naturally contaminated oysters using thermal abuse (137). Temperature abused oyster samples were relayed in Spinney Creek for 14 days and processed, post relay, in the same manner as other oyster homogenates.

V. parahaemolyticus detection - qPCR. V. parahaemolyticus levels in oyster homogenates and water samples were calculated using a Most Probable Number procedure (138) in combination with real time PCR (139). Briefly, APW tubes were scored for turbidity, an aliquot of each turbid APW tube and water sample was transferred into a sterile 1.5 mL centrifuge tube, boiled for 10 mins at 100oC, and spun down at 8000 RPM in a tabletop micro- centrifuge for 5 min. The qPCR was performed in 25 µL reactions containing OmniMix

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Mastermix (Cepheid, Sunnyvale, CA), 75 nM tlh primers, 150 nM tlh TaqMan probe, MgCl2

(Bio-Rad Laboratories, Hercules, CA) final levels 5 mM, 1µL of an internal amplification control (IAC) (BioGX, Birmingham, AL), 75 nM IAC primers, 150 nM IAC probe, and 4 ul of template DNA (boiled oyster homogenate or water). The qPCR parameters in the Cephied Smart

Cycler (Cephied, Sunnyvale, CA) included a hot start step at 95°C for 3 minutes followed by 45 cycles of amplification with a denaturation step at 95°C for 5 sec and annealing/extension at

59°C for 45 sec. The fluorescence readings, dye set, and manual threshold units followed

Nordstrom et al. (139). Vibrio isolates from oyster homogenates that were confirmed as tlh positive were used as positive controls, and nuclease free water served as the no template negative control.

Culture PCR and hemolysin gene detection. In parallel to qPCR, turbid MPN tubes were streaked for isolation onto CHROMagar Vibrio (DRG-International Inc., Springfield, NJ) for V. parahaemolyticus isolation and differentiation at 37°C. Culture plates were incubated for

18 – 20 hrs. Purple isolates evident on CHROMagar Vibrio plates were screened as V. parahaemolyticus using multiplex PCR. Selected phenotypes were streaked for isolation onto tryptic soy agar (TSA) (Thermo Fisher Scientific Inc., Waltham, MA); plates were incubated at room temperature overnight. Isolated colonies on the TSA were used to create broth cultures by transferring one isolate into 3 mL of heart infusion broth (Becton, Dickinson and Company,

Franklin Lakes, New Jersey). Cultures were incubated statically at 37°C, overnight. An aliquot of the overnight culture was transferred to a sterile 1.5 mL micro-centrifuge tube, boiled at

100°C for 10 minutes, and spun down at 8000 RPM for 5 mins to be used as a template for culture PCR.

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IQ Supermix (Bio-Rad, Hercules, CA) was used, following manufacturer’s instructions, to create a mastermix for V. parahaemolyticus culture PCR. V. parahaemolyticus isolates were screened for thermolabile hemolysin (tlh), thermostable direct hemolysin (tdh), and TRH-related hemolysin (trh) genetic markers using primer sets and PCR parameters reported by Panicker et al. (140) in 15 µL reactions. Each PCR included a positive control of V. parahaemolyticus F11-

3A (tdh+/trh+ environmental isolate) gDNA and a no template control using nuclease-free water.

PCR amplicons were visualized using 1.0% agarose (SeaKem LE Agarose, Lonza, Portsmouth,

NH) with GelRed (PHENIX Research Products, Candler, NC).

Statistical analyses. Statistical analyses using log10-transformed data and geometric means of MPN data were carried out using the R package, version 3.2.2 (https://www.r- project.org). The Wilcoxon rank sum test in the R package was used to test the null hypothesis that after 14 days of relay V. parahaemolyticus levels in oysters are not significantly different than Day 0 oyster V. parahaemolyticus levels.

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CHAPTER III

OYSTER MICROBIAL COMMUNITY CHANGES ASSOCIATED WITH VIBRIO PARAHAEMOLYTICUS CONCENTRATIONS IN RELAYED OYSTERS

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INTRODUCTION

Human illness associated with shellfish consumption is a growing public health issue and a significant concern to the shellfish industry. A variety of bacterial pathogens may reside in oysters, yet only some species pose significant threats to humans. Vibrio parahaemolyticus is at the top of bacterial safety concerns for public health officials and the shellfish industry, as it is the leading source of bacterial illnesses from seafood consumption in the United States (60). In the Northeast US, outbreaks of illness caused by V. parahaemolyticus have become much more prevalent over the past 5 years (8, 19) resulting in economic and public health impacts as well as instigating management control measures. Effective control measures must address the effects of environmental and climatic conditions on the ecosystem sources (estuarine waters, sediments, shellfish, plankton and other biota) of V. parahaemolyticus. Many ecosystem matrices harbor different concentrations of Vibrio species during the different months of the year (141). Here we focus on relaying, a strategy involving translocation of oysters from areas with elevated concentrations of Vibrio spp. to areas with low Vibrio concentrations (53, 91, 107) and thus minimize public health concerns for oyster consumers in the Northeast (8).

Oysters are an influential species in many estuaries, as their prolific filter feeding affects the microbiota within them and the surrounding waters (39, 142). In turn, the variable environmental conditions in estuaries can influence the microbiome of the oyster (72, 129).

Dispersion of bio-accumulated and colonized bacteria from an oyster is affected by temporal, episodic and directional changes within the surrounding waters (116, 129, 130, 143, 144). These factors are natural disturbances that play a role in the composition of the microbial community

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(145). Additionally, environmental factors such as temperature and salinity can also cause shifts in oyster microbiomes (114, 141, 143).

Environmental conditions drive selection and abundance of microbial populations in aquatic ecosystems (146). Water temperature is a dominant factor associated with Vibrio populations, especially in cooler climates where they can become undetectable in winter and re- appear in warmer summer conditions when most human illness occurs (81). During the summer, salinity can be a significant factor affecting pathogenic Vibrio spp. concentrations in oysters and the overlying water (57, 118, 119). When salinity levels are elevated, the concentrations of some

Vibrio spp. in oysters can decrease (58, 128), and they can increase under lower salinity conditions (115). Salinity is a factor used in strategies for reducing concentrations of pathogenic

Vibrio spp. in shellfish, including relay (53, 118).

A key consideration in the development of effective strategies to reduce pathogenic

Vibrio spp. in shellfish is the influence of resident microbiota in natural seawater. Water treatment processes that remove natural microbiota (e.g. UV disinfection and filtration) can diminish the effectiveness of relaying for reducing concentrations of Vibrio spp. (105). Space limitations, microbial community interactions and a constant inflow of exogenous material during the filter feeding process can also contribute to microbiome changes within oysters (75,

134, 145, 147). Bacteria associated with filtered food and other particles can become enriched in shellfish tissue by associating with mucus membranes and other available internal surfaces (75,

148, 149). It is not clear if bacteria from surrounding water can actually colonize oysters (141) or are transient residents (75). Those bacteria that do become residents in oysters may also competitively exclude other bacterial species because they are either better suited for the oyster environment (150, 151) or they possess properties that antagonize other bacteria (38, 66, 89,

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113). This exclusion process may affect V. parahaemolyticus populations in oysters and is at the core of this study.

In this study we used an established protocol (152) to investigate the hypothesis that shifts in the microbial inhabitants of the eastern oyster (Crassostrea virginica) during relay are correlated with the reduction of V. parahaemolyticus levels. We observed consistent shifts in oyster microbiomes resulting from the 14-day relay through the summers of 2011 and 2012 when

V. parahaemolyticus levels decreased. In addition, when V. parahaemolyticus levels increased in relayed oysters during 2013, we again observed taxa shifts in oyster microbiomes, however, they were inconsistent and the taxa were completely different from those observed in 2011-12. Our study focused on the co-variation of oyster resident microbiota and V. parahaemolyticus for future in-depth studies of the underlying mechanisms that affect both V. parahaemolyticus populations and microbiomes in shellfish and harvest areas. A better understanding of pathogen- community interaction may lead to better control of human illness and outbreaks that are linked to raw oyster consumption.

RESULTS

Relay of oysters from the Piscataqua River to Spinney Creek successfully reduced V. parahaemolyticus concentrations during 2011-12, but was less successful in 2013. Relaying contaminated oysters from the Piscataqua River to Spinney Creek, a location with historically lower V. parahaemolyticus abundance for 14 days, is a potential strategy for reducing V. parahaemolyticus concentrations to safe levels for human consumption of raw oysters. We hypothesize that during the relay process, V. parahaemolyticus reduction is accompanied by changes in other microbial community members and that some of these may participate in

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pathogen displacement. Therefore, we conducted 11 relay experiments between 2011 and 2013 to obtain quantitative data on V. parahaemolyticus concentrations and in parallel archived samples to evaluate changes in microbial community composition. V. parahaemolyticus concentrations in relayed oysters after less than 14 days were highly variable (131). V. parahaemolyticus concentrations decreased after 14 days of relay in 7 out of 11 relay experiments (Table B.1). Interestingly, three out of four experiments where V. parahaemolyticus actually increased after relay occurred in 2013. We hypothesize that the high success of relay in

2011 and 2012, and subsequent lower success in 2013 reflect differences in inter-annual microbial community composition that could have been driven by the anomalously warm summer and mild winter of 2012 and 2013 (14, 19, 98). The mean monthly sea surface temperatures (SST) recorded during the months of February-August (except April) during either

2012 or 2013 remain the highest for the 17-year record at the A01 buoy in nearby Massachusetts

Bay (http://neracoos.org/datatools/climatologies_display). The mean monthly temperatures for cooler months during this time period are also above the mean for 2001-16. The overall trends for SST data at the A01 buoy and in the Great Bay estuary show steady increases from 2005 through 2012-13(8).

Assignment of oyster and water taxa by operational taxonomic units (OTUs). V4 16S rRNA gene sequence amplifications from DNA of sixty-eight individually barcoded samples, including relayed oysters (n=35) and water (n=33), were sequenced. An average of 65% of the bases from the detected sequences reached a Q score of 30 (99.9% base call accuracy) (Table B.2). After quality filtering and processing (see Methods), 12,115,279 out of 21,763,447 million total paired end reads were clustered into 4,454 Operational Taxonomic Units (OTUs) at 97% sequence identity, with an average of 178,165 (+/- 23,640 s.d.) sequences/sample. Each sample had over

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100,000 sequences, which is considered adequate depth to capture a portion of the community diversity in the oyster and water samples, using shorter reads from the Illumina sequencer (152).

Rarefaction curves for both oyster and water samples indicated sequencing depths did not capture full diversity so conclusions about taxa analyses are considerate of this (Fig. 1A). Six- hundred thirteen genera were identified in the 68 samples and 194 of these genera were classified at the species level. Three of the OTU’s were unclassified. The shorter reads generated by

Illumina sequencing make the identification of rare species more difficult (152), but the depth of taxa identification was sufficient for analyzing bacterial composition changes in oyster and water samples.

Fig. 1. Alpha rarefaction curves for sequences within the observed OTUs. A) All 68 individual samples B) Aggregated by sample type.

Bacterial community composition changed during oyster relay experiments that successfully reduced V. parahaemolyticus concentrations. Paired 0 and 14 Day relayed oyster samples with the largest decrease in V. parahaemolyticus concentrations during relay were used

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to assess changes in microbiome composition (N=4, 1 pair for 2011 and 3 pair for 2012) (Table

1). We first determined if all identified Vibrio taxa co-varied with V. parahaemolyticus concentrations by decreasing in abundance between Day 0 and 14. We hypothesized that all

Vibrio spp. would not respond to relaying the same as V. parahaemolyticus, based in part on previous studies that show variations in V. parahaemolyticus populations in oysters do not reflect trends for the total Vibrio (153, 154). The V. parahaemolyticus concentrations (MPN/g) decreased ~10 fold in oysters from July 2011 while total Vibrio concentrations (sequence abundance counts/g) slightly decreased after 14 days (Table 1). In the 2012 oyster samples with significantly reduced V. parahaemolyticus concentrations we observed substantially increased concentrations of total Vibrio spp. on Day 14 compared to the Day 0 samples. Overall, there was a negative relationship between the concentrations of V. parahaemolyticus and total Vibrio spp. concentrations in all samples (Fig. 2) that was significant (F-test; p<0.001), which is in line with our hypothesis. For every 1% increase in rRNA gene-derived Vibrio genus counts, the V. parahaemolyticus MPN counts decreased by 0.55%. This negative relationship between V. parahaemolyticus and other Vibrio spp. suggests that other Vibrio spp. may have an effect on V. parahaemolyticus concentrations in relayed oysters in water with elevated salinity, although it is also possible that salinity changes during relay are simply more conducive to the growth of these other Vibrio spp.

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Table 1. V. parahaemolyticus concentrations and 16S sequence abundance for Vibrio spp. in paired Day 0 and 14 oyster samples from the four most effective relay experiments: 2011-12.

2011 *Vibrio **Vp Oyster (16s) (MPN/g) Day 0 9 428 July Day 14 5 14 2012 *Vibrio **Vp Oyster (16s) (MPN/g) Day 0 27 103 June Day 14 131 9 Day 0 5 4115 July Day 14 2044 40 Day 0 2 179 August Day 14 944 3 *Total sequence abundance counts that classified as Vibrio **Geometric means of V. parahaemolyticus MPN/g concentrations of triplicate oyster samples

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Fig. 2. Linear regression analysis between V. parahaemolyticus MPN concentrations and Vibrio spp. rRNA gene abundances in all samples. The effect size coefficient (-0.54591) describes the X, Y change correlation. Adjusted R2 (0.3233) explains the variance of the dependent variable (V. parahaemolyticus MPN) to the fitted regression line.

We then analyzed the same four relayed oyster sample pairs to explore differences in all identified bacterial taxa between Day 0 and 14 oyster microbial communities. Two OTUs were unclassified in these eight oysters (representing 0.08% of all sequences in these samples), 471 genera were able to be identified (35% of total sequences) and of these, 144 genera were further identified to the species level (7% of total sequences). We hypothesized that taxa that are relatively more abundant may have greater influence on V. parahaemolyticus concentration reduction during relay, so relatively rare singleton and doubleton OTUs were removed to yield

365 of the 471 genera. Thirty-three (9%) out of the 365 identified genera increased in relative

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abundance (Table 2) and 332 (91%) genera decreased in relative abundance during relay. Of the

33 genera that increased, Vibrio, , Phaeobacter, Corynebacterium and Shewanella were the five most abundant based on relative sequence abundance.

Interestingly, the five most abundant genera encompassed a major fraction (84%) of the total number of sequences identified at the genus level (for the 8 samples) that increased in response to oyster relaying. However, not all these increases were significant. Of the thirty-three genera that increased in abundance, only Corynebacterium and three other genera

(Porphyromonas, Anaerococcus and Verrucomicrobium) increased significantly during 14-day relay (Student’s t-test, p= 0.003, 0.02, 0.008 and 0.03 respectively). The relative abundance for sequences in OTUs identified as these four genera were relatively low compared to the total microbial community (3.5, 0.11, 1.0 and 0.01%, respectively), and accounted for 15% of the total sequence abundance of the 33 genera that increased during relay (Table 2). On the other hand, the Vibrio genus, which increased during relay but not significantly (p= 0.52), represented 21% of sequences of the 33 genera that increased in abundance during relay (Table 2).

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Table 2. Identified genera that increased from Day 0 to Day 14 in oysters from the most effective relay experiments (see Table 1).

*Weighted Total *Weighted Total Genus OTUs Genus OTUs Abundance Abundance Abundance Abundance

Vibrio 1.40% 6.98% 5 Campylobacter 0.02% 0.080% 4 Mycoplasma 0.85% 14.40% 17 Dialister 0.02% 0.080% 4 Phaeobacter 0.26% 1.04% 4 Dietzia 0.02% 0.070% 4 Corynebacterium** 0.18% 3.52% 20 Desulfovibrio 0.01% 0.090% 6 Shewanella 0.16% 1.30% 8 Persicirhabdus 0.01% 0.050% 4 Fusobacterium 0.16% 0.63% 4 Helcococcus 0.01% 0.080% 6 Octadecabacter 0.08% 0.30% 4 Verrucomicrobium** 0.01% 0.100% 8 Pseudoalteromonas 0.07% 0.14% 2 Porphyromonas** 0.01% 0.110% 10 Peptoniphilus 0.07% 0.73% 11 Flavobacterium 0.01% 0.110% 13 Roseobacter 0.06% 0.13% 2 0.01% 0.120% 14 Anaerococcus** 0.06% 1.01% 18 Planctomyces 0.01% 0.100% 19 Rubritalea 0.03% 0.13% 4 Sulfurimonas 0.01% 0.010% 3 Arcobacter 0.03% 0.29% 10 Faecalibacterium 0.004% 0.020% 5 Prevotella 0.03% 0.33% 12 Fusibacter 0.003% 0.010% 3 Desulfococcus 0.03% 0.39% 15 Bacteroides 0.002% 0.020% 10 Veillonella 0.03% 0.10% 4 Ruminococcus 0.0001% 0.001% 9 Halochromatium 0.02% 0.05% 2 *Weighted relative abundance (total abundance / # of OTUs). ** Increased significantly during relay

Correlation analyses of the relationship between V. parahaemolyticus concentrations and the sequence count abundance of all identified genera in all oyster samples (n=33) were then used to determine if other genera decreased in abundance like V. parahaemolyticus or increased like other Vibrio spp. during relay. The relative abundance of 40 identifiable genera significantly correlated (positive or negative) (p<0.05) with V. parahaemolyticus concentrations in the oysters

(Fig. 3). Thirty-two of the forty genera negatively correlated with V. parahaemolyticus concentrations while eight genera positively correlated with V. parahaemolyticus concentrations.

Vibrio, Corynebacterium, Peptoniphilus, Anaerococcus and Prevotella were the only genera identified that increased in abundance after 14-day relay and negatively correlated with V. parahaemolyticus concentrations (Table 2 + Fig 3). Vibrio spp. had the highest relative weighted

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abundance (1.4%) and lowest Spearman ρ coefficient (-0.556) of these 5 genera suggesting once again that Vibrio species are a potentially influential part of the oyster community composition that may affect V. parahaemolyticus concentrations in oysters.

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Fig. 3. The relationship between V. parahaemolyticus concentrations and genera abundance in relayed oyster samples. Spearman ρ (+) = positive correlation, Spearman ρ (-) = negative correlation. (-1) represents the most negative correlation between V. parahaemolyticus concentrations and the abundance of the respective genera. (+1) represents the most positive correlation between V. parahaemolyticus concentrations and the abundance of the respective genera. (0) represents no relationship.

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Almost all (95.5%) of the Vibrio 16S rRNA gene sequences clustered into one large

OTU. The majority of the sequences in this OTU could only be classified to the genus level, but of the 18,802 sequences that met the species classification confidence level (>=60% posterior probability) none were identified as V. parahaemolyticus. This is not surprising given that the sequence read length was short (<200 base pairs) making it difficult to distinguish between individual Vibrio species (particularly closely related species). As well, the detected V. parahaemolyticus concentrations ranged only from 3 to 4115 MPN/g oyster tissue (Table 1), a small fraction of the total microbial population in oysters. A higher abundance of identified

Vibrio sequences in relayed (Day 14) oyster samples suggest that a variety of Vibrio species increased in abundance during relay, and given the observed reduction in V. parahaemolyticus, reflect different responses for Vibrio species to relaying under the conditions of this study.

The taxa composition of relayed oysters does not mimic the composition of the respective harvest (Piscataqua River) and relay (Spinney Creek) waters. One probable mechanism by which the oyster microbiome changes during relay is by ongoing filtration of different taxa from the relay site water and simultaneous removal of some established taxa in the oyster. The oyster and water microbial communities were compared using Principal Coordinate Analysis (PCoA) to see if taxa were different or similar in the two environments. The oyster and water taxa clustered separately from one another (Fig. 4A) and were significantly different, (MRPP; p<0.001, A =

0.1374) (Table 3), confirming that oysters and water microbial communities are different.

Separation of the taxa in the water and oyster samples was clearly distinguishable along the PC2 axis where 27.5% of the sample variation is explained (Fig. 4A). There were a few oyster samples outside of the main cluster including a small tight cluster of oyster samples along the

PC1 axis, which shows a larger portion (33.3%) of the sample variation. Oyster and water

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samples plotted on the same coordinates but colored by sampling year (Fig. 4B) shows the taxa composition of oysters from 2011 and 2012 when relay was successful are similar and differ from taxa in 2013 oysters (MRPP; p=0.004, A = 0.1124), when relay was less successful (Fig.

4B + Table 3). The tight cluster of taxa in water samples, derived from the Piscataqua River and

Spinney Creek sites are distinctly dissimilar from the oyster samples (Fig. 4A) but have a similar taxa composition in 2012 and 2013 (Fig. 4B).

Fig. 4. Principal coordinates analysis of all oyster and water samples based on pair-wise sample Bray-Curtis dissimilarity measures. The ordination is colored by (A) sample type and (B) year to highlight sample similarity patterns related to these variables. PC 1 and PC 2 explain, respectively, 33.3% and 27.5% of the total data variance.

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Table 3. Non-parametric Multi Response Permutation Procedure (MRPP) analysis of Taxa Differences within Sample Matrices and Years – 2011-2013 Relay Experiments.

The difference in community composition (OTUs as determined by Bray-Curtis distances) in the oyster and water sample communities was also analyzed separately. The OTUs in both the oyster and the water samples were significantly different across sampling years. The effect size (MRPP A-value), which measures the magnitude of the differences of the communities between years was four times larger in oyster samples (p<0.001, A=0.1236) compared to water samples (p<0.001, A=0.0336), suggesting higher homogeneity within oyster microbial communities compared with water microbial communities. Additionally, the bacterial communities in all oyster samples from 2011 and 2012 were significantly different (p=0.004,

A=0.1124) compared to those collected in 2013. From 2011-2013 there were no significant differences in Day 0 oyster communities compared to the combined communities in Day 1 through 14 samples during relay. Significant taxa shifts appear to take place in the oyster and

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water matrices between years, but the variation in taxa shifts that occurred during relay were not significant.

The two study sites had consistent differences in conditions, particularly for salinity with the range of salinity being higher at Spinney Creek (25-31 ppt) than at the Piscataqua River (10-

25 ppt). We hypothesize this salinity difference promotes a change in resident microbiota.

Although the taxa in water from the two sites appeared to be similar based on the PCoA plot and were not statistically different (Fig. 4A + Table 3), some specific taxa were found to be significantly different in abundance between the two sites (Fig. 5A). The evidence of specific taxa differences at the two sampling sites supports the notion that some fraction of water microbiota may influence microbial shifts in oysters that may play a part in the effectiveness of relay.

Shifts in oyster community taxa composition were consistent when relay was successful and were different when relay was unsuccessful. Comparing community taxa in oysters where V. parahaemolyticus concentrations were effectively decreased during relay to taxa in oysters where V. parahaemolyticus concentrations did not decrease could provide further information about which taxa may influence oyster relay success. The relative abundance of taxa present in

2011-12 oysters, when relay was consistently effective, was markedly different from 2013 oysters (Fig. 5B + 5C). The more abundant taxa for Day 0 and 14 when relaying was successful in 2011 and 2012 (which included triplicate samples for July 2012) showed highly distinct and consistent patterns between Day 0 and 14 when plotted on a heat map (Fig 5B). This finding showed consistency in both the taxa shifts and in the taxa involved in successful relay experiments, even between different years. The 2011 and 2012 Day 0 and 14 oyster samples have two distinct groups of taxa which reflect the observed decrease in concentrations of V.

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parahaemolyticus in relayed oysters. V. parahaemolyticus concentrations were low for Day 0 and 14 samples and thus did not change during relaying in September 2011, and the respective taxa were not separated from each other, clustering with the taxa for the other 2011/12 Day 14 samples (Fig. 5B). The low concentrations of V. parahaemolyticus may have been due to the taxa already present at Day 0 in the oyster. Overall, relaying that is successful in reducing V. parahaemolyticus concentrations in oysters appears to include consistent changes in a low percentage of the total oyster community taxa, supporting the concept that there are consistent community level interactions with specific taxa that are involved in reducing V. parahaemolyticus concentrations. This did not hold true when relaying was unsuccessful.

We suspected that in 2013 when relaying was not successful in reducing V. parahaemolyticus concentrations that we would observe differences in oyster sample taxa shifts compared to 2011-12. Community analysis of OTUs for all oyster samples from 2011, 2012 and

2013 sampling seasons (Fig 5C) showed Day 0 oyster samples from 2013 had dissimilar taxa compared to those from Day 0 taxa for 2011 and June 2012. These 2013 Day 0 taxa, however, were similar to Day 0 taxa for August and September 2012, suggesting a shift in the microbial community had occurred between June and August 2012, even though the Day 14 taxa in August and September 2012 were consistent with those observed for other 2011-12 Day 14 samples. The main difference between the significant taxa from 2013 Day 14 samples and those for 2011-12 was that the 2013 Day 14 taxa clustered either with their paired Day 0 taxa (indicating no shift during relay) or with a unique group of taxa differing from the Day 14 taxa in 2011-12. Thus, the different changes observed in relayed oyster samples from 2013 compared to 2011-12 samples reflect the observed difference in relay success.

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The clustering of Brochothrix, Carnobacterium and in the 2013 oyster samples (Fig. 5C) is not evident in the 2011 and 2012 heat map (Fig. 5B) but the taxa were abundant to varying degrees in 2011 and 2012 oysters when all three years were analyzed (Fig

5C). These three genera were present at high abundance levels in oysters during 2013 relay based on the bright yellow / white color intensity in the heat map (5C –along x axis). Additionally,

Carnobacterium and Brochothrix positively correlated with the increase in V. parahaemolyticus in relayed oysters (Fig. 3). The presence of these taxa in the heat map when all three years were analyzed supports the interpretation that taxa have different interactions in the oyster microbiome and may variably influence V. parahaemolyticus concentrations during relay (74, 133, 134).

Results from the 2011-13 sample seasons show that relay-induced changes in the oyster microbiomes are associated with different community structures that appear more similar to each other in 2011 and 2012 when relaying was successful and differ from the community structure in

2013 when relaying was not successful.

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Fig. 5. The relative abundance and relatedness between OTUs associated with fresh and relayed oyster and water samples during 2011-13. The heat maps show taxa associations among statistically different samples. Yellow color intensity indicates sequence relative abundance (see key). The dendrograms show hierarchical clustering of OTUs (left) and samples (top). The hierarchal clustering highlights the distribution patterns of the OTUs that differed significantly between Day 0 and 14 samples. Day 14 samples are colored red in the dendrogram and sample labels. 5a. 2012 PR and SC water samples. 5b. All 2011 and 2012 oyster samples. 5c. All 2011, 2012 and 2013 oyster samples.

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Piscataqua River oysters and water have ten core community taxa that are consistently found in Spinney Creek water and relayed oysters. A survey of taxa composition oyster and water samples revealed a group of core taxa in all and within year water and oyster samples (Fig. 6). There were ten taxa that were present in all water and oyster samples, namely, Corynebacterium, Bacillus, Sediminibacterium, Serratia, Pseudomonas,

Lactococcus, , Vibrio, Staphylococcus and Lactobacillus. These ten genera represented a large proportion of the taxa found in all samples for each of the three study years, and almost all (10/12) of the taxa found in all Day 14 samples, suggesting the presence of a small but consistent core microbiome from year to year.

The ten taxa identified as core taxa were consistently among the ten taxa with the highest relative abundance in all oyster, all water and all oyster and water samples from each year (Table

4). Genera within the oyster microbiome were more numerous but less diverse than the water microbiome, possibly reflecting a host (oyster) effect for colonizing bacteria (145, 155). The highest number of taxa was identified in 2012 samples, with slightly fewer in 2013 and about half as many in 2011, whereas the diversity was highest in 2013 and lowest in 2011, which probably reflect the inter-annual changes in oyster microbiomes. Vibrios were included in the top ten identified taxa in all water, 2012 and 2013 samples but not in all oyster and 2011 samples.

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Fig. 6. Venn diagram of the the number of identified taxa found in 100% of the oyster and water samples by year and Day 14 of relay. Venn Diagram - Oliveros, J.C. (2007-2015) Venny. An interactive tool for comparing lists with Venn's diagrams. http://bioinfogp.cnb.csic.es/tools/venny/index.html

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Table 4. The diversity and relative abundance of taxa in oysters, water and both water and oysters in each study year (2011, 2012 and 2013). Oyster Diversity Water Diversity Richness 889 Richness 733 *Evenness 0.320 *Evenness 0.475 Genus Relative Genus Relative Rank #Genus Abundance Rank #Genus Abundance 1 Pseudomonas 0.2948 1 Pseudomonas 0.1071 2 Bacillus 0.2650 2 Streptococcus 0.0887 3 Sediminibacterium 0.0721 3 Bacillus 0.0617 4 Serratia 0.0498 4 Acinetobacter 0.0602 5 f__Brachyspiraceae 0.0295 5 Corynebacterium 0.0568 6 Mycoplasma 0.0246 6 f__Staphylococcaceae 0.0547 7 f__Bacillaceae 0.0187 7 Vibrio 0.0543 8 Lactococcus 0.0142 8 Serratia 0.0363 9 c__Mollicutes 0.0125 9 o__Streptophyta 0.0318 10 k__Bacteria 0.0115 10 Anaerococcus 0.0277

2011 Diversity 2012 Diversity 2013 Diversity Richness 434 Richness 872 Richness 780 *Evenness 0.273 *Evenness 0.416 *Evenness 0.436 Genus Relative Genus Relative Genus Relative Rank #Genus Abundance Rank #Genus Abundance Rank #Genus Abundance 1 Pseudomonas 0.4757 1 Pseudomonas 0.2299 1 Bacillus 0.2253 2 Bacillus 0.2366 2 Bacillus 0.0752 2 Pseudomonas 0.1133 3 Serratia 0.0903 3 Sediminibacterium 0.0748 3 Streptococcus 0.0570 4 Sediminibacterium 0.0496 4 Serratia 0.0536 4 Staphylococcus 0.0386 5 f__Bacillaceae 0.0186 5 Streptococcus 0.0502 5 Acinetobacter 0.0365 6 f__Brachyspiraceae 0.0157 6 Acinetobacter 0.0415 6 Vibrio 0.0358 7 Lactococcus 0.0129 7 Corynebacterium 0.0373 7 Corynebacterium 0.0319 8 k__Bacteria 0.0105 8 Vibrio 0.0345 8 o__Streptophyta 0.0267 9 c__Mollicutes 0.0094 9 Staphylococcus 0.0316 9 Anaerococcus 0.0246 10 g__Mycoplasma 0.0085 10 f__Leuconostocaceae 0.0299 10 Serratia 0.0244

*Evenness is on a scale of 0 to 1. 1 = an equal proportion of all genera present. 0 = one genus completely dominant in the group and all other genera at a very low abundance. #Classifications not at the genus level are noted with a letter in front of an underscore to represent the classification level (k= , c= class, o= order, f= family, g= genus).

Identification of cultivable microbiota in relayed oysters. Taxa identified by 16S rRNA gene sequencing were a guide for targeting potentially influential cultivable taxa that are in 2014 relayed oyster samples where concentrations of V. parahaemolyticus were successfully reduced.

We did this to design enrichment and cultivation methodologies so that isolates from successful relay trials could be utilized to further examine the relative competitiveness in culture with V.

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parahaemolyticus. Thirty-eight isolates differing by colony characteristics on Marine agar were classified into two phyla. Ninety-four percent of the sequences from the isolates classified to phylum , while the remaining 6% belonged to the . Sixty-four percent of the isolates were classified to Vibrio at the genus level, including one each that classified as V. harveyi, V. aestuarianus, and V. shilonii. Vibrio species were expected to respond well to Marine agar, and the prevalence of Vibrio species in the 2014 relayed oysters is also in accordance with earlier described results and other studies (155). Genus Pseudomonas comprised 17% of the isolates, including some identified as P. pseudoalcaligenes and P. umsongensis. The isolates belonging to the Firmicutes classified to families Bacillaceae and Exiguobacteraceae.

Many of the previously described results suggest that non-V. parahaemolyticus Vibrio species may influence V. parahaemolyticus reductions during oyster relay, and simple competition assays were used to investigate this. A variant V. parahaemolyticus strain with an intergenic transposon insertion generating an erythromycin-resistance that did not differ in specific growth rate and final biomass when compared to the wild type V. parahaemolyticus was used for assessment of competitiveness with Vibrio alginolyticus and Vibrio fluvialis (Table 5) under the culture conditions. After two days (48 h) of co-culture, the erythromycin-resistant V. parahaemolyticus strain did not out-compete either V. alginolyticus or V. fluvialis, based on the relative realized growth rates (W values) for the competition cultures compared to pure cultures.

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Table 5. Vibrio spp. used in the competition assay with environmental strain G3654 of Vibrio parahaemolyticus from Spinney Creek. Source Genus Species Competitive Characteristic Reference (ATCC #)

Community member in oyster A clinical isolate from Romalde Vibrio fluvialis microbiome, no known Maine oysters - 2016 et. al, 2014 biocontrol agent properties

Community member in oyster Romalde A clinical isolate from a microbiome et. al, 2014 Vibrio alginolyticus Maine - 2016 Verschuere Biocontrol agent properties et. al, 2000

DISCUSSION

Recent studies have reported on the composition and dynamics of the oyster microbiome

(62, 64, 72, 145, 156). Vibrio spp. found naturally in estuarine waters can persist at concentrated levels in shellfish matrices (157) so we wanted to investigate oyster microbiome dynamics in relation to V. parahaemolyticus concentrations during pre-harvest relay treatment. Continued exposure of the oysters to new conditions (e.g. higher salinity) resulted in reductions in V. parahaemolyticus levels, and, in turn consistent and significant changes to the oyster microbiome. This utilizes 16S rRNA gene sequencing to help understand how oyster relay works to eliminate from live oysters. We provide evidence for consistent shifts in relayed oyster community taxa following a significant relay-associated change in environmental conditions, i.e., from low salinity water to water with elevated salinity, that reflect reductions in

V. parahaemolyticus concentrations. One apparent explanation for these results is that changes in

V. parahaemolyticus concentrations during relay were influenced by the changing microbial

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community in relayed oysters. The shifts in taxa during relay were consistent when relay was successful in reducing V. parahaemolyticus levels, although the taxa shifts were both variable and different when the relaying was not successful. Further research on taxa shifts during successful relaying are needed to gain a better understanding about the environmental conditions and potential biological mechanisms that may cause these changes to occur.

Multiple factors, including the different niches found within shellfish, interactions between microbial communities associated with different estuarine ecosystem matrices, and environment conditions, can influence the microbiota that reside within shellfish (56, 66). Many factors can influence the density of different Vibrio species in the oyster microbial community

(125, 141, 145, 158, 159), including filtration from the estuarine water column followed by either bio-accumulation and persistence within the oyster microbiome, host defense related elimination, or depuration back into the water column (74, 147). Prolonged exposure of oysters to changes in environmental conditions (120), including significant changes in salinity, has also been shown to influence the colonized microbiota in oysters (154, 160), and is another significant factor influencing the displacement of Vibrios from oysters (114). Additionally, physical disturbance and other stresses on host shellfish can cause profound changes in their microbiomes (145). The process of handling and harvesting oysters can disturb the host and cause changes in the microbiome resulting in increased Vibrio levels that can last for varying time (161). Translocation of oysters by relay is a disturbance to the host that can initially increase Vibrio levels (131, 160), suggesting other possible changes in the oyster microbiome.

Whether successful relaying of oysters to remove V. parahaemolyticus is a function of bio-accumulated taxa in the oyster from water or the change in environmental conditions that affect the oyster and its microbiome is not something that can be easily distinguished. In this

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study, the oyster microbiome taxa differences between Day 0 and Day 14 samples were not the same as the observed differences between the water microbiomes at the harvest and relay sites.

This suggests that the mechanism for reducing V. parahaemolyticus in relayed oysters is probably a complex function of environmental conditions, host response to relaying and interactions between the relay water microbiome and the oyster microbiome.

In 2013, after nearly two years of near-record high sea surface temperatures (SST) in the

Northwest Atlantic and the Great Bay estuary (8, 98) relaying was coincidently unsuccessful for reducing V. parahaemolyticus concentrations in oysters. The elevated SST in the GBE during

2011-13 can be considered a period of disturbance to the Great Bay estuary ecosystem, including oysters, as regional species shifts and invasions were reported during this time period (162, 163).

Environmental conditions can induce changes in whole ecosystem biomes that can have profound effects on Vibrio levels (13, 96, 164). Some of the taxa that were distinct in 2013 are possibly related to ecosystem changes caused by the higher regional SST. Changes in oyster microbiomes were coincident with increases in SST in the Great Bay estuary and were apparent with a shift in taxa profiles for Piscataqua River oysters starting in August 2012 and continuing into 2013. The abundance of Brochothrix, which has been implicated in shrimp spoilage (165,

166), Carnobacterium that can be a fish pathogen or probiotic in aquaculture and expresses chitinase like V. parahaemolyticus (167) and Bacillus, arguably the most studied potential probiotic (86, 87, 90, 168, 169) were significantly (p < 0.05) elevated in 2013 oyster samples.

Regional changes in water temperatures are thus a likely important factor that can cause changes in oyster microbiomes and influence the success of relaying to reduce V. parahaemolyticus concentrations. Thus, climate-induced ecosystem changes, like significant increases in water temperature and extreme rainfall/runoff with associated reductions in estuarine salinity, can have

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significant public health implications (14, 163, 170). The increased incidence of vibriosis and oyster-borne outbreaks of V. parahaemolyticus illnesses in the region over this time period (19,

92) is evidence of this disturbance and points to Vibrio spp. as a useful indicator of climate impacts on human illness occurrence (171).

Inter-species competition between V. parahaemolyticus and other species is one potential explanation for why relaying can be successful. Closely related Vibrio spp. appeared to be potential competitors of V. parahaemolyticus in relaying, based on consistently observed inverse shifts in abundance, even though some limited and preliminary co-culture experiments did not bear this out. Similar genetic capacity in Vibrio spp. for persistence in different ecological conditions is a factor that may enhance the competitiveness of other Vibrio spp. with V. parahaemolyticus under certain conditions. Vibrio spp. include pathogenic and probiotic species that can be numerically dominant in oysters and seawater (172, 173). V. parahaemolyticus can also be displaced by other bacteria that are better suited to environmental conditions because they have antagonistic and anti-bacterial capabilities (41, 174). The potential for another Vibrio species to out-compete V. parahaemolyticus is consistent with the negative correlation between

16S-identified Vibrio species and V. parahaemolyticus MPN counts. As the microbiome shifts in composition in disturbed oysters, it is likely that other Vibrio spp. and species better suited to the new higher salinity environment displace V. parahaemolyticus populations.

In this study we analyzed oyster samples directly from relay experiments to generate initial findings on potentially important microbiological factors that underlie how successful relay works. Future research can lead to identifying what bacterial community taxa could be used to influence the reduction of naturally occurring V. parahaemolyticus in relayed oysters to help prevent human illness.

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MATERIALS AND METHODS

Oyster and water sample collection. Oyster and water samples collected during relay experiments (131) were used for community member analyses. Briefly, oysters (Crassostrea virginica) were harvested by diving or tonging from an oyster bed in the upper Piscataqua River

(43°10’08.49”/70°49’42.54”) in Dover, New Hampshire during June to September from 2011-

2014 and relayed to Spinney Creek in Eliot, Maine. Historically the Piscataqua River has lower salinity levels and higher V. parahaemolyticus concentrations in comparison to Spinney Creek.

Enough oysters were harvested and transported to allow for analysis of V. parahaemolyticus levels in three separate sub-samples of 12 animals (per homogenate) at pre-determined sample times. Water samples (500 mL) were also taken at the two sites using sterile Nalgene (Rochester,

NY) bottles. Oysters were suspended in the water column of Spinney Creek within a basket attached to a mooring ~1 m from the water surface. On the respective sampling days, (Day 0 and

Day 14) oysters were removed from the basket and transported to the lab in an insulated cooler with ice packs for immediate analysis.

Sample processing and determination of V. parahaemolyticus concentrations in oyster and water samples. V. parahaemolyticus concentrations in oyster homogenates and water samples were calculated using the FDA Bacteriological Analytical Manual three-tube MPN method in combination with real time PCR. APW tubes were scored for turbidity, a 1.0 mL aliquot of each turbid APW tube and water sample was transferred into a sterile 1.5 mL centrifuge tube, pelleted at 8000 RPM (6093 x g) in a Sorvall Legend Micro 17 tabletop micro-centrifuge with a Fiberlite

Micro 24x2 rotor (Thermo Scientific, Waltham, MA) for 5 min and boiled for 10 mins at 100oC.

The qPCR was performed in 25 µL reactions containing OmniMix Mastermix (Cepheid,

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Sunnyvale, CA), 75 nM tlh primers, 150 nM tlh TaqMan probe, MgCl2 (Bio-Rad Laboratories,

Hercules, CA) final concentration 5 mM, 1µL of an internal amplification control (IAC)

(BioGX, Birmingham, AL), 75 nM IAC primers, 150 nM IAC probe and 4 ul of template DNA.

The qPCR parameters in the Cephied Smart Cycler (Cephied, Sunnyvale, CA) included a hot start step at 95°C for 3 minutes followed by 45 cycles of amplification with a denaturation step at

95°C for 5 sec and annealing/extension at 59°C for 45 sec. The primers, fluorescence readings, dye set, and manual threshold units followed Nordstrom et al. (139).

Sample DNA extraction for 16S rRNA gene sequence analysis. Total bacterial DNA was extracted from frozen water and oyster samples and fresh oyster samples using an E.Z.N.A. Soil

DNA Kit (Omega Bio- Tek, Norcross, GA) following the manufacturer’s protocol. Quality and integrity of extracted DNA was visualized following electrophoresis on a 1.5% agarose gel stained with gel red. Oyster meat samples from each month of the three sampling seasons (2011-

2013) and the respective water samples were selected in order to allow for 16S rRNA gene sequencing for microbial community trend analysis during the 14-day relay experiments. A representative set of samples were chosen to analyze 16S rRNA gene sequences in water and oyster samples at day 0 and day 14 of relay during the months of the sampling seasons from

2011-2013. Oyster homogenate and water samples were preserved for sequencing experiments by freezing sample material in 50 mL polypropylene conical tubes (Corning, Corning, NY) at -

80oC. In 2014, Sanger sequencing analysis was performed on amplicons from bacterial isolates cultivated on Marine agar from fresh oyster samples during the sampling season as an additional analysis to determine trends in cultivable taxa during relay.

16S rRNA gene sequence amplification, library construction, and Illumina sequencing. A total of 68 water and oyster meat samples from 2011-2013 were utilized to generate 16S rRNA

65

gene amplicon libraries. Extracted DNA was amplified in triplicate from water and oyster samples to normalize and decrease amplification bias in the library before Illumina sequencing.

The V4 hypervariable region of the 16S rRNA gene was targeted using forward and reverse primer sequences along with thermal cycler parameters listed in Caporaso et al. (152).

Amplicons generated from PCR were visualized on 1.5% agarose. PCR reagents were removed from the amplicon libraries using a MoBio (Carlsbad, CA) ULtraclean PCR Clean-up kit. Pooled amplicon libraries were quantified using a Qubit Fluorometer (Invitrogen, Carlsbad, CA).

Libraries from 2011-2013 oyster and water samples were sequenced using the Illumina (San

Diego, CA) NGS HiSeq 2500 sequencer following published protocols (152, 175). Paired-end

150 base sequences were generated for each library and all samples were loaded into a single lane on the Illumina flow . Fastq files were generated using CASAVA software version 1.8.3 from Illumina to de-multiplex the samples based on the 12 base pair Golay barcodes. De- multiplexing statistics were generated using Illumina software.

Sequence analysis. 16S rRNA gene sequence libraries were processed using an AXIOME II pipeline (176). This automated pipeline makes use of a variety of marker gene analysis tools and ensures reproducibility of analyses. The paired-end sequences were assembled with PANDAseq

(177) with a quality threshold of 0.9. UPARSE was used for clustering of the marker genes at

97% sequence identity (178) with a de novo chimera check. The most frequently observed sequence in an OTU was chosen as its representative sequence. Taxonomic classifications were generated by the Ribosomal Database Project (RDP) classifier, version 2.2, via the QIIME 1.8.0 package (179). RDP was trained against the GreenGenes 13_8 revision reference set, with the default posterior probability of 60% used for the classification cutoff. The 60% cutoff was used to determine the taxonomic classification (consensus lineage) down to the lowest level

66

confidence criterion. Operational taxonomic unit (OTU) table creation was performed by

QIIME. The OTU table was randomly subsampled at ten different sequencing depths (evenly spaced between 10 and 182210 sequences per sample) ten times for each depth. The alpha diversity statistics were calculated for each table, and the mean and standard deviation of these statistics were plotted.

Statistical analysis. Sequence data were normalized to adjust for disproportionate sequence count bias by dividing the sequence counts for each OTU by the total number of sequences in the sample. The percent relative abundance raw data for relay samples were organized by consensus lineage. The abundance results for each lineage were then sorted and categorized based on whether they had a majority or minority of OTUs increase in sequence relative abundance during relay. A sum total of the abundance data for each of the OTUs that increased was calculated and weighted against the amount of OTUs identified for each lineage. This initial analysis was a general assessment to understand which genera increased during relay experiments.

Additionally, percent abundance data was used for discovery odds ratio testing to determine the OTU abundance counts that were significantly different between sample groups.

We assessed which OTUs differed significantly during relay in oyster and water samples by the discovery odds ratio test in the R package, metagenomeSeq version 1.10.0. Normalized data were log-scaled and resulting p-values adjusted by false discovery rate (FDR) correction.

Significantly different OTUs (p<0.05) were visualized in a heat map with the hierarchical clustering by complete linkage of the samples and OTUs plotted on the axes. The point of mapping only the significantly different OTUs on heat maps was to determine if there is a distinction between and/or consistent genera in the harvest and relay water samples and Day 0 and 14 relayed oyster samples from 2011 to 2013.

67

Oyster V. parahaemolyticus concentrations, as determined by MPN/qPCR, were compared to the relative abundance of each identified genus in all oyster samples with a

Spearman correlation (non-parametric rank correlation) to examine which genera significantly correlated with V. parahaemolyticus concentrations in relayed oysters. Significance of the correlation was assessed by F-test, and p-values (< 0.05) were adjusted with FDR for multiple test correction. A linear model was used to analyze the relationship between the log MPN V. parahaemolyticus concentrations and the log relative abundance of Vibrio sequence reads for all samples.

The vegan package in R was used to create Bray-Curtis principal coordinates analysis

(PCoA) ordinations for all samples. In this ordination, each point represents the microbial community of a sample, and the distance between points in the two-dimensional plot approximates pairwise distances between samples. Non-parametric Multi Response Permutation

Procedure (MRPP) analysis on Bray-Curtis dissimilarities was used to assess the dissimilarity of oyster and water samples by quantifying abundance-weighted differences between the microbial communities from each sample. MRPP was used to confirm, statistically, whether certain sample groupings contained distinct microbial communities. The MRPP A value ranges from 0 to 1, where A=0 implies that the sample dissimilarities can be explained by chance alone rather than the given sample grouping, and A=1 implies that all samples within a given group are identical and distinct from other groups. Shannon Diversity was calculated for each sample in a rarefied (evenly subsampled) OTU table to understand the richness and evenness of taxa diversity in the oyster and water samples in relay experiments from 2011-2013.

Competition assay. The fitness of Vibrio species relative to V. parahaemolyticus was determined using methods based on related studies (38, 86). Pure cultures of the competitor

68

species (Table 4) and V. parahaemolyticus were incubated in Marine Luria Bertani (MLB) broth

(per liter: 10 g tryptone, 5 g yeast extract, Instant Ocean to provide 25 ppt salinity; pH = 7.0) in a

Tecan Infinite M200 plate counter (Männedorf, Switzerland) at 25°C for 24 h time periods.

Optical density readings at 600 nm were used to determine lag and exponential phase duration, specific growth rates (µ) average rates of biomass increase (m), or the Malthusian parameter, in

24 hours for each species:

mx = (lnXd/lnXo)/d

Where mx is the Malthusian parameter for species x, Xd=CFU reading after 1 day (d=1; 24h) &

Xo=CFU reading at T=0.

For competition experiments, we used V. parahaemolyticus strain G3654 isolated from

Spinney Creek oysters in 2013, a Vibrio alginolyticus clinical strain from a 2016 wound infection, and a Vibrio fluvialis clinical strain isolated from Maine oysters in 2016 (Table 4).

These species were chosen based on them being regional isolates from the Gulf of Maine and on preliminary results from an initial competition experiments between V. parahaemolyticus and 7 other Vibrio species, with V. alginolyticus being a potential competitor and V. fluvialis being a species that V. parahaemolyticus can out-compete. The V. parahaemolyticus strain G3654 was first modified to be resistant to erythromycin.

The ERM mutant and wild type V. parahaemolyticus strains were cultured in the plate counter to determine any negative fitness effects on the mutant. The Vibrio species were grown together in 200 µl MLB broth at 25°C in 1.0 ml wells of the plate counter. All cultures were pre- conditioned for one 24-h period in the same media and plate counter, and then transferred to new

69

plates for 2 consecutive 24-h incubations using 1:100 diluted cultures at the beginning of each

24-h incubation. Viable cell counts were determined in triplicate using Marine agar ± erythromycin for total culture counts and to select for the constructed ERM-resistant V. parahaemolyticus strain. The viable cell counts for the competitors were determined by subtracting the V. parahaemolyticus strain counts from the total plate counts. We spread 10 µl of decimally diluted cultures onto the agar media using the rounded bottom end of small sterile test tubes and incubated the plates overnight at room temperature (~22°C). Triplicate incubations of pure cultures for each competitor were run on the same plate counter, along with triplicate blank wells containing un-inoculated media. The natural logarithms of each competitor after 3 days of competition will be used to determine relative realized growth rates (W), which is the ratio of m values between the competitor and V. parahaemolyticus under the competition conditions:

Wab = ma / mb

for competitors A and B. The w value for competition conditions was compared to the w value calculated based on parallel pure, non-competitive cultures of V. parahaemolyticus and the two competitors, to determine which species is more competitive.

Nucleotide sequence accession numbers. All sequence data referred to in this article have been deposited in the European Nucleotide Archive (ENA) database

(http://www.ebi.ac.uk/ena) under the accession number PRJEB20315.

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APPENDICES

88

APPENDIX A

CHAPTER II SUPPLEMENTAL MATERIAL Fig. A.1 Table A.1

89

Fig A.1. Lab procedure for processing oysters after field collection. Oyster homogenates consisted of 12 live oysters.

90

Table A.1. 2011-2014 Oyster and Water Sample MPN Metadata

Year Month Day MPN Matrix Source 2011 7 0 330 Oyster PR 2011 7 0 720 Oyster PR 2011 7 0 330 Oyster PR 2011 7 2 279 Oyster PR 2011 7 2 27.9 Oyster PR 2011 7 2 183 Oyster PR 2011 7 7 330 Oyster PR 2011 7 7 72 Oyster PR 2011 7 7 330 Oyster PR 2011 7 14 9 Oyster PR 2011 7 14 9 Oyster PR 2011 7 14 37 Oyster PR 2011 8 0 330 Oyster PR 2011 8 0 28 Oyster PR 2011 8 0 138 Oyster PR 2011 8 2 330 Oyster PR 2011 8 2 330 Oyster PR 2011 8 2 330 Oyster PR 2011 8 7 225 Oyster PR 2011 8 7 128 Oyster PR 2011 8 7 23 Oyster PR 2011 8 14 45 Oyster PR 2011 8 14 63 Oyster PR 2011 8 14 28 Oyster PR 2011 9 0 72 Oyster PR 2011 9 0 12.9 Oyster PR 2011 9 0 63 Oyster PR 2011 9 7 330 Oyster PR 2011 9 7 0.28 Oyster PR 2011 9 7 1380 Oyster PR 2011 9 14 36 Oyster PR 2011 9 14 330 Oyster PR 2011 9 14 6.3 Oyster PR 2011 11 0 2.79 Oyster PR 2012 6 0 36 Oyster PR 2012 6 0 200 Oyster PR 2012 6 0 150 Oyster PR

91

2012 6 7 7.4 Oyster PR 2012 6 7 3 Oyster PR 2012 6 7 3 Oyster PR 2012 6 10 3 Oyster PR 2012 6 10 3 Oyster PR 2012 6 10 3 Oyster PR 2012 6 14 6 Oyster PR 2012 6 14 11 Oyster PR 2012 6 14 9.2 Oyster PR 2012 7 0 2400 Oyster PR 2012 7 0 2400 Oyster PR 2012 7 0 12100 Oyster PR 2012 7 7 3.6 Oyster PR 2012 7 7 3.6 Oyster PR 2012 7 7 16 Oyster PR 2012 7 10 3 Oyster PR 2012 7 10 3 Oyster PR 2012 7 10 2100 Oyster PR 2012 7 14 9.4 Oyster PR 2012 7 14 6.2 Oyster PR 2012 7 14 1100 Oyster PR 2012 8 0 6.2 Oyster PR 2012 8 0 200 Oyster PR 2012 8 0 4600 Oyster PR 2012 8 7 6.2 Oyster PR 2012 8 7 3 Oyster PR 2012 8 7 3 Oyster PR 2012 8 10 3 Oyster PR 2012 8 10 9.4 Oyster PR 2012 8 10 29 Oyster PR 2012 8 14 3 Oyster PR 2012 8 14 3 Oyster PR 2012 8 14 3 Oyster PR 2012 9 0 38 Oyster PR 2012 9 0 21 Oyster PR 2012 9 0 3 Oyster PR 2012 9 7 93 Oyster PR 2012 9 7 6.1 Oyster PR 2012 9 7 43 Oyster PR 2012 9 10 11 Oyster PR

92

2012 9 10 3 Oyster PR 2012 9 10 9.2 Oyster PR 2012 9 14 3 Oyster PR 2012 9 14 3 Oyster PR 2012 9 14 3 Oyster PR 2012 10 0 3 Oyster PR 2013 6 0 150 Oyster PR 2013 6 0 1500 Oyster PR 2013 6 2 110000 Oyster PR 2013 6 2 46000 Oyster PR 2013 6 7 430 Oyster PR 2013 6 7 460 Oyster PR 2013 6 10 24000 Oyster PR 2013 6 10 46000 Oyster PR 2013 6 14 3600 Oyster PR 2013 6 14 38000 Oyster PR 2013 7 0 1200 Oyster PR 2013 7 0 430 Oyster PR 2013 7 2 460000 Oyster PR 2013 7 2 46000 Oyster PR 2013 7 2 110000 Oyster PR 2013 7 7 11000 Oyster PR 2013 7 7 150000 Oyster PR 2013 7 10 15000 Oyster PR 2013 7 10 1100000 Oyster PR 2013 7 10 16000 Oyster PR 2013 7 14 3600 Oyster PR 2013 7 14 15000 Oyster PR 2013 9 0 9300 Oyster PR 2013 9 7 430 Oyster PR 2013 9 14 290 Oyster PR 2013 10 0 930 Oyster PR 2013 10 7 94 Oyster PR 2013 10 14 62 Oyster PR 2013 11 0 3 Oyster PR 2014 6 0 30 Oyster PR 2014 6 0 30 Oyster PR 2014 6 14 36 Oyster PR 2014 6 14 290 Oyster PR 2014 7 0 930 Oyster PR

93

2014 7 0 150 Oyster PR 2014 7 14 1100 Oyster PR 2014 7 14 1100 Oyster PR 2014 8 0 110000 Oyster PR 2014 8 0 930 Oyster PR 2014 8 14 110 Oyster PR 2014 8 14 3.8 Oyster PR 2012 6 0 3 Water PR 2012 7 0 240 Water PR 2012 8 0 23 Water PR 2012 9 0 9.2 Water PR 2013 6 0 15 Water PR 2013 7 0 93 Water PR 2013 9 0 920 Water PR 2013 10 0 11 Water PR 2014 6 0 3 Water PR 2014 7 0 93 Water PR 2014 8 0 93 Water PR 2012 7 14 9.2 Water PR 2012 8 14 9.2 Water PR 2012 9 14 3 Water PR 2011 7 0 3 Water SCS 2011 7 2 11 Water SCS 2011 7 7 4.6 Water SCS 2011 8 0 4.6 Water SCS 2011 8 2 3 Water SCS 2011 8 7 11 Water SCS 2011 9 0 3 Water SCS 2011 9 7 3.8 Water SCS 2012 6 0 3 Water SCS 2012 6 7 3 Water SCS 2012 6 10 3 Water SCS 2012 7 0 93 Water SCS 2012 7 7 7.4 Water SCS 2012 7 10 240 Water SCS 2012 8 0 6 Water SCS 2012 8 7 6 Water SCS 2012 8 10 93 Water SCS 2012 9 0 3 Water SCS 2012 9 7 3 Water SCS

94

2012 9 10 3 Water SCS 2013 6 2 2300 Water SCS 2013 6 7 110 Water SCS 2013 6 10 1100 Water SCS 2013 7 0 1200 Water SCS 2013 7 2 460 Water SCS 2013 7 7 7500 Water SCS 2013 7 10 230 Water SCS 2013 9 0 6.1 Water SCS 2013 9 0 7.4 Water SCS 2013 9 7 3 Water SCS 2013 9 7 11 Water SCS 2013 10 0 29 Water SCS 2013 10 7 11 Water SCS 2014 6 0 3 Water SCS 2014 6 14 9.2 Water SCS 2014 7 0 24 Water SCS 2014 7 14 2.3 Water SCS 2014 8 0 21 Water SCS 2014 8 14 3 Water SCS 2011 7 14 11 Water SCS 2011 8 14 3 Water SCS 2011 9 14 20 Water SCS 2012 6 14 3 Water SCS 2012 7 14 21 Water SCS 2012 8 14 3 Water SCS 2012 9 14 3 Water SCS 2013 6 14 43 Water SCS 2013 7 14 930 Water SCS 2013 7 14 150 Water SCS 2013 9 14 21 Water SCS 2013 10 14 93 Water SCS

95

APPENDIX B

CHAPTER III SUPPLEMENTAL MATERIAL Tables B1-B2

96

Table B.1. Oyster and water sample V. parahaemolyticus MPN/g concentrations and 16S count metadata for samples submitted for 16S rRNA gene sequencing

97

Table B.2. CASAVA Quality Data for Oyster and Water Samples

98